Stage-specific gene expression in Teladorsagia circumcincta (Nematoda: Strongylida) infective larvae and early parasitic stages☆
Introduction
Dramatic physiological changes occur during the transition from the free-living to the parasitic phase of a nematode’s life cycle. These physiological changes are initiated, and accompanied by, up- and down-regulation in expression levels of suites of genes associated with changes in the nematode’s environment and with nutrition, development, the onset of sexual maturation and establishment in the host (for recent reviews see Nisbet et al., 2004, Nikolaou and Gasser, 2006). A detailed understanding of these processes provides an opportunity to define drug and vaccine targets in parasitic helminths. Previous studies of the mechanisms underlying the transition from infective to parasitic stages of nematodes have used various molecular technologies, the more recent employing microarrays generated from pools of expressed sequence tags (ESTs) derived from cDNA libraries (e.g. Moser et al., 2005). By utilising clustering software prior to selection of ESTs to be printed on the microarrays, redundancy can be reduced, however the majority of genes represented by the ESTs on the microarray slides are unlikely to be differentially expressed between two test populations. For example, only 2.7% of the genes represented by 4191 ESTs derived from an Ancylostoma caninum L3 cDNA library were shown to be differentially regulated between infective and parasitic stages using this method (Moser et al., 2005). Stage-specific gene expression has been implied via the study of relative representation of ESTs in cDNA libraries synthesised from different developmental stages of nematodes (e.g. Hoekstra et al., 2000, Thompson et al., 2005) however doubts now exist over the merits of this approach of comparing differences in gene expression (Thompson et al., 2006). In the present study, we have used suppression subtractive hybridisation (SSH, Diatchenko et al., 1996) to allow the enrichment of genes expressed in a stage-specific manner between infective, exsheathed L3 (xL3) and early L4 Teladorsagia circumcincta prior to gene expression profiling by microarray. These technologies have enabled identification of suites of differentially expressed genes which, via bioinformatic analyses, have provided insights into the biochemical and physiological processes associated with both the free-living stages and their adaptation to a parasitic lifestyle.
Section snippets
Parasite material
Teladorsagia circumcincta L3s were harvested from coproculture and exsheathed by exposure to sodium hypochlorite as detailed in Jackson et al. (2004). L4s were collected at 8 days p.i. from the abomasal mucosa of sheep infected with 50,000 L3s using previously published methods (Knox and Jones, 1990). The worms were washed extensively in PBS, snap-frozen and stored in liquid nitrogen.
SSH
Frozen nematodes were homogenised in liquid nitrogen using a pre-cooled mortar and pestle. TRIZOL® reagent
Suppression subtractive hybridisation (SSH)
The analysis of subtraction efficiency, performed using 18,424 cloned plasmids (9212 from the xL3-specific library and 9212 from the L4-specific library) confirmed the high efficiency of the SSH procedure. Un-subtracted cDNA from xL3s hybridised almost exclusively to colonies generated from the xL3-specific subtracted library (Fig. 1A) and un-subtracted cDNA from L4s hybridised almost exclusively to those colonies generated from the L4-specific subtracted library (Fig. 1B), demonstrating the
Discussion
Here, suppression subtractive hybridisation has proven to be a highly effective tool for enriching cDNA libraries for stage-specific and stage-enriched transcripts from xL3 and L4 T. circumcincta. Overall the “shift” of gene expression from L3 to L4 appears to reflect the changes in nutrition, aerobic/microaerobic metabolism and rapidity of growth (e.g. collagen synthesis in L4) associated with the transition to parasitic lifestyle.
The transcript for one particular metabolic enzyme, GTP-CH,
Acknowledgements
Funding support through SEERAD is gratefully acknowledged. We also thank Liz Jackson and Dave Bartley (Parasitology Division, MRI) and David Kennedy, Jim Williams and Roy Davie (Clinical Division, MRI) for the production of parasite materials and expert animal care. Thanks also to Clare Booth, SCRI for sequencing the ESTs, Gill Campbell, Rowett Research Institute for printing the microarrays and Jill Sales, BIOSS for assistance with statistical analysis.
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