344
ParasitologyToday, vol. 8, no. I0, 1992
explain the drug's selective action against them. It seems more likely that these parasites have unique proteins. The stereo-specificity of PZQ action is further indication that it exerts its effects via interaction with a protein. The EDs0 for L-PZQ eliciting contraction 2~ or for inducing tegumental disruption 22 in Schistosoma mansoni is about half that of the racemic PZQ. The action of PZQ on schistosomes is also dependent on the developmental stage of the schJstosome 22'23, with juvenile worms being much less susceptible than adult worms, This stage specificity could be well explained by stage-specific differences in protein expression. W e believe that all of this information taken together supports the hypothesis that PZQ interacts with a specific, unique protein involved in Ca 2+ homeostasis in trematodes and cestodes (Fig, 2). A better understanding of the selective action of PZQ against these parasites will require the identification of both the molecule with which PZQ interacts and the site of this interaction within the worms. Identification of the molecular targets of PZQ action, whether protein or lipid, may be accomplished through w o r k with whole
worms or worm homogenates, but it is clearly of interest exactly where in the worm these molecules are located and what physiological mechanisms are altered there. Studies involving the use of whole parasites will provide little information about the site of PZQ action; rather, this will require new approaches that use specific cells and tissues of the parasites.
References
I World Health Organization (I 989) Document WHO/SCHISTO/89.102 2 Becket, B. et al. (1980) Z. Parasitenkd. 63, 113-128 3 Shaw, M,K. and Erasmus, D,A. (1987) Parasitology 94, 243-254 4 Fetterer, R.H., Pax, R.A. and Bennett, J.L. (I 980) Eur.J. Pharmacol.64, 31-38 5 Sabah, A.A. et al. (I 985) Exp. Parasitol. 60, 348-354 6 Harder, A., Andrews, P. and Thomas, H. (1987) Parasitol.Res. 73, 245-249 7 Klaassen, C.D. and Eaton, D.L. (1991) in Casarett and Doull's Toxicology(Amdur, M.O., Doull, J. and Klaassen,C.D., eds), pp 12-49, Pergamon Press 8 Bricker, C.S. et al. (1983)Z. Parasitenkd 69, 61-71 9 Brindley, P.J. et al. (1989) Mol. Biochem. Parasitol. 34, 99-108 10 Sauma, S.Y., Tanaka, T.M. and Strand, M. ( 1991) Mol. Biochem.Parasitol.46, 73-80
I I Gomme, J. and Albrecthsen, S, (I 988) Camp. Biochem. Physiol.90, 651-657 12 Teilens,A.G.M., van den Heuvel,J.M. and van den Bergh, S.G.(1990)Mol. Biochem.Parasitol. 39, 195-202 13 Thompson, D.P., Pax, R.A. and Bennett, J.L. (I 982) Parasitology85, 163-178 14 Depenbusch,J.W. et al. ( t 983) Parasitology87, 61-73 15 Wolde Mussie, E. et al. (1982) Exp. Parositol. 53, 270-278 16 Xiao, S.H. et al. (1984)J. Parasitol. 70, 177-179 17 Nechay, B.R., Hillman, G.R. and Dotson, M.J. (1980)J. Parasitol.66, 59(~600 18 Pax, R.A., Thompson, D.P. and Bennet:, J.L. (1983) in Mechanism of Drug Action (Singer, T.P., Mansour, T.E. and Ondarza, R.N., eds), pp 187-196, Academic Press 19 Harder, A., Goossens, J. and Andrews, P. (1988) Mol. Biochem.Parasitol.29, 55-60 20 Schepers,H. et al. (I 988)Biochem. Pharmacol. 37, 1615-1623 21 Andrews, P. (1985) Pharmacol. Ther. 29, 129-156 22 Xiao, S.H. and Catto, B.A. ( 1989)j. Infect. D~s. 159, 589-592 23 Sabah, A.A. et al. (1986) Exp. Parasitol. 61,294-303 24 Shaw, M.K. (1990) Parasitology 100, 65-72 Tim Day and ]ames Bennett ore at the Department of Pharmacology and Toxicology and Ralph Pax is at the Deportment of Zoology, Michigan State University, East Lansing, MI 48824, USA.
Transformation of Caenorhabditis elegans With Genes From Parasitic Nematodes W.N. G rant Our knowledge of many aspects of the molecular biology of animal parasitic nematodes has rapidly expanded in recent years but the classical genetic analysis of this group of organisms has yet to emerge as a viable discipline. For example, it is not possible to routinely perform crosses between single males and females to examine the genetic basis of even simple phenotypes such as anthelmintic resistance. This has meant that the function of many cloned parasite genes can only be inferred from sequence comparison with genes from other organisms where the function is known, or by correlation of DNA polymorphisms linked to the gene with phenotypic differences between strains or individuals. In the absence of classical genetic techniques, a molecular solution is to transform a
suitable host with the gene of interest, but what de/fnes a suitable host? Here, Warwick Grant describes recent work that aims to provide such a host. As more molecular data on a diverse range of parasitic nematodes have accumulated, two general features have emerged. First, the function of genes that have been cloned using nonfunctional criteria (eg, immunological criteria for potential vaccine and immunodiagnostic antigen genes) has often proven elusive. T w o good examples of this are two candidate vaccine antigen genes cloned from Trichostrongylus colubriformis. One contains a domain with striking homology to the gut active peptide valosin ~ and the second contains a
globin-like domain 2. These homologies suggest possible functions for these antigens, but how can these hypothetical functions be tested? If this problem arose in yeast or Drosophila, the answer would be to first turn to the genetic system of those organisms to look for informative mutants so that the link between disruption of the gene and its normal function could be made, In the absence of the genetic means to do this, transformation of an appropriate host with the candidate gene has often been employed as a means of establishing or confirming function. For parasitic nematodes, transformation is the only option available and the choice of transformation host is an important consideration. Ideally one would like a host whose biology is as similar as ~) 1992.ElsevierSciencePublishersLtd.(UK)
345
Parasitology Today, vol. 8, no. I O, 1992
possible to that of the gene's original owner so that the biological function of the gene's product will be faithfully represented. For example, there is little point in transforming E. coil with a parasite collagen gene if ~t doesn't have a cuticle into which the collagen protein can be inserted. The second general feature to emerge from recent work is that several basic aspects of nematode molecular biology appear to be conserved between diverse nematode genera but not between the Nematoda and other phyla. An excellent example of this is a mechanism of messenger RNA (mRNA) trans-splicing which shares some features with mRNA processing from all eukaryotes, but the details of which are nematode specific3'4. Nematode collagen genes display a similar blend of generalized and nematodespecific featuress, with the latter presumably reflecting the specialized role of collagen in the cuticle. This reinforces the point made above, that the choice of transformation host is crucial if expression of the gene in question is to produce a phenotype related to its normal function, in vivo, and narrows the field of potential candidates considerably. C a e n o r h a b d i t i s elegans as a
T r a n s f o r m a t i o n Host
The free-living nematode Caenorhabditis elegans has been intensively studied from many points of view for two decades6. Initial interest was focussed on aspects of basic biology that seemed remote from the concerns of most parasitologists but, as the field has matured sufficiently for generalizations to be made, it has become clear that this organism has much to offer us at a fundamental level. Most: important in the current context is the fact that C. elegans is a nematode with a powerful and sophisticated genetic system and that it can be transformed. The basic method of transformation of C. elegans is extremely simple and requires little specialized equipment (Fig. I, Box I and Refs 7-9). Cloned DNA is injected directly into the ovary of an adult hermaphrodite and 20-50 transformants are then recovered in the first generation (F I) progeny of the injected worm. These F I transformants are of two types. The majority do not pass the transformed phenotype on to succeeding generations, ie. are transiently transformed, but approximately 10%, ie. 2-5 worms per injected hermaphrodite, are heritably trans-
Cloned PraurgSr~eisgtene~gegenCa)ndidate/
n
/7
/
Needle filled
with cloned DNA
Worm ovary injected with cloned DNA
1
Transformed progeny containing parasite gene
1
Transformants to be assayed for drug resistance
Fig. I. Transformation of C. elegans by microinjection of cloned DNA into the ovary. A finely drawn glass needle is inserted into the distal portion of each gonad arm of an adult hermaphrodite which has been immobilized on a dry agarose pad. The gonad arm is filled with DNA solution and the worm is then allowed to recover and lay eggs. Each successfully injected worm will produce up to 50 transformed progeny.
formed and can give rise to an indefinite succession of transformed progeny in the following generations. The transforming DNA is inherited as an extrachromosomal array composed of many copies of the original sequence7'9, so the pattern of inheritance is not Mendelian and varies to some extent between transformants. However, it is stable in
the sense that transformed progeny can be recovered indefinitely at each generation, ie. permanent transgenic strains can be established. What are the pertinent features of this system? First, C elegans generally reproduces as a self-fertilizing hermaphrodite, so that the establishment of clonal transgenic strains from each of the initial
Box I. Equipment for Transformation • An inverted microscope equipped with differential interference contrast (Normarski) optics and, if possible, a free-sliding oil cushion stage to allow for the easy positioning of the worms for injection. • A micromanipulator to hold and manoeuvre the needle for injection. Injections are perfomed from a low (<5 °) angle, so the micromanipulator must be able to be positioned at approximately the same level as the microscope stage and adjusted so that the needle is almost horizontal. • A needle puller to prepare needles for injection. • A vibration-free table for the microscope and micromanipulator. This is not absolutely essential, but it certainly helps. • An incubator capable of maintaining 20°C, required for the maintenance of C. elegans.
346 transformants is straightforward and rapid, with a generation time of only four days at 20°C. C. elegans also produces occasional males that can be used as a means of transferring genes (including transgenes on extrachromosomal arrays) between strains, so that once a transgenic strain has been established the phenotype conferred by the introduced D N A can be investigated in a range of genetic backgrounds using conventional genetic methods without the requirement of making a new transformed strain for each combination required. Second, D N A of virtually any form can be used. There is no requirement for specific sequences in the transforming DNA. For example, the original reports used plasmid D N A devoid of any nematode sequences at all7. Consequently, no special manipulation of the cloned D N A is necessary before it can be used for transformation: plasmid, lambda, cosmid and even yeast artificial chromosome D N A have been successfully used. Furthermore, co-injected DNAs are incorporated into a single chimeric array and co-expressed. A dominant marker for transformation is available (the rol-6 gene, which confers an easily scored change in the movement of the worms) 9 and this can be simply mixed with the D N A of interest, ie. there is no need to physically link the marker and the D N A of interest, in vitro, before injection because the worm does this effectively postinjection, in viva. It is not necessary, therefore, to rely on a readily detectable phenotype from expression of the parasite transgene, because the transformants can be detected in the first instance by their expression of the rol-6 marker. Third, the injected D N A must contain all the sequences necessary for its own expression, ie. it must be genomic D N A with the correct promoters, splice signals, etc. When the injected D N A is from a parasite, the C. elegans transcription machinery must be able to recognize and correctly interpret these signals. How often this requirement will be fulfilled is not known but it has the fortuitous spin-off that successful expression in this system will, by definition, delineate the key regulatory elements flanking the gene. Finally, promoter function and the sites and levels of expression of genes can be examined in this system 1°-~2 These studies suggest that, in general, the introduced genes function in much the same way as do their normal chromosomal counterparts but that some care must be taken in interpreting
Parasitology Today, vol. 8, no. I0, 1992
the results. This appears to relate more to the level 8-~° than to the timing or location of expression, and is presumably associated with the varying degrees to which the worm can regulate many copies of genes normally present at two copies per genome. Transformation with Parasite Genes How can this technology be applied to parasites? Two obvious avenues are the investigation of parasite genes cloned for 'practical' reasons and, more speculatively, to ask whether there are functional parasite homologues of known C. elegans genes. Resistance to the benzimidazoles (BZs) is due to a decrease in the amount of drug bound to parasite tubulin 13 and evidence has been presented implicating a single ~-tubulin locus in this resistance in Haemonchus contortus ~4, although other loci may be involved Is. We have similar data for T. colubriformis (W.N. Grant and L.J. Mascord, unpublished). These data show that a I3tubulin restriction fragment length polymorphism (RFLP) is correlated with resistance but do not constitute proof that the tubulin gene linked to the RFLP causes the resistance. A J3-tubulin gene that appears to be associated with BZ susceptibility in T. colubriformis has been used to transform C. elegans (W.N. Grant, unpublished) and a transgenic strain carrying the parasite gene as an extrachromosomal array with the rol-6 marker has been established. Does the parasite tubulin transgene have any effect on the BZ susceptibility of C. elegans? The preliminary answer is affirmative. Resistance to BZs is generally either dominant or partially dominant in C. elegans h6. When either of the parents in a cross between resistant and susceptible C. elegans carries the parasite I]-tubulin transgene, the degree of dominance of the resistance is significantly reduced compared to a control cross without the parasite gene, ie. the parasite gene produces a phenotype consistent with its proposed function. The hypothesis that particular tubulin genes may determine different levels of BZ susceptibility in the parasite can now be tested by examining their differing effects on the BZ susceptibility of transgenic C. elegans. In order to explore the function of parasite homologues of C. elegans genes, a T. colubnformis genomic clone containing a sequence that crosshybridizes to the gene-specific sequence from a C. elegans collagen gene (the dpy-13
gene, which is involved in specifying cuticle morphology w) has been isolated and used to transform dpy-13 mutants (W.N. Grant and D. Riddle, unpublished). Our expectation was that the parasite gene would complement or rescue the dpy-13 mutation but, to our surprise, the transformants exhibited a much more severe derangement of normal cuticle formation than did the original mutant. This is still consistent with a role for this parasite sequence in cuticle formation and suggests that the parasite gene product interferes with, rather than complements, cuticle formation in a dpy-13 background. Prospects These results show that at least two parasite genes can be maintained and expressed in C. elegans and that the phenotypes they produce are consistent with their proposed functions. As more parasite genes are cloned and as the C. elegans genome sequencing project ~8 uncovers more genes with potentially interesting parasite homologues, the availability of a transformation system will become increasingly important. The technique is simple and yields rapid results. Give it a try with your favourite gene!
References I Savin, K.W. et ol. (1990) Mol. Biochem. Parasitol. 41, 101-108 2 Frenkel, M.J. et at. (1992) MoL Biochem. Parasitol. 50, 27-36 3 Nilson, T.W. (1989) Exp. Parasitot. 69, 413-416 4 Donelson, J.E. and Zeng, W. (1990) Parasitology Today 6, 327-334 5 Kingston, I.B. ( 1991) Parasitology Today 7, 11-15 6 Wood, W.B. (1988) The Nematode Caenorhabditis elegans,Cold Spring Harbor Press 7 Stinchcomb,D.T. et al. (I 985)Mol. Cell. Biof. 5, 3484-3496 8 Fire,A. (1986) EMBOJ. 5, 2673-2680 9 Mello, C.C. et al. (1991) E_MBO J. 10, 3959-3970 10 Fire, A. and Waterston, R.H. (1989) EMBOJ. 8, 3419-3428 I Fire, A., White Harrison, S. and Dixon, D. (1990) Gene 93, 189-198 2 Way,J.C.and Chalfie,M. (1988)Cell 54, 5-16 3 Lacey,E. (I 988) Int. J. Parasitol. 18, 885-936 14 Roos, M.H. (1990) Parasitology Today 6, 125-127 15 Geary, T.G. et al. (1992) Mol. Biochem Parasitol. 50, 295-306 16 Driscoll, M. eta/. (1989) J. Cell Biol. 109, 2993-3003 17 van Mende,N. et al. (1988) Cell 55, 567-576 18 Sulston,J. et al. (I 992) Nature 356, 37-41 Warwick Grant is at the CSIRO Division of Animal Health, Private Bag, Armidale, NSW 2350, Australia.