Studies utilizing schistosomes are greatly hindered by the unavailability of genetically uniform schistosome material. We have routinely maintained 5 male and 5 female clones of Schistosoma mansoni by serial microsurgical transplantation of sporocysts into Biomphalaria glabrata snails for 2.5 years (Cohen and Eveland, 1984, Expl Parasit. 57: 15-19). During our studies it became apparent that this technology would be enhanced greatly by the cryopreservation of clones. Although sporocyst cryopreservation has not been reported for any trematode species, it seemed likely that this stage in the schistosome life cycle could be cryopreserved because of its relative undifferentiation. In addition, cryopreserving sporocysts would allow: the storage of virtually unlimited numbers of clones with interesting phenotypes (e.g., drug resistance, high or low infectivity or immunogenicity) in multiple locations; the easy transportation of clones between laboratories; a reduction in the risk that point mutations may occur during subpassaging; and the preservation of new field isolates and their genetic diversity. Although cryopreservation of S. mansoni schistosomula has been reported (James and Farrant, 1977, Trans. R. Soc. trop. Med. Hyg. 71: 498-500; Stirewalt et al., 1979, Expl Parasit. 48: 272-281; James, 1981, Expl Parasit. 52: 105-116), genetically identical cercariae cannot be perpetuated by this method, because recombination occurs when adults derived from thawed schistosomula pair within their mammalian host. We have successfully cryopreserved S. mansoni sporocysts in our laboratory. The microsurgical tranplantation techniques are described elsewhere (Cohen and Eveland, 1984, loc. cit.). Using aseptic procedures, parasitized hepatopancreas and ovotestis containing sporocysts were excised from S. mansoni-infected B. glabrata snails, placed in sterile Chernin's Balanced Salt Solution (BSS-Chernin, 1963, J. Parasit.), and cut into fragments. The fragments were put into BSS containing either 15-20% dimethyl sulfoxide (DMSO-Fisher, Fair Lawn, NJ), or 15-20% glycerol (Sigma, St. Louis, MO), respectively, and frozen in sterile 1.2 ml Nunc conical bottom, screw cap cryotubes (Vangard International, Neptune, NJ). Small quantities of tissue were frozen in each tube with just enough cryoprotectant added to completely cover the tissues. This facilitated rapid thawing and ensured that only freshly-thawed material was used. Tubes were then incubated at 22-24 C for 0.5 hr, 4 C for 0.5 hr, and -20 C for 0.5 hr, successively, then transferred into a Revco freezer (-70 C). After several days the contents were thawed by rapidly pipetting 37 C BSS into the tube. Tissues were then washed twice with BSS to remove the cryoprotectant. The thawed sporocysts were implanted immediately into anesthetized B. glabrata. Snails were maintained routinely in the dark (Cohen and Eveland, 1984, loc. cit.) at 28 C and exposed to light once a week to check for cercarial emergence, which began after 4 wk. The results are summarized in Table I. These results demonstrate that S. mansoni sporocysts are easily cryopreserved. DMSO was a suitable cryoprotectant but glycerol was not. Infectivities of thawed sporocysts were only slightly lower than those reported for unfrozen sporocysts from miracidially infected snails (Cohen and Eveland, 1984, loc. cit.). Studies are in progress to optimize the conditions for sporocyst cryopreservation and to determine whether subpassaged sporocysts can also be cryopreserved.