IMONFISH PARASITES
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CULTURE OF SOME COMMON FISH 
PARASITES
FOR EXk'ERIMENTAL STUDIES
Heino Beckert
Zoology-Entomology Department 
Series
Fisheries No. 5
AGRICULTURAL EXPERIMENT STATION
AUBURN UNIVERSITY
E. V. Smith, 
Director
AuunAabm
May 1967

CULTURE OF SOME COMMON FISH PARASITES
FOR EXPERIMENTAL STUDIES*
Heino Beckert
Instructor
I. Introduction
Fish parasite cultures in the laboratory constitute an im-
portant aspect of research on control of these organisms. Basic
information about the organism is a prerequisite to effective
parasite control. Necessary information includes host range, life
cycle, and special environmental conditions either advantageous or
detrimental to the parasite, such as the organism's response to
temperature, oxygen content, pH, and salinity of the water. To
obtain information of this type the parasite must be available in
sufficient numbers to permit study and experimentation.
A number of parasitic organisms have been cultured in the
laboratory. Characteristically, internal parasites were cultured
in vitro, usually on tissue cultures. Uzmann and Hayduk (1963)
successfully cultured Hexamita salmonis, an intestinal flagellate
of young trout and salmon. Williams, Hopkins, and Wyllie (1961)
cultured the strigeid trematode Diplostomum phoxini in vitro.
Jensen, Stirewalt, and Walters (1965) cultured cercariae of Schistosoma
Supported by the Southeastern Cooperative Fish Parasite and
Disease Project.
mansoni in vitro for as long as 112 days. Meerowitch (1965) raised
Trichinella spiralis from larvae to immature adults in vitro.
Hoffman (1958) reared metacercariae of Posthodiplostomum minimum
to infertile adults. Hoffman and Putz (1964) maintained a population
of Gyrodactylus macrochiri on 6 month old bluegills in running
water in the laboratory. Hlond (1966) reported an in vitro culture
of Ichthyophthirius multifiliis on a carp slime medium. However,
personal communication with Mgr. Hlond, Instytut Zootechniki, Zator
k. Oswiecimia, Poland, revealed that only the tomite state of I.
multifiliis was reared and the parasite did not complete its life
cycle in vitro. Tidd (1963) cultured Lernaea cyprinacea in the
laboratory and infected goldfish as well as tadpoles and young
adults of Rana pipiens with this parasite. Lahav and Sarig (1964)
cultured L. cyprinacea in the laboratory, describing in detail 
the
life cycle of this organism. Dr. Hoffman, Eastern Fish Disease
Laboratory, Leetown, W. Va. describes in a personal communication
a culture method for both Ichthyop thirius and L. cyprinacea.
These parasites were maintained by dividing an aquarium with a
screen and placing infected fish in 
one end and uninfected, non-
immune fish in the other end. When the initially infected fish
died they were replaced by more nonimmune hosts.
Most fish obtained from a natural habitat show some 
degree of
parasitism. However, the fact that the incidence of parasitism is
usually low in wild fish unless an epizootic is occurring, 
and the
fact that wild fish are usually parasitized by more than one species
of parasites make it difficult to obtain specific 
information about
a single type of parasite. Therefore,it is necessary to obtain pure
cultures of specific parasites under laboratory conditions to elimi-
nate variable environmental conditions that occur in a natural
situation.
The following conditions should be met for culturing a fish
parasite in the laboratory:
(1) An adequate supply of host fish is important. It is
most desirable to use fish from the same population, 
that is indi-
viduals from the same parent stock, or at least fish spawned at the
same time and in the same pond.
(2) Prospective host fish must be removed to an isolated en-
vironment as soon as possible to prevent natural infections 
by para-
sites and disease organisms. It is best to store young host fish
in small ponds to facilitate their removal by seining. These storage
ponds must be large enough to permit carrying out an adequate
feeding program. Ponds with a surface area of 0.05 acre 
and an aver-
age depth of 2.5 feet have been used with good results at Auburn.
Where facilities are available, host fish can be stored 
in troughs
with running water. If an adequate flow of water can be maintained,
a relatively large number of fish can be kept and fed in such a
facility. The chief advantage of using troughs is that the fish can
be observed and treated against parasites and diseases prior 
to
experimental infection. This is not possible in ponds 
because
close and regular observation of the fish population is not feasible.
Where possible fish can be spawned in the laboratory and the fry
reared to fingerling size in steel troughs with running water. This
ensures parasite-free experimental fish that are not immune to para-
sites.
4
(3) It is essential that host fish used for maintaining a
parasite culture be healthy and if at all possible free of disease
and parasites. Periodic examinations of the fish should be made.
(4) The parasite culture is best maintained in a volume of
water of manageable size. r'orty-liter aquaria and 100-liter feeding
troughs have been successfully used at Auburn.
(5) Since the life cycles of many parasites are influenced
by temperature, it is desirable to maintain the parasite culture in
an environment of relatively constant temperature.
(6) Dechlorinated water with a pH value of 7 or 8 is best
suited for maintaining healthy host fish.
II. Laboratory Culture of Ichthyophthirius multifiliis
The organism. Ichthyophthirius is a one-celled, ciliated
parasite of fishes that affects skin and gills, causing a ccndition
known as white spot disease, or ichthyophthiriasis (Fig. 1 and 2).
This organism is one of the most cosmopolitan and destructive
parasites of fresh water fishes. Observations on the life cycle
of I. multifiliis agreed with those given by Kudo (1960).
At maturity the parasite, lodged just under the host's
epithelium, leaves the host and swims free in the water (Fig. 3).
A short time after leaving the host, the parasite attaches itself
to aquatic vegetation, stones, or other suitable substrate and
encysts. The cytoplasm divides into as many as 1,000 small round
daughter cells (Fig. 4 to 7). The daughter organisms or tomites
(Fig. 8 and 9) break through the cyst wall, become elongate and
swim actively about in search of a host. Once in contact with a
host, the tomites bore into the skin where they parasitize the fish.
Depending on the temperature the parasite reaches maturity in a
5
period of time ranging from a few days to several weeks.
Host fish. A number of host species for culturing Ichthyophthyrius
have been used at Auburn. Carp, goldfish, bluegills, tilapia,
fathead minnows, golden shiners, channel catfish and white catfish
were used. Scaly fish were found to be unsatisfactory because in
the course of a heavy infection with the parasite these fish secrete
large amounts of mucus and lose scales; this interferes with the
recovery of the mature parasites.
Exposure of host fish to the parasite. Ten channel catfish
or white catfish fingerlings each were stocked into 40-liter
(10 gallons) aquaria filled with dechlorinated tap water. Then, a
single heavily infected catfish fingerling was killed and placed
into each aquarium. The water temperature was kept between 70
and 80aF. Four or 5 days later, small white spots could be seen on
the live fish. The resulting infection was usually heavy enough to
maintain the culture and to allow experimentation with the parasite.
It was not possible, however, to produce mild, uniform infections
consistently. Only when host fish were exposed to large numbers of
the parasite did uniform but heavy infections result. A minimum of
200 trophozoites per 10 fish in 40 liters of water should be used.
Recovery of mature parasites. The parasites should not be
collected until they have almost reached maturity, that is until they
are about to leave their host to% encyst. When premature parasites
are obtained by scraping them from the fish, the resulting number
of daughter organisms is very small, yielding only a mild infection
upon subsequent exposure to other fish. On the other hand, the para-
sites should be collected before any of them leave the fish since
6
Fig. 1. Channel catfish fingerling infected with I.
multifiliis. The white spots covering the
fish are trophozoites lodged 
just under the
host's epithelium.
Fig. 2. Gill filaments of a channel catfish finger-
ling parasitized by I. multifiliis.
Fig. 3. Mature trophozoite of I. multifiliis showing
the characteristic horse-shoe shaped
macronucleus.
Fig. 4. Trophozoite of I. multifiliis after the first
division. The two daughter cells are encased
by a clear cyst wall.
Fig. 1
Fig. 2
Fig. 3 
FgFig. 4
8
Fig. 5. Trophozoite of I. multifiliis after the second
division.
Fig. 6. Cyst of I. multifiliis.
Fig. 7. Cyst of I. multifiliis showing the cyst wall and
containing almost mature tomites.
Fig. 8. Tomite of I. multifiliis just after leaving the
cyst.
Fig. 9. Tomite of I. multifiliis showing the strong
ciliation of the orgonism.
Fig. 7
Fig. 5
Fig. 8
Fig. 6 FgFig. 9
10
any parasite swimming in the water or being encysted on the aquarium
bottom is for all practical purposes lost for use in the culture.
Since all of the host fish are exposed to the parasite at the
same time and since most of the parasites are in the same stage of
development when introduced to the host fish, most of the parasites
are equally far along in their development when collected.
The most efficient method of recovering and collecting mature
Ichthyophthirius is pithing a heavily infected host fish and placing
it in a petri dish filled with dechlorinated tap water. Shortly
after the host's death, the parasites leave the fish and swim about
in the water.
To avoid a low oxygen situation by leaving the dead fish
in the container with the parasites that already have left the host,
it is desirable to remove the pithed fish after 
about 1 hour and
place it into another dish. After several consecutive changes
about 1 hour apart, a large number of healthy and normal trophozoites
of the parasite can be collected in the dishes.
Counting of trophozoites. In culturing Ichthyophthirius in
the laboratory, it is often desirable to count a specific number of
parasites for further use in infecting other fish. The counting
of trophozoites is greatly facilitated 
by placing the dish con-
taining the parasites on a dark background. Mature trophozoites
are usually 0.5 to 1.0 am. in diameter and of a whitish or gray
color. Against a dark background the trophozoites can easily be
counted with the unaided eye. However, a large-diameter magnifying
glass and strong illumination are helpful and less fatiguing. The
best time for counting trophozoites is immediately after the parasites
11
have begun to settle to the bottom of the dish. At that time the
parasites adhere to the glass instead of actively swimming through
the water and are easily collected.
If only a small number of trophozoites is required, individual
parasites can be sucked up with a finely drawn-out pipette and
placed in a staining dish. The parasites are then poured into
another aquarium to infect a new host population.
Where larger numbers of trophozoites are needed, it is desirable
to draw with a grease pencil eight wedge-shaped sections on the
bottom of the dish containing the pithed fish. If, for instance,
300 trophozoites are needed, this number can be counted and the
sections containing the desired parasites marked with a grease
pencil. Since the parasites are adhering to the glass, they can
be washed off into an aquarium with the aid of a wash bottle. This
washing off must be done gently to prevent injury to the parasites.
Care should be taken in avoiding great temperature differences
between the water in dishes containing the parasites and the water
in aquaria containing the fish to be infected.
III. Laboratory Culture of Cleidodiscus pricei
The organism. Cleidodiscus pricei (Fig. 10) is a common gill
parasite of channel catfish. It is a monogenetic trematode, namely
this gill fluke does not have an intermediate host and exhibits no
polyembryony like digenetic trematodes such as the liver fluke. After
copulation the adult lays adhesive, three-lobed eggs (Fig. 11 and 12)
that cling to the mucus on the gills or drop off the fish into the
water.
12
Fig. 10. Adult of C. pricei attached to a gill filament
of a channel catfish. The posterior end of the
worm is armed with a set of hooks with which
the parasite fastends itself to the host's gill
epithelium. The anterior end of the worm has
four eye-spots.
Fig. 11.
Fig. 12.
Fig. 13.
Fig. 14.
Fig. 15.
Adult C. pricei with fully formed egg just before
it was laid.
Egg of C. pricei with its characteristic spine.
When the larva is ready to emerge the operculum
at one of the corners breaks open.
Larva of C. pricei. The four eye-spots and the
hooks can be distinguished.
Larva of C. pricei showing a tuft of cilia at the
posterior end and a lateral band of cilia around
the midsection. The band of cilia at the anterior
end is not visible.
Gill filaments of a channel catfish infected
with C. pricei.
Fig. 10
Fig. 11
Fig. 13
Fig. 12
Fig. 14Fg.1
/
Fig. 15
14
Upon hatching, 
the ciliated 
larva (Fig. 13 and 
14), looking
much like a miracidium 
of a digenetic 
fluke, swims rapidly 
about
in search of a host 
fish. After the 
larval form has reached 
the
gills, the cilia are lost; it attaches 
itself to a gill filament
and assumes the 
appearance of 
a. miniature adult 
worm (Fig. 15).
Exposure of host fish 
to the parasite. Rarely 
can a gill fluke-
free channel catfish 
be found. Use can 
be made of this fact 
by
placing 10 or 
15 channel catfish 
fingerlings in 
a 
4
0-liter aquarium.
By not changing 
the water, the 
parasite population 
is allowed to
build up. After several 
weeks there is 
usually a heavy infection
present on most 
fish. These heavily 
infected fish 
can be placed
into aquaria with 
lightly infected 
fish and the culture 
may thus
be maintained.
The disadvantage 
of this culture method 
is that, while the 
fish
are being held in 
aquaria, feeding 
is not possible. Consequently,
all fish are in 
a very poor physical 
condition. Weaker 
individuals
may die and the parasite 
culture may be lost 
as the host population
density decreases.
When channel catfish 
are being held in aquaria, 
even without
feeding, a certain 
amount of debris accumulates 
on the bottom. If
the fish have a gill 
fluke infection, many 
of the eggs can be 
found
in that debris. 
This material 
can be syphoned 
into a container,
preferably a urine sedimentation 
glass, to concentrate the debris.
The debris should 
be checked microscopically 
for gill fluke eggs,
and introduced into 
aquaria containing 
channel catfish to 
maintain
the culture.
At Auburn, debris 
containing fluke 
eggs was collected 
and
15
aliquots of the material introduced into several aquaria. The
culture was maintained in this manner for several months.
When gills infected with adult worms are removed from a channel
catfish, placed in a petri dish and covered with water, the parasites
are in a stress situation. It was observed that within 30 minutes
most adult worms had laid one egg each. The eggs can easily be re-
moved from the gill tissue with a fine pipette. Most of the eggs
collected in this manner yielded infective larvae. By this method
a relatively large number of eggs can be obtained for experimentally
infecting new host populations.
Use of feeding troughs with running water. Since the disad-
vantage of culturing gill flukes in aquaria is the resulting
poor condition of the fish, an experimental system was used at
Auburn that allowed light feeding of the fish while being exposed
to the parasite.
At first, a 100-liter stainless steel feeding trough was stocked
with 50 channel catfish fingerlings. A very small amount of water
was passed through the trough and the fish were fed approximately
1 per cent of their body weight each day. The fish were examined
for gill flukes twice weekly for 4 weeks. No significant in-
crease in the gill fluke population was found. It was assumed
that most of the parasite eggs and larvae were washed from the
trough.
To determine if better results in culturing gill flukes
could be obtained by impeding the flow of water through feeding
troughs, the following procedure was used: One feeding trough
was used as control; the water entered from the faucet and went
16
b.
Fig. 16. a. Regular feeding trough, b. feeding trough
with baffles.
out through a standpipe. In the other trough 
a series of Styro-
foam baffles was used to impede the water flow and prevent loss
of eggs and larvae by the running water (Fig. 16). Each trough
was stocked with 50 channel catfish fingerlings that were fed a
maintenance diet of 1 per cent of their body weight each day. The
water temperature was 550F. Prior to stocking, 10 fish from the
experimental population were checked for infection with C. pricei.
Periodic counts of gill flukes were made from five fish from each
trough and the number of parasites found was recorded. All gill
17
fluke counts were made froma the outer right gill (Fig. 
17 and 18).
On the average, the fish harbored 6.3 gill flukes per outer
gill prior to stocking. The parasite population on the fish stocked
into the feeding trough without baffles decreased during the 65-day
experimental period. It can be assumed that the unimpeded water
flow carried away a sizeable portion of parasite eggs and larvae.
On the other hand, the parasite population increased on the fish
held in the feeding trough with baffles. It appears that with a
decrease in water flow Cleidodiscus pricei increases in number on
the host. An experimental set-up similar to the one described
would be well suited for culturing C. pricei in the laboratory
in large numbers.
Collecting gill flukes. The best method to collect and count
gill flukes for quantitative determinations is to remove one or
several gill arches, rinse them gently with a wash bottle and place
them into a vial half filled with water. The vial should then be
placed in a freezer to remain for 24 hours. Freezing relaxes the
flukes and causes them to loosen their hold on the gill filaments.
After being frozen for 24 hours, the vial content is thawed and
shaken briskly, using an agitator with vortex action. This
shaking removes most of the mucus clinging to the flukes and makes
the parasites clearly visible (Mizelle, 1936). The contents of the
vial is then poured into a small dish, preferably one with a grid
bottom and placed on a dark background. Under a low power dissect-
ing microscope, the gill flukes can readily be seen and counted.
18
Mean
**-*. 
Range
12
L 11
0 9
0 8
0.
' 7
-, 6
0 3
L4
S0 1 2 3 4 5 6 7
Sample No.
Fig. 17. Average number of Cleidodiscus pricei on the gills
of channel catfish fingerlings held in a 100-liter feeding trough
without baffles for 65 days. Samples were taken at approximately
9-day intervals.
19
Mean
Range
18
17
m 16
En 15
L 14
i 13
0
L 12
S11
cL 10
9
8
7
6
L, 5
.0 4
E
: 3
C
2
< 1
0 1 2 
3 4 5 
6 7
Sample No.
Fig. 18. Average number of Cleidodiscus pricei on the gills
of channel catfish fingerlings held in a 100-liter feeding 
trough
with baffles for 65 days. Samples were taken at approximately 
9-
day intervals.
20
IV. Laboratory Culture of Lernaea cyprinacea
The organism. Lernaea cyprinacea is a parasitic 
copepod in-
fecting primarily cyprinid and catastomid hosts, but may parasitize
many other freshwater fishes.
This parasite is commonly known as anchor worm because its
anterior end if modified into an anchor-like structure (Fig. 19 a)
that enables the parasite to lodge itself firmly under the skin
of its host.
Studies of the life cycle of this parasite (First Annual Report,
Cooperative Fish Parasite and Disease Project, July 1, 1964 to
June 30, 1965; Agricultural Experiment Station, Auburn University)
show that mature eggs develop into parasitic forms after 
having
undergone as much as seven molts, going through 
the naupliar and
metanaupliar stages (Fig. 19 b and c). The 
first parasitic form,
namely the first copepodid stage (Fig. 19 d) 
infects fish. Five
more molts occur. Mating takes place during the 
last copepodid
stage. The male does not develop further. The 
female burrows
into the skin where it becomes embedded. The 
characteristic anchors
keep the parasite affixed to its host. Under good 
conditions one
female will produce several pairs of egg 
sacs. A temperature of
approximately 800F is optimal for this parasite.
Host fish. Since the anchor worm parasitizes 
readily all
cyprinids, goldfish or carp lend themselves readily 
for use as
host fish. These species are hardy and can 
be kept in aquaria for
a considerable period of time when fed lightly.
Collection of parasite eggs. As the adult female 
develops,
the worm's posterior portion can be seen 
protruding from the host's
b.
Anchor
a.
d.
Egg sacs
Fig. 19. Stages in the life cycle of Lernaea cyprinacea.
a. adult, b. nauplius, c. metanauplis, d. copepodid stage.
(After Putz and Bowen, 1964).
Adult anchor worm protruding from the abdomen
of a goldfish. (Photograph by Dr. H. S. Swingle)
Almost mature egg sacs of an anchor worm.
(Photograph by Dr. H. 
S. Swingle)
Naupliar stage of L. cyprinacea. (Photograph by
Dr. R. Allison)
Shed skin of one of the naupliar stages.
(Photograph by Dr. R. Allison)
22
Pig. 20.
Pig. 21.
Fig. 22.
Pig. 23.
4-,
~
I
...........
Fig. 22Fi.2
Fig. 20
Fig. 21
Fig. 23
24
skin (Fig. 20). A pair of egg sacs (Fig. 21) is formed at the
posterior end. At first these egg sacs are greenish, 
with maturity
they become whitish. At that point they should be removed from the
adult. This is best done by removing the host 
fish from the aquarium,
holding it under a strong light. Preferably the fish should 
be
removed from the aquarium with a wire basket rather than with a
net. This will reduce the chance of injury to the worms. The
fish should be held firmly in a paper towel. The egg sacs can
then be pinched off with a pair of sharp forceps. Care should be
taken that the adult worm is not injured or accidentally dislodged
in the process.
Incubation of the eggs. Once the egg sacs are recovered from
the parasite they should be placed in a small dish filled with
dechlorinated water. Water temperature should be 80*F. The dish
should then be placed in an incubator set at 80*F.
Most eggs will hatch within 24 hours and the still free-living
organisms will undergo a series of molts. The larvae (Fig. 22)
should be checked frequently. Shed skins (Fig. 23) are evidence
that molting is taking place. At 804F approximately 72 hours will
elapse before the first copepodid stage is reached. This stage is
characterized by on elongation of the abdomen, which differentiates
it from the last metanaupliar stage.
Exposure of host fish to the parasite. It is best to introduce
only the first copepodid stage to the host fish since earlier,
non-parasitic stages may be eaten by the fish, thus reducing the
chance for infection. Water containing the infective larvae should
25
be poured gently into the aquaria containing previously unexposed
fish. The water in the aquaria should be of the same temperature
as that in which the larvae were kept.
At Auburn it was not possible to obtain uniform, light infections
in the laboratory. However, fairly satisfactory 
results were ob-
tained by exposing 3, 6-inch goldfish in a 40-liter aquarium 
to
the first copepodid stages obtained by incubating one egg sac.
If a too light infection results, this method of infecting 
fish
could be varied by increasing the infection rate.
The fish in the aquaria should be fed lightly. Starving fish
will feed on the protruding anchor worms of other fish.
Symptoms of anchor worm infection. Two or 3 days after
introduction of the parasitic stages, the fish usually show lack
of mobility, folding of fins while staying close 
to the bottom of
the aquaria.
Later small bloody lesions appear on the base of fins and on
the sides of the body. The fish may show jerking 
movements and rub
themselves on the bottom or sides or the aquarium. Approximately
10 days after exposure to the first copepodid stage, small anchor
worms can be seen on the fish.
26
SUMMARY
Ichthyophthirius multifiliis was cultured in the laboratory by
the following method: Ten non-immune channel catfish or white
catfish fingerlings each were stocked into 40-liter aquaria filled
with dechlorinated tap water. Then, either a recently killed,
heavily infected catfish fingerling or at least 200 trophozoites
of the parasite were introduced into each aquarium. After the new
host fish became visibly infected, the parasites were recovered
by killing the host fish, placing one fish into a petri dish filled
with water. The trophozoites left the host almost immediately
after its death and swam in the water. They were collected with a fine
pipette at that time or later after they began to adhere to bottom
of the dish.
Cleidodiscus pricei was cultured in the laboratory by crowding
15 channel catfish fingerlings, which usually have a mild, natural
infection, into one aquarium. The parasite population then gradually
built up. Gill fluke eggs can be collected either by syphoning them
off the bottoms of aquaria containing infected fish or by removing
gills harboring the worm and collecting the eggs as they are being
laid. To avoid starving the host fish in aquaria, 100-liter
feeding troughs equipped with baffles to impede the water flow were
stocked with 50 channel catfish fingerlings, having a natural,
light parasite infestation. By feeding the fish approximately 1
percent of their body weight each day, the hosts remained healthy
and the parasite population was allowed to build up. The baffle
system prevented, to a large extent, the loss of parasite eggs and
larvae.
27
Lernaea cyprinacea was cultured in the laboratory 
on cyprinid
fishes. Mature egg sacs were removed from the adult 
parasite and
incubated at 80*F. for approximately 72 hours. At that time in-
fective larvae had developed. The 
first copepodid stage was then
introduced to non-immune host fish. Three 6-inch cyprinids in a
40-liter aquarium were exposed to the first copepodid larvae ob-
tained by incubating one egg sac. The water temperature was between
75 and 80
0
F.
Literature Cited
Hlond, S. Experiments In Vitro Culture of Ichthyophthirius
multifiliis. FAO World Symposium on Warm Water Pond Fish
Culture. FR:IX/E-4. Rome, Italy. 1966.
Hoffman, G. L. Experimental Studies on the Cercaria and Metacercaria of
a Strigeid Trematode, Posthodiplostomum minimum. Exp. Parasit.,
7:23-50. 1958.
Hoffman, G. L. and Putz, R. E. Studies on Gyrodactylus macrochiri
n. sp. (Trematoda:Monogenea) from Lepomis macrochirus. Proc.
Helm. Soc. Wash., 31:76-82. 1964.
Jensen, D. V., Stirewalt, M. A., and Walters, M. Growth of Schistosoma
mansoni Under Dialysis Membranes in Rose Multi-purpose Chambers.
Exp. Parasit., 17:15-23. 1965.
Kudo, R. R. Protozoology. 4th Edition. Charles C. Thomas, Publisher,
Bannerstone House. Springfield, Ill. 1960.
Lahav, M. and Sarig, S. Observations on the Biology of Lernaea
cyp~rnacea L. in Fish Ponds 
in Israel. Bamidgeh, 16:77-86. 
1964.
Meerovitch, E. Studies on the In Vitro Axenic Development of
Trichinella spiralis. 1. Basic Culture Technique, Pattern of
28
Development, and the Effects of 
the Gaseous Phase. Helm. Abst.,
34(2):1298. 1965.
Mizelle, J. D. New Species of Trematodes from the Gills of Illinois
Fishes. A. Midl. Nat., 17:785-806. 1936.
Putz, R. E. and Bowen, J. T. Parasites of Fresh Water Fishes. IV.
Miscellaneous. The Anchor Worm (Lernaea cyprinacea) and Related
Species. U.S. Dept. of Interior. Fish and Wildlife Service.
Fisheries Leaflet 575. 1964.
Tidd, W. A. Transfer of Larvae of Lernaea cyprinacea L. from
Goldfish to the Leopard Frog, Rana pipiens. J. Parasit.,
51(2-2) No. 157. 1965.
Uzman, J. R. and Hayduk, S. H. In Vitro Culture of the Flagellate
Protozoan Hexamita salmonis. Science, 140:290-292. 1963.
Williams, M. 0., Hopkins, C. A., and Wyllie, M. R. The In Vitro
Cultivation of Strigeid Trematodes. III. Yeast as a Medium
Constituent. Exp. Parasit., 11:121-127. 1961.