NIAID Schistosomiasis Resource Center

Maintenance of prepatent snails

Most laboratories that obtain infected snails on this contract receive them a few days after exposure to miracidia. The procedures listed below should serve as a guide to the maintenance of these species during the prepatent period, and one from which modifications can be made to suit each laboratory's needs. Several publications give in-depth discussions on the maintenance of Biomphalaria glabrata, Bulinus truncatus, and Oncomelania hupensis sp. snails (1-7).

Several things should be considered before maintaining snails.

Temperature. For best growth snails should be maintained between 24-27C. For infected snails, there is generally a higher level of mortality at temperatures around 27C, although the prepatent period will be longer at the lower temperature ranges. Each laboratory should maintain the temperature for their particular needs.

Water. Water that has a low chlorine content is best for rearing snails. Water that is suitable is often that obtained after passage through a charcoal, or carbon, filter, then aged (aerated) for 1-2 days before use. Alternatively, some laboratories rely on a local source of commercially-obtained bottled water, or on water obtained from an aquarium that has been set up at least several days in advance. The laboratory should test whether the water is suitable first by placing uninfected snails into it and observing their behavior and/or whether any snail mortality occurs after a few hours or days.

Container. Biomphalaria and Bulinus sp. snails can be maintained in almost any laboratory container, but do particularly well in shallow pans, as long as they are not overcrowded and the water changed once or twice a week.

Food. Most laboratories use lettuce leaves as the primary food source for Biomphalaria and Bulinus sp snails. Romaine lettuce is a good variety to use, but the leaves should be stripped from the rib and washed before use. If rearing small juvenile snails however a better food source is Nostoc, grown over autoclaved mud (see SOP for maintaining Oncomelania sp). Alternatively, Romaine leaves wilted by heat also can serve as a food source for juvenile snails, although care should be taken with the amount given, since bacterial growth in the container can quickly become a problem.

Lighting conditions. Snails can be kept under almost any lighting condition. Infected Biomphalaria and Bulinus sp. snails however will be stimulated to shed more cercariae under a light source. If one needs large numbers of cercariae at any one time, the snails can be placed in darkness for 1-2 days before shedding under light to maximize the harvest.

Routine maintenance of prepatent and shedding snails

1) Check temperature daily, and maintain a constant level of water in the container.

2) Remove dead snails as often as possible, and the remains of lettuce (for Biomphalaria and Bulinus sp.) before adding fresh lettuce.

3) Change water at least once a week, and add a pinch of lime (powder) or CaCO3 to each container. Contaminated water can be poured into a container with bleach for disinfection.

4) Infected snails should be gently washed/cleaned before checking for cercarial emergence. Biomphalaria glabrata and Bulinus truncatus snails can be checked for cercarial production around 4-6 weeks after exposure to miracidia (depending on the maintenance temperature). Place snails individually into small containers with water, and place under a light source for 1-2 hrs. Cercariae that emerge can be seen in the water using a dissecting microscope. Maximal cercarial production occurs around 2 weeks after initial cercarial emergence, although snails usually continue to release cercariae throughout their lifetime. Every time the infected snails are used for cercarial collection, they can be returned to the containers and remain for 3-5 days before the next shedding. Oncomelania sp. snails can be examined around 4-6 weeks after exposure to miracidia for evidence of patency by determining the presence of daughter sporocysts, visible through the shell. The sporocysts appear as whitish mottling in the hepatopancreas. This method permits selection of around 85% of the total snails that ultimately will become positive. Visually-negative snails are kept separately until patency, followed by crushing at the time the visually-positive snails are used. Cercarial shedding is not a routine way to screen infected Oncomelania snails. The positive snails are crushed for maximal collection of S. japonicum cercariae for mouse infections.

5) For infecting mammals, obtain cercariae from at least 20-30 snails to insure a balanced sex ratio (see SOP for exposure of mammals to cercariae).

Exposure of mammals to cercariae

Below is a brief description of exposure methods to mice and hamsters with cercariae of the three schistosome species. For most laboratories, the mouse is the preferred host for S. mansoni and S. japonicum, although the hamster is used by some investigators as the definitive host for S. mansoni. The normal laboratory host for S. haematobium is the hamster.

Percutaneous exposure of mice

Two methods are commonly used for percutaneous exposure of mice to cercariae: (1) tail exposure, and (2) abdominal exposure.

Tail exposure

1. S. mansoni cercariae are placed in a 12-mm x 75-mm glass or plastic test tube and conditioned water added to within 10-mm from the top of the tube.

2. A mouse is placed in a restraining tube, with its tail extending from the bottom of the tube. (Note: restraining tubes of various sizes can be obtained from commercial vendors).

3. Wipe the tail with a gauze sponge moistened with conditioned water to remove bedding debris and insert the tail into the tube containing cercariae.

4. Expose the mouse for approximately 1 hr, remove the mouse from the restraining tube and return it to its cage.

5. To estimate the success of cercarial penetration, examine the contents of the tube by emptying the contents into a petri dish, staining with iodine and examining with a dissecting microscope. Count intact cercariae and the detached cercarial bodies (if any). Under ideal conditions at least 90% of the cercariae should have penetrated.

Abdominal exposure

This method is the preferred method for exposure to S. japonicum cercariae, but can also be used for exposing to S. mansoni.

1. Anesthetize the mouse with an ip injection of a suitable drug (e.g. sodium pentobarbital, 60mg/kg).

2. Shave the abdomen with animal clippers.

3. Wipe the abdomen with moistened gauze.

4. Place the mouse on its back in a 10-cm watch glass, or in a slotted board, so that involuntary movements will not disturb the cercarial suspension.

5. For S. mansoni exposures, place a 18-mm high x 10-mm wide stainless steel ring on the abdomen, and pipette the desired number of cercariae into the ring. Since S. japonicum cercariae are notoriously "sticky", adhering readily to plastic or glass, other means of delivery are needed. Thus, S. japonicum cercariae are usually placed on the abdomen using a small hair-loop connected to the end of a Pasteur pipette.

6. Expose the mouse to cercariae for approximately 1-hr, remove the suspension from the ring (for S. mansoni), then return the mouse to its cage, without wiping the exposed skin site.

Abdominal exposure of hamsters

Identical procedures as that for mice can be used for exposing hamsters on the abdomen with either S. mansoni or S. haematobium cercariae.

Injection procedures

Some laboratories infect mice or hamsters by injecting cercariae subcutaneously (subQ) or intraperitoneally (ip), although this is not the natural infection route. For this, estimate the injection dose of cercariae by pulling a suspension into a 1-ml syringe fitted with a 21-ga. needle. Express the suspension into a counting dish, stain with iodine and count intact cercariae. Once the dose is determined, inject the desired number of cercariae by standard subQ or ip injection. Due to the sticky nature of S. japonicum cercariae, injection methods are not recommended.

Breeding snails

Biomphalaria and Bulinus sp.

Methodologies and maintenance conditions for Biomphalaria and Bulinus sp. snails are almost identical.

1. Adult snails are placed into shallow plastic trays (10" x 7") and maintained at a density of 10-15 per tray, and fed romaine lettuce. The trays are changed weekly, at which time egg masses are scraped from the wall as well as from the surface of the lettuce.

2. The egg masses are placed in petri dishes with conditioned water and incubated under ceiling light for one week, until hatching.

3. Approximately 50-100 newly hatched snails are transferred to petri dishes with Nostoc muscorum (Cyanobacteria) and mud, and are placed under continuous illumination. One the algae has been consumed (approx. 1 week) the snails are transferred to a fresh dish of Nostoc.

4. This process is repeated until the snails reach approximately 3-5mm in diameter, which is a suitable size for exposure to miracidia.

Oncomelania sp.

Most maintenance conditions, including temperature, water source, and lighting are the same as the conditions for rearing Biomphalaria and Bulinus (see above). Containers for the breeders, and the food source however are different. Nostoc muscorum is used as the major food source for Oncomelania sp. snails. N. muscorum suspensions are inoculated weekly into petri dishes containing a mound of autoclaved mud and sterilized water with 0.067% NaNO3. They are kept at 27°C for at least one week before use. The breeders are maintained in a petri plate containing Nostoc grown over the autoclaved mud base (see references 1 and 2).

1. Five to seven pairs of adult Oncomelania snails are placed in one petri dish containing mud that is placed along the periphery, and a small quantity of Nostoc. The dishes are kept under constant light.

2. Using a dissecting microscope, examine the mud for eggs. Each egg will be encased individually in a gelatinuous capsule. Transfer the eggs into a petri dish, along with some snail feces, mud and Nostoc.

3. At room temperature, and under constant light, most eggs will hatch within 3 weeks.

4. Newly hatched snails are placed in newly established dishes containing a small amount of Nostoc and mud, and allowed to grow under constant light.

5. After reaching 2-3 mm in shell length, the snails are ready for exposure to miracidia.

Exposing snails to miracidia

Snails can be exposed to miracidia either en masse or individually. For en masse exposure, miracidia can be withdrawn with a Pasteur pipette, under a dissecting microscope, and placed in a container with numerous snails. Typically around 10 miracidia per snail gives a good infection rate. For individual exposures, place each snail in a well of a 24-well plate, add a minimum amount of conditioned water, then add the appropriate number of miracidia. In both cases expose snails to miracidia for at least 1-2 hrs before placing them in appropriate rearing containers.

References

1) Bruce, J.I.,, Radke, M.G., and Davis, G.M. 1971. Culturing Biomphalaria and Oncomelania (Gastropoda) for large-scale studies of schistosomiasis. Biomedical Report No. 19, 406th Medical Laboratory, U.S. Army.

2) Bruce, J.I. and Liang, Y-S. 1992. Cultivation of schistosomes and snails for researchers in the United States of America and other countries. Journal of Medical and Applied Malacology 4:13-20.

3) Lewis, F.A., Stirewalt, M.A., Souza, C.P., and Gazzinelli, G. 1986. Large-scale laboratory maintenance of Schistosoma mansoni, with observations on three schistosome/snail host combinations. Journal of Parasitology 72:813-829.

4) Lewis, F. 1998. Schistosomiasis. Current Protocols in Immunology, John Wiley & Sons, Inc. 19.1.1-19.1.28.

5) Liang, Y-S., Bruce, J.I., and Boyd, D.A. 1987. Laboratory cultivation of schistosome vector snails and maintenance of schistosome life cycles. Proceedings of the First Sino-American Symposium 1:34-48.

6) Standen, O.D. 1951. Some observations upon the maintenance of Australorbis glabratus in the laboratory. Annals of Tropical Medicine and Parasitology 45:80-83.

7) Stirewalt, M.A. 1954. Effect of snail maintenance temperatures on development of Schistosoma mansoni. Experimental Parasitology 3:504-516.


		Standard Operation Procedures
		Maintenance of prepatent snails Most laboratories that obtain infected snails on this contract receive them a few days after exposure to miracidia. The procedures listed below should serve as a guide to the maintenance of these species during the prepatent period, and one from which modifications can be made to suit each laboratory's needs. Several publications give in-depth discussions on the maintenance of Biomphalaria glabrata, Bulinus truncatus, and Oncomelania hupensis sp. snails (1-7). Several things should be considered before maintaining snails. Temperature. For best growth snails should be maintained between 24-27C. For infected snails, there is generally a higher level of mortality at temperatures around 27C, although the prepatent period will be longer at the lower temperature ranges. Each laboratory should maintain the temperature for their particular needs. Water. Water that has a low chlorine content is best for rearing snails. Water that is suitable is often that obtained after passage through a charcoal, or carbon, filter, then aged (aerated) for 1-2 days before use. Alternatively, some laboratories rely on a local source of commercially-obtained bottled water, or on water obtained from an aquarium that has been set up at least several days in advance. The laboratory should test whether the water is suitable first by placing uninfected snails into it and observing their behavior and/or whether any snail mortality occurs after a few hours or days. Container. Biomphalaria and Bulinus sp. snails can be maintained in almost any laboratory container, but do particularly well in shallow pans, as long as they are not overcrowded and the water changed once or twice a week. Food. Most laboratories use lettuce leaves as the primary food source for Biomphalaria and Bulinus sp snails. Romaine lettuce is a good variety to use, but the leaves should be stripped from the rib and washed before use. If rearing small juvenile snails however a better food source is Nostoc, grown over autoclaved mud (see SOP for maintaining Oncomelania sp). Alternatively, Romaine leaves wilted by heat also can serve as a food source for juvenile snails, although care should be taken with the amount given, since bacterial growth in the container can quickly become a problem. Lighting conditions. Snails can be kept under almost any lighting condition. Infected Biomphalaria and Bulinus sp. snails however will be stimulated to shed more cercariae under a light source. If one needs large numbers of cercariae at any one time, the snails can be placed in darkness for 1-2 days before shedding under light to maximize the harvest. Routine maintenance of prepatent and shedding snails 1) Check temperature daily, and maintain a constant level of water in the container. 2) Remove dead snails as often as possible, and the remains of lettuce (for Biomphalaria and Bulinus sp.) before adding fresh lettuce. 3) Change water at least once a week, and add a pinch of lime (powder) or CaCO3 to each container. Contaminated water can be poured into a container with bleach for disinfection. 4) Infected snails should be gently washed/cleaned before checking for cercarial emergence. Biomphalaria glabrata and Bulinus truncatus snails can be checked for cercarial production around 4-6 weeks after exposure to miracidia (depending on the maintenance temperature). Place snails individually into small containers with water, and place under a light source for 1-2 hrs. Cercariae that emerge can be seen in the water using a dissecting microscope. Maximal cercarial production occurs around 2 weeks after initial cercarial emergence, although snails usually continue to release cercariae throughout their lifetime. Every time the infected snails are used for cercarial collection, they can be returned to the containers and remain for 3-5 days before the next shedding. Oncomelania sp. snails can be examined around 4-6 weeks after exposure to miracidia for evidence of patency by determining the presence of daughter sporocysts, visible through the shell. The sporocysts appear as whitish mottling in the hepatopancreas. This method permits selection of around 85% of the total snails that ultimately will become positive. Visually-negative snails are kept separately until patency, followed by crushing at the time the visually-positive snails are used. Cercarial shedding is not a routine way to screen infected Oncomelania snails. The positive snails are crushed for maximal collection of S. japonicum cercariae for mouse infections. 5) For infecting mammals, obtain cercariae from at least 20-30 snails to insure a balanced sex ratio (see SOP for exposure of mammals to cercariae). Exposure of mammals to cercariae Below is a brief description of exposure methods to mice and hamsters with cercariae of the three schistosome species. For most laboratories, the mouse is the preferred host for S. mansoni and S. japonicum, although the hamster is used by some investigators as the definitive host for S. mansoni. The normal laboratory host for S. haematobium is the hamster. Percutaneous exposure of mice Two methods are commonly used for percutaneous exposure of mice to cercariae: (1) tail exposure, and (2) abdominal exposure. Tail exposure 1. S. mansoni cercariae are placed in a 12-mm x 75-mm glass or plastic test tube and conditioned water added to within 10-mm from the top of the tube. 2. A mouse is placed in a restraining tube, with its tail extending from the bottom of the tube. (Note: restraining tubes of various sizes can be obtained from commercial vendors). 3. Wipe the tail with a gauze sponge moistened with conditioned water to remove bedding debris and insert the tail into the tube containing cercariae. 4. Expose the mouse for approximately 1 hr, remove the mouse from the restraining tube and return it to its cage. 5. To estimate the success of cercarial penetration, examine the contents of the tube by emptying the contents into a petri dish, staining with iodine and examining with a dissecting microscope. Count intact cercariae and the detached cercarial bodies (if any). Under ideal conditions at least 90% of the cercariae should have penetrated. Abdominal exposure This method is the preferred method for exposure to S. japonicum cercariae, but can also be used for exposing to S. mansoni. 1. Anesthetize the mouse with an ip injection of a suitable drug (e.g. sodium pentobarbital, 60mg/kg). 2. Shave the abdomen with animal clippers. 3. Wipe the abdomen with moistened gauze. 4. Place the mouse on its back in a 10-cm watch glass, or in a slotted board, so that involuntary movements will not disturb the cercarial suspension. 5. For S. mansoni exposures, place a 18-mm high x 10-mm wide stainless steel ring on the abdomen, and pipette the desired number of cercariae into the ring. Since S. japonicum cercariae are notoriously "sticky", adhering readily to plastic or glass, other means of delivery are needed. Thus, S. japonicum cercariae are usually placed on the abdomen using a small hair-loop connected to the end of a Pasteur pipette. 6. Expose the mouse to cercariae for approximately 1-hr, remove the suspension from the ring (for S. mansoni), then return the mouse to its cage, without wiping the exposed skin site. Abdominal exposure of hamsters Identical procedures as that for mice can be used for exposing hamsters on the abdomen with either S. mansoni or S. haematobium cercariae. Injection procedures Some laboratories infect mice or hamsters by injecting cercariae subcutaneously (subQ) or intraperitoneally (ip), although this is not the natural infection route. For this, estimate the injection dose of cercariae by pulling a suspension into a 1-ml syringe fitted with a 21-ga. needle. Express the suspension into a counting dish, stain with iodine and count intact cercariae. Once the dose is determined, inject the desired number of cercariae by standard subQ or ip injection. Due to the sticky nature of S. japonicum cercariae, injection methods are not recommended. Breeding snails Biomphalaria and Bulinus sp. Methodologies and maintenance conditions for Biomphalaria and Bulinus sp. snails are almost identical. 1. Adult snails are placed into shallow plastic trays (10" x 7") and maintained at a density of 10-15 per tray, and fed romaine lettuce. The trays are changed weekly, at which time egg masses are scraped from the wall as well as from the surface of the lettuce. 2. The egg masses are placed in petri dishes with conditioned water and incubated under ceiling light for one week, until hatching. 3. Approximately 50-100 newly hatched snails are transferred to petri dishes with Nostoc muscorum (Cyanobacteria) and mud, and are placed under continuous illumination. One the algae has been consumed (approx. 1 week) the snails are transferred to a fresh dish of Nostoc. 4. This process is repeated until the snails reach approximately 3-5mm in diameter, which is a suitable size for exposure to miracidia. Oncomelania sp. Most maintenance conditions, including temperature, water source, and lighting are the same as the conditions for rearing Biomphalaria and Bulinus (see above). Containers for the breeders, and the food source however are different. Nostoc muscorum is used as the major food source for Oncomelania sp. snails. N. muscorum suspensions are inoculated weekly into petri dishes containing a mound of autoclaved mud and sterilized water with 0.067% NaNO3. They are kept at 27°C for at least one week before use. The breeders are maintained in a petri plate containing Nostoc grown over the autoclaved mud base (see references 1 and 2). 1. Five to seven pairs of adult Oncomelania snails are placed in one petri dish containing mud that is placed along the periphery, and a small quantity of Nostoc. The dishes are kept under constant light. 2. Using a dissecting microscope, examine the mud for eggs. Each egg will be encased individually in a gelatinuous capsule. Transfer the eggs into a petri dish, along with some snail feces, mud and Nostoc. 3. At room temperature, and under constant light, most eggs will hatch within 3 weeks. 4. Newly hatched snails are placed in newly established dishes containing a small amount of Nostoc and mud, and allowed to grow under constant light. 5. After reaching 2-3 mm in shell length, the snails are ready for exposure to miracidia. Exposing snails to miracidia Snails can be exposed to miracidia either en masse or individually. For en masse exposure, miracidia can be withdrawn with a Pasteur pipette, under a dissecting microscope, and placed in a container with numerous snails. Typically around 10 miracidia per snail gives a good infection rate. For individual exposures, place each snail in a well of a 24-well plate, add a minimum amount of conditioned water, then add the appropriate number of miracidia. In both cases expose snails to miracidia for at least 1-2 hrs before placing them in appropriate rearing containers. References 1) Bruce, J.I.,, Radke, M.G., and Davis, G.M. 1971. Culturing Biomphalaria and Oncomelania (Gastropoda) for large-scale studies of schistosomiasis. Biomedical Report No. 19, 406th Medical Laboratory, U.S. Army. 2) Bruce, J.I. and Liang, Y-S. 1992. Cultivation of schistosomes and snails for researchers in the United States of America and other countries. Journal of Medical and Applied Malacology 4:13-20. 3) Lewis, F.A., Stirewalt, M.A., Souza, C.P., and Gazzinelli, G. 1986. Large-scale laboratory maintenance of Schistosoma mansoni, with observations on three schistosome/snail host combinations. Journal of Parasitology 72:813-829. 4) Lewis, F. 1998. Schistosomiasis. Current Protocols in Immunology, John Wiley & Sons, Inc. 19.1.1-19.1.28. 5) Liang, Y-S., Bruce, J.I., and Boyd, D.A. 1987. Laboratory cultivation of schistosome vector snails and maintenance of schistosome life cycles. Proceedings of the First Sino-American Symposium 1:34-48. 6) Standen, O.D. 1951. Some observations upon the maintenance of Australorbis glabratus in the laboratory. Annals of Tropical Medicine and Parasitology 45:80-83. 7) Stirewalt, M.A. 1954. Effect of snail maintenance temperatures on development of Schistosoma mansoni. Experimental Parasitology 3:504-516.