**3.2 Cryopreservation of nematode without cryoprotectant additive**

The simplest method for nematode cryopreservation was reported on ruminant nematodes. Infective juveniles were cooled directly in liquid nitrogen vapor after being unsheathed by sodium hypochlorite, and suspended in physiological saline (Campbell and Thomson, 1973; Van Wyk et al., 1977). For example, infective juveniles of sheep nematodes (*Haemonchus contortus, Ostertagia circumcincta, Trichostrongylus axei, Trichostrongylus colubriformis, Nematodirus spathiger and Oesophagostomum columbianum*) and the bovine nematodes (*Haemonchus placei, Ostertagia ostertagi, Nematodirus helvetianus, Oesophagostomum radiatum, Cooperia pectinata and Cooperia punctata*) survived these simple cryopreservation procedures.

James (1985) suggested that the presence of natural cryoprotectants plays a role in the cryosurvival of domestic animal parasitic nematodes. Bai et al. (2004) demonstrated in *Steinernema carpocapsae* and *Heterorhabditis bacteriophora* that the cryo-survival rates of infective juveniles are positively correlated with worm concentration during the cryoprotectant glycerol incubation step. The survival rates ranged from about 20% to 100%, which was proportional to worm concentration of 120-12,000 per ml (Bai et al., 2004). Infective juveniles indeed produce cryoprotecting molecules such as trehalose and glycerol,

Cryopreservation of Embryos from Model Animals and Human 271

unfrozen worms (James, 1981). On the other hand, 70% of the *Onchocerca microfilariae* were

A similar vitrification methodology was also applied to the plant parasitic nematode *Meloidogyne Graminicola*, a rice root-knot nematode (Bridge & Ham 1985). The second-stage larvae were pre-incubated in 10% ethylene glycol at 37C for 15 minutes and then incubated in 40% ethylene glycol for 30-45 minutes. The worms were then rapidly cooled in liquid nitrogen in the same manner as James (1981) on *Schistosoma mansoni*. This two-stage vitrification procedure on plant nematodes was later modified by Triantaphyllou & McCabe (1989) who replaced the glass coverslip slivers with a small strip of chromatography paper as a carrier device. A survival rate of up to 90% was obtained. The author reported that the modified two-step method produced satisfactory results on other plant parasitic nematodes also, such as some *Meloidogyne* and *Heterodera* species (Triantaphyllou & McCabe, 1989).

An entirely different treatment methodology was developed primarily for vitrifying the entomopathogenic nematodes in the genera *Steinernema* and *Heterorhabditis*, which are effective as a biological control agent for insect pests in agriculture (Popiel & Vasquez, 1991). *Steinernema carpocapsae* were pre-incubated in 22% glycerol at room temperature for 24 hours and then in ice cold 70% methanol for 10 minutes. After removal of the majority of methanol by centrifugation, concentrated worms in methanol (i.e. 20l) were spread on a small strip of filter paper before plunging into liquid nitrogen for rapid cooling. Up to 95% post-thaw survival rate could be obtained. On the other hand, a post-thaw survival rate of about 55% was obtained on *Heterorhabditis bacteriophora* when the optimal 14% glycerol was

Curran et al. (1992) further optimized the protocol by replacing the centrifugation with a filtration step to remove glycerol prior to the methanol incubation. The optimal conditions for glycerol incubation for a number of entomopathogenic nematodes were also determined. Optimal glycerol pre-incubation conditions were determined to be 18% glycerol for 24 hours for *Steinernema carpocapsae*, 17% glycerol for 72 hours for *Heterorhabditis bacteriophora* and 13.8% glycerol for 72 hours for *Steinernema feltiae* and *Steinernema glaseri* (Curran et al., 1992). Other than that, 167 entomopathogenic nematodes were found to be able to survive the cryopreservation treatments, proving the feasibility of cryobanking of these worms. The mean survival rate of the *Steinernema* species is 58% (ranging from 25% to 97%) and that of

Based on the modification by Curran et al. (1992), Nugent et al. (1996) optimized cryopreservation on seven isolates of *Heterorhabditis*. Up to 8 days of pre-incubation in 11% or 15% glycerol is optimal for cryopreserving a couple of isolates. Nugent (1996) also found that glycerol can be replaced by DMSO in the pre-incubation step. For example, incubation of isolate HI82 in 8% DMSO for 3 days yielded a survival rate of about 80%, similar to those

To the best of our knowledge, unlike mammals, there have been no studies of interactions between nematodes and cryoprotectants. Conversely, different protocols have been developed independently by different groups for specific worm species. Whether the different protocols are indeed applicable to other groups of nematodes or not requires

viable and remained infective after vitrification and thawing (Ham et al., 1981).

**3.5 Vitrification by a glycerol/methanol two-step procedure** 

*Heterorhabditis* species is 51% (ranging from 25% to 87%).

used (Popiel & Vasquez, 1991).

pre-incubated in 15% DMSO.

in response to thermal and other environmental stresses (Jagdale & Grew, 2003; Qiu & Bedding, 2002). The trehalose content in *Steinernema carpocapsae* increases from 4% to 8% after being incubated in 22% glycerol for 18 hours, before the animals are further processed for cryopreservation (Popiel & Vasquez, 1991). Production of natural cryoprotectants by the animal itself could, therefore, be the key to good animal survival in this cryopreservation procedure. Exploring an efficient way to induce the production of the natural cryoprotectants can improve cryo-survival. Identifying the molecular pathway responsible for cryoprotectant production may help make cryopreservation of these species simpler.
