A method for securing the snail, Lymnaea stagnalis jugularis (Say), free from bacteria

A method for securing the snail, Lymnaea stagnalis jugularis (Say), free from bacteria

EXPERIMENTAL PARASITOLOGY 16, 7-13 (1964 A Method for Securing jugularis (Say), the Snail, Lymrzuea stagnaZis Free from Bacterial Frank The Unive...

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EXPERIMENTAL

PARASITOLOGY 16, 7-13

(1964

A Method for Securing jugularis (Say),

the Snail, Lymrzuea stagnaZis Free from Bacterial

Frank The University (Submitted

of

E. Fried1

South Florida,

for publication,

Tampa, Florida

20 February

1963)

Individual eggs obtained from isolated egg masses of the snail Lymnaea stagnalis jugularis (Say) were surface sterilized with the aid of a dilute solution of sodium hypochlorite. After subsequent washing in a favorable salt solution (designated SX), 90% of such treated eggs would hatch. The yield of living specimens that were obtained bacteria-free ranged up to 100%.

The in vitro propagation of snails, free from microbial contamination, is of considerable interest to the physiologist and parasitologist. Such experimental material can facilitate critical and much needed studies on snail metabolism and nutrition and enhance the development of tissue culture techniques; additionally, it can provide reliable systems with which one may study and evaluate selectively introduced parasites. It is with this latter intent that the present work has been undertaken in the course of investigations on the nature of the biochemical relationship between the snail, Lymnaea stagnalis jugularis (Say), and the trematode, Fascioloides magna. Chernin (1957) secured bacteriologically sterile Australorbis glabratus, and Chernin and Schork (1959) reported growth of this pulmonate snail on a diet of autoclaved yeast and formaldehyde-killed Escherichia coli. Michelson (1959) demonstrated a technique 1 This work was initiated at The Rockefeller Institute, New York, during the .tenure of a U.S. Public Health Service Research Fellowship from The National Institute of Allergy and Infectious Diseases. It was continued in Tampa with assistance from The University of South Florida and NSF Grant G19950.

whereby Viviparus japonicus could be obtained bacteriologically sterile. The method to be described, modified from that which Chernin (1957) employed for Australorbis, includes innovations specifically favoring the manipulation and study of L. stagnalis jugularis by the present author. As will be shown, this potentially large pulmonate snail can now be routinely obtained free from microbial contamination without the use of antibiotics. MATERIALS

AND

METHODS

Maintenance of Stock Snails The colony of L. stagnalis jugularis (Say) used in this work was originally obtained from Minnesota and was kept in the laboratory over 5 years. Stock snails (Fig. 1) as a rule were maintained in S- and lo-gallon aquaria with undergravel filters, constant aeration, and low-level illumination of one U-watt bulb per aquarium. A satisfactory diet included daily additions of lettuce, and commercial Pablum Mixed Cereal sprinkled sparsely over the surface of the water. Powdered calcium carbonate was added less frequently and distilled water was used to maintain a constant water volume. The snails were kept at about 2 1“C.

8

FRIEDL

FIG. 1. .4 pulmonate

Procuring

snail. L. slagnalis

jr~gzdaris (Say), on the bottom of one of the stock aquaria

Egg Masses

Egg masses, considered in good condition when firm and not grossly overgrown with microorganisms (Fig. 2), were carefully removed by means of a spatula and forceps from the sides of the stock tanks within a few days after they were produced. They were placed in stack finger bowls of IO-cm diameter containing 20-30 ml of a salt solution to be referred to as SX (Table I). After about 12 days at room temperature (usually 20”-26’C) the young snails as a rule appeared to be well developed within their egg membranes (Fig. 3). The hatching of a few individuals has been considered indicative of an egg mass ready to be used. Some egg masses taken from aquaria which contained much microorganismic growth were subjected to a “pretreatment.” These were immersed for 10 minutes at room temperature

in a solution containing Clorox (a commercial solution of 5.25:/( sodium hypochlorite by weight) diluted 11400 with SX. Customarily these “pretreated” masses were kept in 5 ml of SX solution in cotton stoppered, SO-ml Erlenmeyer flasks instead of finger bowls. This procedure was not employed when material could be obtained in good condition directly from the stock tanks. Disaggregation

of Individual

Eggs

A choice egg mass was placed in 10 ml of SX in a Petri dish and slit lengthwise with fine, alcohol-flamed scissors. It was then transferred to a 16 X 150 mm tube containing 10 ml of SX, capped by the thumb. and shaken vigorously until the eggs were freed of their surrounding matrix. This jellylike material with adherent bubbles generally floated to the top and could easily be

SECURING

Lymnaea

FIG. 2. An egg mass of the type used to supply eggs for obtaining

I TIC. 3. A 12-day-old egg of L. stagnalis jugularis rou ended by its egg membrane.

9

FREE FROM BACTERIA

showing

axenic L. stagnaZis jugularis.

a well-developed

but unhatched

snail su,r-

FRIEDL

10

Constitution _~~. NaCl

Milligrams

Salts

Distribution

TABLE I of SX Salt SolutionQ per liter

1000

KC1

50

NaaHPO,

25 100

MgSO,-7H,O CaCJ

a7

(Anhydrous)

NaHCOa Disodium

120 25

EDTAb

Trace elementsc

Micrograms

per liter 10

CuSO~.5H20

100

FeCI,

100

H3BO3

100

MnSO,.4H,O

10

ZnCl,

U Phenol red, 2.5 mg per liter, was added as a pH indicator. A precipitate, which formed after sterilization by autoclaving, was resuspended before use or dissolved by bubbling exhaled air through the solution. The solution eventually reached a pH of about 8.4 when exposed to air. b Disodium ethylenediaminetetraacetate. c Conveniently supplied from a 1000 x stock suspension.

removed. The eggs, suspended three times in IO-ml changes of SX, were then ready to be surface sterilized. Surface Sterilization The following steps were carried out using sterile technique. Disaggregated eggs were placed in 5 ml of a l/400 dilution of Clorox in SX in a 16 X 150 mm tube. Here they were kept for 10 minutes at room temperature with occasional agitation. Subsequent transfers were made by means of capillary (Pasteur) pipettes. After the hypochlorite bath, the eggs were sedimented through four 16 X 150 mm tubes, each containing 10 ml of SX. They were allowed to remain in each tube about 2 minutes prior to the next transfer. Finally, the eggs were transferred to 10 ml of SX in a Petri dish of lo-cm diameter t,o be individually placed in separate tubes.

of Eggs

Under a binocular dissecting microscope, and by using a flamed nichrome loop of about 1.4 mm inside diameter, eggs were transferred from the Petri dish to individual, cotton-stoppered, 16 X 150 mm tubes containing 2 ml of SX. Unhatched snails were clearly visible within their egg membranes and could at this time be chosen for viability and generally favorable appearance. Any with developmental abnormalities were discarded; eggs containing the largest and most active snails were selected. At this point sample eggs placed in bacteriologic culture medium provided a test of the surface sterilization process. Egg-containing tubes were then set aside to be observed at intervals for hatching. Although higher room temperatures have been used, a temperature of about 21°C was most frequently employed. Sterility

Testing

Routine sterility tests were carried out with 15 ml of Fluid Thioglycollate Medium (Baltimore Biological Laboratory) in 18 x 150 mm tubes. In addition, other supplementary media (veal or heart infusion broth) have been used as needed. Snails (except those inoculated into the medium immediately after surface sterilization) with a minimal amount of SX solution were inserted into the thioglycollate medium with capillary pipettes and generally crushed against the side of the tube. Additional sterility tests were occasionally made with some of the SX solution remaining in the tube from which the snail was removed. Tubes to be tested were agitated to suspend particulate matter prior to removal of either snail or fluid samples. Sterility tests were incubated for at least 24 hours at 37°C and held at room temperature for a week or more prior to evaluation. Subcultures were prepared when the first sterility test was considered inconclusive.

SECURING

Hatching

11

TABLE II and Sterility Test Results on L. stagnalis Eggs Surface Sterilized Sodium Hypochlorite and Placed in SX Solution NO.

Expt.

snails

1 2 3

14 15 5

Total Over-all Over-all

Lymnaea FREE FROM BACTERIA

Per cent hatched

Hatching prior to:

A. Egg masses “pretreated” 93 100 60

G percentage of snails hatching: 91 percentage of hatched snails found

bacteria-free:

with

Per cent of hatched snails found bacteria-free

6 days 3 days 6 days

85 93 100

90

B. Egg masses not “pretreated” 1 2 3 4 5 6 7 Total Over-all Over-all

4 10 9 6 7 9 10

12days 13 days 8 days 7 days 4 days 6 days 7 days

75 60 100 100 100 100 100

3 percentage of snails hatching: 91 percentage of hatched snails found RESULTS

The snail eggs generally hatched within several days after treatment (Table II, Fig. 4), although eclosion sometimes failed to take place even after several weeks had elapsed. A sample of 68 treated snails ranged from 1.0 to 2.0 mm (mean, 1.6 mm) in shell length within 1 week after hatching. (These measurements were made with the aid of an ocular grid through the culture tube walls and do not include exclusively bacteria-free individuals.) The hatchability and general vitality of the young snails seemed to depend upon judicious choice of vital specimens prior to individual tubing. Such hatched specimens, kept in 2 ml of SX at room temperature, could be generally maintained without feeding for 1 week but began to show a noticeable increase in mortality during the second week. One factor contributing to the loss of snails was their tendency to crawl out of the salt solution and subsequently to dry on the side of the tube. Constant surveillance was necessary

bacteria-free:

100 33 89 66 57 100 100

80

so that such snails could be detected and shaken down into the fluid before excessive dehydration took place. DISCUSSION

The method described above has proved to be successful in the absence of antibiotics for the routine isolation of bacteria-free L. stagnalis jugularis (Say). The data included illustrate the variability usually encountered with the method. Of over 500 snails already treated, the over-all incidence of contamination did not differ appreciably from that indicated by the experiments shown. Sodium hypochlorite, previously reported by Chernin (1957) as a surface disinfectant for A. glabratus eggs, has been demonstrated in this work to be effective for L. stagrralis jugularis. Other workers (Lapage, 1933 ; Glaser and Stoll, 1940; Weinstein and Jones, 1956) have recognized it to be a useful compound for similar disinfection procedures on other organisms. The salt solution employed (SX) was de-

FRIEDL

FIG. 4. Young L. stagnaZis jrrgularis men was about 18 days old.

newly hatched from a surface sterilized

rived initially as a modification of the ‘,handling solution” used by Chernin (1957). A comparison with “handling solution” will reveal a lower total solute content in the present formulation of SX (the concentrations of most of the major salts having been reduced by one-half) plus the addition of certain biologically significant trace substances. The chelating agent, disodium EDTA, has also been included as a carrier for certain of these trace substances. After treatment and selection, newly hatched snails possess, as far as can be seen, no gross abnormalities in appearance or activity (Fig. 4) ; nevertheless, their degree of physiologic normality under bacteria-free conditions remains yet to be demonstrated by extended periods of in vitro culture. ACKNOWLEDGMENTS

The author is grateful for the hospitality assistance extended to him in the laboratory

a.nd of

egg. This bacteria-free

speci-

Dr. Norman R. Stall while a Guest Investigator of The Rockefeller Institute. He is indebted to Dr. Stall personally for the help given him when problems concerning research were encountered and for his comments on this and previous manuscripts. Information regarding the identity of the snails used in this work was most kindly provided by Drs. H. A. Rehder and J. P. E. Morrison of the U.S. National Museum. REFERENCES

E. 1957. A method of securing bacteriologically sterile snails (Australorbis glabratus). Proceedings of the Society for Exfierimental Biology and Medicine 96, 204.210.

CHERNIN,

CIIERNIN, E., AND SCHORK, A. R. 1959. Growth in axenic culture of the snail Australorbis glabratus. American Journal of Hygiene 69, 146.160. GLASER, R. W., AND STOLL, N. R.

1940. Exsheathing and sterilizing infective nematode larvae, Journal of Parasitology 26, 87-94. G. 1933. Cultivation of todes. A’ature 131, 5X3-584.

LAPAGE,

parasitic

nema-

SECURING

Lymnaea

E. H. 1959. A technique for securing and maintaining bacteriologically sterile prosobranch snails. Transactions of the American Microscopical Society 78, 256-261.

MICHELSON,

FREE

FROM

BACTERIA

13

P. P., AND JONES, M. F. 1956. The in vitro cultivation of Nippostrongylus muris to the adult stage. Journal of Parasitology 42, 213-236.

WEINSTEIN,