AMER. ZOOL., 14:821-824 (1974).
A Method of Raising Clones of the Hydroid Phialidium gregarium
(A. Agassiz, 1862) in the Laboratory
STEVEN G.
WORTHMAN
Department of Biological Structure, University of Washington,
School of Medicine, Seattle, Washington 98195
SYNOPSIS. The hydroid Phialidium gregarium can be successfully cultured to sexual
maturity at any time of the year and maintained for years in a closed system marine
aquarium. Artificial sea water was used in place of natural sea water and was renewed by one-quarter of its volume each 4 weeks, Freshly hatched Artemia nauplii
were used as the sole food source. Methods for obtaining colonies of a single genotype
from planulae are described. Subciilturing pieces from parental stock is accomplished
by a new method. A new technique for removing hydroid colonies from their substrate for histological study is described.
INTRODUCTION
We have found a closed system aquarium
provides adequate culture conditions for
growth and sexual reproduction of Phialidium gregarium hydroids kept for periods
of up to 3 years. Such maintenance provides the investigator with a rare opportunity to study developmental processes o£
clones of animals the year round under
controlled laboratory conditions. The
methods used to establish colonies of
Phialidium and their subsequent maintenance are the concern of this paper.
MATERIALS AND METHODS
Colonies of Phialidium gregarium were
cultured in a 190-liter marine aquarium
(Dayno Aqua Lab. Model 203, Dayno
Sales Co., Lynn, Mass.). Water circulation through a calcite filter in the base
of the aquarium was 13 liters/min which
This research was supported by USPHS Grant
GM-00136 from the National Institutes of Health
and by Grant GB-8094 from the National Science
Foundation.
I would like to thank Dr. E. C. Roosen-Runge
for his unfailing guidance and help developing the
culture system described, especially with reference
to the capillary tube method of subculturing. Also,
I would like to express thanks to Dr. R. L. Fernald,
Director of the Friday Harbor Laboratories, for
allowing me the use of that facility.
821
helped maintain pH. near 8.2. Oxygenation was aided by two jets of water which
perturb the surface layer. The operating
temperature range was 10.5 C to 13.5 C.
Colonies did not tolerate temperatures
above 15 C for long without showing signs
of degeneration.
Synthetic sea salt and trace elements
(Dayno Sales Co., Lynn, Mass.) were dissolved in stainless steel-distilled water and
the salinity maintained between 30—33 ppt
by periodic addition of distilled water. Onequarter of the total volume of the aquarium
was changed each 4 weeks and synthetic
sea water made 3 days earlier was added
at each change. Such water renewals helped
sustain a pH range of 7.6 to 8.2 and kept
ammonia levels below 0.02 ppm. The
metabolic balance of the aquarium, water
was easy to maintain because of the small
biomass in culture compared with the relatively large water volume in circulation.
The glassware used was embryologically
clean, having had no contact with detergents or toxic chemicals. Microscopic slides
used as substrates were soaked overnight
and rinsed in five changes of culture water.
Hydroid colonies were initiated from
planulae collected at the Friday Harbor
Laboratories of the University of Washington according to the method of RoosenRunge (1970). Planulae were allowed to
settle on glass slides spread over the bottom of 2-liter Pyrex rectangular dishes.
822
STEVEN G. WORTHMAN
Water containing day-old planula larvae
was gently poured into the dishes. The containers were then loosely covered with
polyethylene sandwich wrap to inhibit
evaporation, and immersed almost to their
tops in the aquarium by being placed on
plexiglass shelves hung 4 cm beneath the
water surface. The dishes remained undisturbed for 4 days, at which time 60 to 80%
of the planulae had settled. Slides having
multiple settlings were carefully thinned
so that the most healthy appearing stolon
was ultimately the only genotype populating that slide. After examination, each
slide was marked for identification using a
diamond stylus and placed into a 300-ml
Pyrex bowl which was later covered with
a square plexiglass sheet. The bowls were
kept in contact with aquarium water on
a shelf and remained undisturbed for 3
days or until functional hydranths made
their appearance. Then feeding was begun
with a Pasteur pipet under a dissecting
scope. Watchmaker's forceps were used to
feed bits of dissected Artcmia to young
feeding hydranths too small to ingest whole
brine shrimp (Werner, 1968). In a few
days when primary hydranths could hold
two or three Arlemia, feeding was regularized by the following procedure. Five
slides were fitted into a plexiglass slide
top
'
front
TI«W
'
end
FIG. 2. Diagram of the plastic feeding container, 500 ml volume. All measurements are in
centimeters.
holder (Fig. 1) and immersed into a 500-ml
bath of Artemia (Fig. 2) for 6 min. Care
was taken to ensure that the bath
equilibrated with aquarium temperature
(12.5 C) and was free of debris that would
foul the colonies. After feeding, the cultures, still in the slide holder, were hung
from two plexiglass rails running the
width of the aquarium. In this position,
the slides lay parallel to the drift of the
current, long axis vertical, with the top
frame of the culture holder 10 cm beneath
the water surface.
Method of subculluring new colonies
I-J.7H
I" I
When a culture had expanded over both
sides of a slide (3 to 4 months), it was
used as parental stock for subculturing
clones by vegetative transplantation. A
stainless steel dissecting needle was used
176
to cut 0.5-mm pieces of stolon and attached
hydranths out of the colony. The older
stolons were the most easily removed and
damaged least by the process.
To fasten the excised pieces, we invented
HI
a new method employing a capillary tube
that proved to be more efficient than tying
front vi«w
and
pieces to the slide. The excess perisarc was
FIG. 1. Diagram of the plastic slide holder used
trimmed away and the diameter of the
to carry five slides standing vertically in the
stolon measured with an eyepiece micromgrooves of the frame. All measurements are in
centimeters. Slides need not be removed only at
eter using a stereo dissecting scope. A
the ends of the frame because the depth of the capillary tube 1.5 mm long was selected
grooves is such that the slides are mobile and can
and measured for an inside diameter
be removed by lifting them out of the bottom
groove regardless of their location along the frame. slightly larger than the perisarc of the cutU
I
I
LABORATORY CULTURE OF
ting to be transplanted. A small quantity
of silicone grease (Dow Corning Stopcock
Grease) was spread on a dry slide. The
slide was then placed into a 300-ml Pyrex
bowl full of culture water, and the selected
capillary was submerged and pressed into
the grease with its long axis parallel to the
slide surface. The cutting was transferred
with a large bore eyedropper to the transplant bowl and its stolon maneuvered into
the free end of the capillary. For proper
positioning, the stolon must hang over
the edge of the tube and touch the slide
with its hydranth remaining upright. The
stolon attaches within 3 days if the transplant remains undisturbed. Feeding the
single hydranth was begun immediately
with a Pasteur pipet. Later when two or
more hydranths appeared, colonies were
fed by the submersion method previously
described.
Phialidium
HYDROID
823
in absolute ethanol before the standard
paraffin embedding procedure was begun.
Use of a clear plastic substrate has other
advantages if one wishes to keep a photographic record of colonial growth. Reference lines are easily scratched into plastic
and measurements made in terms of such
a reference coordinate system lend themselves well to computer analysis of growth
data.
REMARKS
Our primary objective in raising colonies of Phialidium is to establish healthy
parental stock of singular genotype that
can be used to study developmental processes in the laboratory. The maintenance
of constant culture conditions and the
establishment of a regular feeding schedule is as important for the proliferation
of Phialidium gregarium (Roosen-Runge,
Removing colonies from the substrate for 1970) as it is for Cordylophora (Fulton,
histological purposes
1962, 1963) and Campanularia (Crowell,
1957). Although some investigators feed
When it was desirable to remove pieces hydroids twice a day, maintenance of
of a colony intact for histological prepara- Phialidium colonies was obtained by one
tion, a plastic substrate was used. Plexiglass feeding per day. Colonial growth is slowed
sheets measuring 5.1 cm X 7.6 cm X u-16 and significant hydranth regression occurs
cm were dipped once into a 2% parlodion- when colonies are fed less than once a day.
butylacetate solution (Mallinckrodt Chemi- Hydroids vary in their potential to thrive
cal Work, New York). This parlodion in the aquarium. One reason, for example,
overlay adheres tightly to plexiglass and is that a hydroid which cannot regulate
cannot be removed by abrasion as it can the size of its hydranths to accommodate
from glass. After drying, the substrate was Artemia may not feed even if food is
detoxified by soaking it in three changes present. Algae and diatoms encrust such
of culture water over an 8-hr period. When slow-growing hydroids, interfering with
the colonies grown on the substrate were their development.
to be removed, they were fixed and dehyThriving colonies undergo a period of
drated in a graded series of alcohols up apparent dormancy following a period of
to absolute alcohol. Cultures were then sexual reproduction and growth, but inplaced into a 15-cm petri dish containing creased feeding can occasionally interrupt
absolute ethanol and 0.45 M butylacetate such a trend. It is often necessary to scrape
(mol wt 116.16). Within about 5 min the one-half of a slide clean on one side with
parlodion film loosened and growing tips a razor blade in order to renew growth
of the stolon could be selectively lifted from the old stolon mat. This method is
from the slide with a dissecting needle and like transplanting the colony to a new subexcised with iiidectomy scissors. More rapid strate and regrowth proceeds in a similar
removal could be effected b) increasing fashion.
concentrations of the solvent, butylacetate.
The traditional method of making transCuttings were cleaned of debris and washed plants by tying cuttings to a slide with
824
STEVEN G. WORTHMAN
thread was not used for Phialidium because employing the capillary tube technique allows smaller pieces of a colony
to be used for regenerating a stolon. Regrowth seemed to be more rapid if small
cuttings were transplanted. Also, unlike
the capillary technique, tying with thread
did not ensure that single hydranths would
remain upright in feeding position.
Removing a colony or piece of stolon
from its substrate has been a problem for
investigators, especially when it is to be
used for histological study. Methods of
removal differ depending on the nature of
the substrate used for growth. Scraping can
be employed if the colony grows over a
dense algal mat (Hale, 1960). If no such
mat is available and the colony grows on
glass, the wax technique may be used to
remove stolons from slides (Braverman,
1971). The first method was judged inconvenient for our purposes and the second
method, though practical, does not remove
all of the stolon of Phialidium. When a
particular part of a stolon must be recovered, the plastic slide coated with a
parlodion film is quite satisfactory.
When hydroids are raised in the laboratory, the process narrows the range of
genotypes gathered from the wild. For
example, some planulae do not settle on
slides in the allotted amount of time.
Thinning those that do settle is necessary
to make room for stolon growth, and again,
not all of the colonies thrive under the
given culture conditions. Selection of hydroids for growth in a closed system is,
therefore, only an approximation to the
selection conditions that actually exist in
the wild.
REFERENCES
Braverman. M. 1971. Studies on hydroid differentiation. VII. The hydrozoan stolon. J. Morphol.
135:131-152.
Crowell, S. 1957. Differential responses of growth
zones to nutritive level, age, and temperature in
the colonial hydroid Campanularia. J. Exp. Zool.
134:63-90.
Fulton, C. 1962. Environmental factors influencing
the growth of Cordylophora. J. Exp. Zool. 151:
61-78.
Fulton, C. 1963. The development of a hydroid
colony. Develop. Biol. 6:333-369.
Hale, L. J. 1960. Contractility and hydroplasmic
movements in the hydroid Clytia johnstoni.
Quart. J. Microscop. Sd. 101:339-350.
Roosen-Runge, E. C. 1970. Life cycle of the hydromedusa Phialidium gregarium (A. Agassiz, 1862)
in the laboratory. Biol. Bull. 139:203-221.
Werner, B. 1968. Polypengeneration und Entwicklungsgeschichte von Eucheilota maculata (Thecata-Leptomedusae). Mit einem Beitrag zur
Methodik der Kultur mariner Hydroiden. Helgolander Wiss. Meercsunters. 18:136-168.