International Journal for Parasitology 36 (2006) 475–483 www.elsevier.com/locate/ijpara
Heritable transgenesis of Parastrongyloides trichosuri: A nematode parasite of mammals Warwick N. Grant a,*, Stephen J.M. Skinner a,b, Jan Newton-Howes a, Kirsten Grant a, Gail Shuttleworth a, David D. Heath a, Charles B. Shoemaker a,c a
AgResearch Ltd, Wallaceville Animal Research Centre, P.O. Box 40063, Upper Hutt, New Zealand b Living Cell Technologies NZ Ltd, P.O. Box 23566, Hunters Cnr, Auckland, New Zealand c Department of Biomedical Sciences, Tufts School of Veterinary Medicine, North Grafton, MA 01536, USA Received 1 August 2005; received in revised form 28 November 2005; accepted 2 December 2005
Abstract Germline transformation of a parasitic nematode of mammals has proven to be an elusive goal. We report here the heritable germline transformation of Parastrongyloides trichosuri, a nematode parasite whose natural hosts are Australian possums of the genus Trichosurus. This parasite can undergo multiple free-living life cycles and these replicative cycles can be maintained indefinitely in the laboratory. Transformation was achieved by microinjection of DNA into the ovary syncytium of either free-living or parasitic adult females. By selecting for the transgenic progeny of successive free-living life cycles, it was possible to establish and maintain transgenic lines. All three transgenic lines tested were shown capable of establishing patent infections in possums and to transmit the functional transgene to their progeny. The transgene, driven by the Pt hsp-1 promoter, was constitutively expressed in intestinal cells at all stages of both parasitic and free-living life cycles, although gene silencing appears to occur in some transgenic progeny. This is the first report of heritable transgenesis in a parasitic nematode of a mammal and we discuss a variety of previously inaccessible experimental avenues that will now be possible with this powerful model system. q 2005 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved. Keywords: Transgenic; Nematode parasite; Parastrongyloides trichosuri; Microinjection; Brushtail possum; Functional genomics
1. Introduction Parasitic nematodes of mammals are a large and diverse group of organisms that cause tremendous morbidity and economic loss but research on these pathogens has been hindered by the absence of methods to manipulate gene expression and define gene function (Grant and Viney, 2001; Viney et al., 2002; Boyle and Yoshino, 2003). This shortcoming is more evident now that substantial genomics resources are becoming available in the form of parasitic nematode expressed sequence tag (EST) collections (Parkinson et al., 2003), some of which have been the subject of extensive bioinformatic and gene expression analyses (McCarter et al., 2003; Mitreva et al., 2004a,b; Parkinson et al., 2004; Thompson et al., 2005) and of several growing genome sequence databases. There have been several recent reports of * Corresponding author. Tel.: C64 4 922 1506; fax: C64 4 922 1580. E-mail address:
[email protected] (W.N. Grant).
the application of RNAi (Fire et al., 1998) to parasitic nematodes (Hussein et al., 2002; Urwin et al., 2002; Aboobaker and Blaxter, 2003; Lustigman et al., 2004; Issa et al., 2005) but none for heritable germline transformation of a nematode parasite of mammals. Nematode parasites of mammals are difficult organisms work with, as they are generally obligate parasites that cannot complete a life cycle in vitro. In most cases, the parasitic stages of the life cycle cannot be manipulated while in their hosts nor be maintained in a healthy state outside their hosts. This limits ready access to only those stages of the life cycle, which develop outside the host, or to those aspects of their biology expressed during short-term in vitro maintenance of the parasitic stages. Early attempts to define gene function (e.g. anthelmintic resistance) relied on expression of parasite genes in the free-living nematode Caenorhabditis elegans (Grant, 1992; Kwa et al., 1995) and this continues to be a useful method for studying parasite genes and their promoters where orthology is likely (for example, see Couthier et al., 2004).
0020-7519/$30.00 q 2005 Australian Society for Parasitology Inc. Published by Elsevier Ltd. All rights reserved. doi:10.1016/j.ijpara.2005.12.002
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Its obvious weakness is that although some aspects of a parasite’s nematode biology may be recapitulated in C. elegans, it is unlikely that parasite-specific biology will be faithfully assayed in this system. There are several reports of transient expression systems utilising parasites removed from their host (Davis et al., 1999; Jackstadt et al., 1999; Higazi et al., 2002, 2004) but the applicability of these models is limited by the fact that parasitic stages can be maintained in vitro for only short periods of time. Lok and Massey (2002) have reported progress towards transgenesis in Strongyloides stercoralis, a human parasite which in common with other Strongyloides species, has an unusual life cycle, which can include a single free-living generation (Shiwaku et al., 1988; Harvey et al., 2000). In this work (Lok and Massey, 2002), free-living stage adults were injected with plasmid DNA and some of the F1 embryos expressed a reporter gene encoded on the injected plasmid; however, these eggs did not hatch and no viable transgenic worms could be obtained. Parastrongyloides trichosuri is a natural parasite of Australian brushtail possums of the genus Trichosurus (Mackerras, 1959; Viggers et al., 1998) but can also be maintained in the Australian marsupial sugar glider, Petaurus breviceps (Nolan, personal communication), which can be obtained in many parts of the world as an exotic pet. It is a close relative of the Strongyloides spp. (Dorris et al., 2002) and like many Strongyloides spp., can undergo a free-living as well as a parasitic life cycle (Fig. 1(a)). The key difference is that P. trichosuri is the only parasitic nematode species known that can be maintained indefinitely in a free-living life cycle or induced to enter the parasitic life cycle by simple manipulation
of the L1 stage culture conditions (Grant et al., 2006). During the free-living phase, P. trichosuri is a dioecious nematode of similar size and biology to C. elegans. Although somewhat more delicate and less fecund than C. elegans hermaphrodites, free-living P. trichosuri females can survive microinjection. We report here, using P. trichosuri, what we believe is the first heritable genetic transformation of a mammalian nematode parasite, including successful passage of a transgenic worm through its mammalian host. The transgene was expressed during all P. trichosuri life cycle stages, including the parasitic phase in the host, and was also inherited through many generations. We have thereby developed a mammalian nematode parasite model in which it is possible to ‘knock-in’ genes to test their function, including genes expressed during the parasitic phase of the life cycle. Given that parasitic nematodes infect more than two thirds of the world’s human population and are a plague both on livestock and agricultural crops, the potential benefits to agricultural, veterinary and medical biotechnology of a genetically manipulable parasite as a research tool are large. 2. Materials and methods 2.1. Parasitology and free-living cultures Parasitological procedures and free-living cultures were essential as previously described (Grant et al., 2006). Briefly, adult female parasites for microinjection were obtained at necropsy from the small intestine of possums 3–6 weeks post transdermal infection with 5000 infective stage L3s (iL3s). Adult female free-living worms were obtained from cultures,
Fig. 1. The life cycle and development of Parastrongyloides trichosuri. (a) The life cycle of P. trichosuri; (b) Nomarski photomicrograph of the ovary of a free-living adult (100!); and (c) higher magnification of b (200!) at the point at which the microinjection needle penetrates the ovary syncytium.
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established from eggs or freshly hatched L1s, on 1/10 NGM agar (standard NGM, Sulston and Hodgkin, 1988) containing 1/10th the concentration of peptone. Post-microinjection freeliving cultures were maintained on the same medium. Infective larvae were produced by permitting free-living cultures on 1/10 NGM to proceed through several generations without subculturing. 2.2. Transgenesis The construction of a reporter plasmid, pPt hsp1Pro:NLS:GFP/lacZ (pPt hspPro), which is derived from the vector pPD96.02 (Fire, 1992; Fire et al., 1990) consists of the promoter region from the Pt hsp-1 heat-shock gene driving expression of a green fluorescent protein (GFP)/b-galactosidase fusion protein reporter gene containing a nuclear localisation signal (NLS).The utility of this plasmid in producing transgenic C. elegans has been described previously (Newton-Howes et al., 2006). This plasmid was injected into the ovary syncytium of adult female P. trichosuri using the method described for microinjection of C. elegans (Mello et al., 1991; Mello and Fire, 1995). Two types of microinjection were carried out. First, adult parasitic females were removed from the small intestine of possums 3–6 weeks p.i., the females were separated from the males (by manipulation under a dissecting microscope) into warm (w35 8C) 0.9% saline and transferred to dry 1% w/v agarose pads (Mello et al., 1991; Mello and Fire, 1995). The ovary syncytium was microinjected with a solution containing pPt hspPro plasmid at 80–100 ng/ml in H2O (or in worm injection buffer) combined with an approximately equal concentration of Sau3A-digested P. trichosuri genomic DNA. Worms were injected singly in order to minimise the period spent on the agarose pads. Injected worms were rehydrated in situ with PBS and transferred to NCTC insect tissue culture medium (Sigma) in wells of 24 well plates (one to three worms per well) and maintained in a 5% CO2 atmosphere at 35–37 8C. Following 18–24 h incubation under those conditions, eggs and hatched L1 larvae produced by the adults were transferred to 1/10 NGM plates seeded with Escherichia coli HB101, supplemented with an extract of possum faeces (Grant et al., 2006) and cultured further at 20 8C. Subsequent maintenance of free-living worms was as previously described (Grant et al., 2006). Alternatively, microinjection was performed as described above into the ovary syncytium of adult free-living females, which had been cultured, on 1/10 NGM at 20 8C (Grant et al., 2006). The anatomy of the female reproductive system of P. trichosuri is analogous to that of C. elegans (White, 1988) and S. stercoralis (Lok and Massey, 2002). It consists of a centrally located vulva and uterus, with a symmetrical reflexed two-armed ovary that extends anteriorly and posteriorly. The distal portion of the gonad, past the reflex point, consists of a syncytium composed of nuclei destined to be packaged into ova and then fertilised in the proximal half. Transgene DNA was injected into the proximal portion of this syncytium (arrows, Fig. 1(b) and (c)). The injected free-living females
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were rehydrated in situ on the agarose pad as for parasitic adults and then transferred to a seeded 1/10 NGM plate (three to five worms per plate) with 5–10 male free-living worms of the same age. In either case, the F1 generation were free-living and were expanded through subculture to an F2 generation. Each F2 culture was, where possible, founded by a small (!10) population of F1 worms picked from a single injection plate. In general, only a single positive F2 transgenic culture was kept and maintained as a transgenic ‘line’ to increase the likelihood that each line was the product of a unique transgenesis event. To screen for transgenic worms, F2 or later generation cultures were split: half of the culture was transferred to fresh, seeded 1/10 NGM plates to establish F3 and later generations and then maintained as a transgenic line by subculture to fresh plates at each generation. The remaining F2 worms were stained for b-galactosidase activity (Fire, 1992) and/or subjected to single worm PCR to detect the expression and/or presence of the transgene, respectively. Reporter gene expression in pPt hspPro plasmid is under the control of the Pt hsp-1 heat-shock promoter. The Pt hsp-1 gene is most closely homologous to the C. elegans hsp-1 gene and, like the C. elegans hsp-1 gene, Pt hsp-1 is constitutively expressed and induced only moderately (4- to 8-fold) by heat-shock (Newton-Howes et al., 2006). Consequently, worms were not subjected to heat-shock prior to staining for b-galactosidase activity. Single worm PCR detection of the transgene utilised nested primers located in the coding region of the GFP and the C. elegans derived unc-54 gene 3 0 untranslated region of the reporter plasmid. Single worms were prepared for PCR by digestion in 25 ml of lysis buffer (60 mM Tris, 0.6 mM EDTA, 1.2% Tween) containing 2 mg/ml proteinase K for 14 h at 42 8C, followed by 1 h 95 8C and 1 min 4 8C. The PCRs were carried out in a 25 ml volume using 2 ml of a 1/10 dilution of the worm lysate as template and Platinum Taq polymerase (Invitrogen) with 1.6 mM [Mg2C], 30 cycles of amplification and an annealing temperature 55 8C with a 1 min extension time (at 72 8C). Primers for the first round reactions were AGTCGAATTCGGCCGCTGTCATCAGAG (Forward) and TCAGAAGCTTAAACAGTTATGTTTGGTATATTG (Reverse). Second round PCR was carried out using the same reaction conditions, with 2 ml of a 1/10 dilution of the first round reaction as template and primers TACATGCTCTTTCTCCCTGTGC (Forward) and GCATCGTGCTCATCAATACTTGT (Reverse). Note that lysates that did not yield a PCR product were interpreted as being derived from non-transgenic worms. To ensure that there was template DNA able to be amplified from all lysates and thus remove any false negative results, a PCR for P. trichosuri actin was included as a positive control for template quality for all lysates. Actin PCR was carried out using the same reaction conditions as for the transgene with the primers CATGTGGAAGGGCATATCCT (Forward) and GAATACGTTGCTGCCCTTGT (Reverse). In some cases, Southern blots of the PCR products were carried out using a digoxigenin-labeled PCR product (Boehringer) generated from
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the microinjected plasmid as per the manufacturer’s instructions. 2.3. Infection of possums with transgenic worms Three transgenic F2 cultures (independently derived from different injected free-living females) were permitted to continue through several generations without subculture. All three yielded infective larvae after 10–14 days at 20 8C and each was used to infect one possum (i.e. one line per possum) at w5000 larvae per possum (under PC2 level containment as required by regulation in New Zealand). All experiments were performed in accordance with the 1987 Animal Protection (Codes of Ethical Conduct) Regulations of NZ, with approval from the Animal Ethics Committee of Wallaceville Animal Research Centre. Prior to infection, 40 individual iL3s were removed and subjected to single worm PCR to estimate the proportion of transgenic worms in the infecting dose. The possums were monitored by faecal egg count from day 12 p.i. and faeces collected when a positive faecal egg count was established. Eggs were prepared from these faeces and freeliving cultures established on 1/10 NGM (Grant et al., submitted). Adult F1 free-living worms (96 for each transgenic line) were assayed by single worm PCR (males only) and b-galactosidase staining for the presence and expression of the transgene. Once PCR and b-galactosidase status for the freeliving F1 of each line had been established, the parasitic adults from each possum were collected at necropsy and assayed by PCR (96 worms for each animal) and b-galactosidase staining. 2.4. Inheritance of the transgene in free-living worms The transgenic ‘lines’ established using the methods described above were clearly derived from a founder population that consisted of the progeny from more than one injected female and contained transgenic and non-transgenic individuals. To test whether the transgene was inherited by 100% or some lesser fraction of the progeny of a given transgenic parent, we carried out single male–female matings where at least one parent was transgenic. Single late L4 or early adult stage free-living (presumably virgin) females and a single male from the same transgenic culture were transferred to a small (!5 mm) spot of E. coli HB101/fecal extract on a 1/10 NGM plate. The parents and a sample of eight progeny from each fertile single pair were genotyped by PCR. 3. Results 3.1. Expression of a transgene in free-living worms The incidence of transgenic progeny produced following pPt hspPro microinjection of free-living adult females was determined by chromogenic staining for b-galactosidase (Fire, 1992). Typically we pooled 3–5 microinjected females onto a plate containing 10–20 uninjected males and cultured for several days prior to staining. Not surprisingly, given the pooling of parental injected worms, we found that
the percentage of positively stained worms varied widely between different injection experiments. In general, the highest frequencies of transgenic progeny were obtained following the injections of young females that contained less than five mature eggs in their uteri. From 3 to 5 microinjected females, we usually found at least some positively staining progeny among the F1s (5 of 17 worms in Fig. 2(a)) and in rare cases we found that virtually all progeny express b-galactosidase (15 of 15 worms in Fig. 2(b)). The post-injection survival of injected females varied from 30% to as high as 80–90% when young, free-living females were used. The occurrence of transgenic progeny when the reporter plasmid, pPt hspPro, was microinjected into the gonad of parasitic adult females was also variable and generally not as high as from injection of free-living females. The postinjection survival of parasitic females was difficult to judge because they rarely survived for more than 48 h after removal from the host (whether injected or not) and ceased laying eggs after 18–24 h. Varying the time post-infection at which the parasitic females were removed from the host did not appear to improve the frequency of transgenic progeny from, and the survival and fertility of, the parasitic females following injection (not shown). Expression of the GFP/b-galactosidase fusion protein reporter gene in pPt hspPro was driven by the Pt hsp-1 heatshock promoter region (Newton-Howes et al., 2006). The Pt hsp-1 gene is the likely orthologue of the C. elegans hsp-1 gene. Like the C. elegans hsp-1 gene, Pt hsp-1 is constitutively expressed and is induced only moderately (4- to 8-fold) to heatshock. When pPt hspPro was expressed in transgenic C. elegans, b-galactosidase activity was detected in gut cell nuclei in all developmental stages (Newton-Howes et al., 2006). The b-galactosidase staining in free-living P. trichosuri transformed by pPt hspPro is clearly localised to gastrointestinal cells with an expression pattern very similar to that previously seen with this plasmid in transgenic C. elegans. Although the reporter contains a NLS, we did not always observe the clear nuclear localisation in P. trichosuri that was seen in C. elegans. Although in some cases (e.g. Fig. 2(c) and (d)) we did find that most or all of the staining localised to gut cell nuclei, the staining of the gut cells was often more diffuse (Fig. 2(e)). The cause of the variable nature of the nuclear localisation is not clear. 3.2. Inheritance of the transgene in free-living worms Transgenes which result from microinjection in C. elegans are generally maintained as genetically unstable extrachromosomal arrays composed of tandem copies of the injected DNA. The characteristic feature of this mode of inheritance is that ! 100% of the progeny of a transgenic parent inherit the transgene and that expression of the transgene is mosaic in each transgenic worm (Mello et al., 1991; Mello and Fire, 1995). Early observations with P. trichosuri transgenesis were that only a portion of microinjected worms’ progeny appeared transgenic and their expression patterns were variable and mosaic: for example, anything from one or two gut cells to all
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Fig. 2. b-Galactosidase staining of transgenic free-living worms. (a) and (b) Fields showing stained and unstained worms from two transgenesis experiments; (c) detail of nuclear staining in the gut cells of an adult free-living male, note the punctuate nuclear localisation of the b-galactosidase activity; (d) the same detail for a free-living adult female; (e) a free-living male from a different experiment with more diffuse localisation of b-galactosidase staining; and (f) shows transgene expression in an embryonated egg.
gut cells were stained in individual worms (Fig. 2(a)–(f)). To investigate this observation further, a series of cultures were established from single male and virgin female worms from a single transgenic line, which contained transgenic (b-galactosidase expressing) worms. Eight progeny from each fertile single male/female pair were tested for the presence of the transgenic GFP coding region by PCR, as were the two parents. This test was carried out for two independently obtained transgenic lines, which should carry different extrachromosomal arrays. As shown in Fig. 3, some but not all of the progeny inherited the transgene where at least one of the parents was transgenic: this is as expected for extrachromosomal inheritance. Note that in the examples shown, one of the transgenes consistently produced a closely spaced doublet, both bands of which contained sequences, which hybridised to the GFP probe (Fig. 3, lower panel). We interpret this to indicate that a small rearrangement may have occurred during the assembly of the presumed extrachromosomal array, although most transgenic P. trichosuri lines we examined did not show this rearrangement (e.g. Fig. 3, upper panel). Similar rearrangement of transgene sequences has been observed in C. elegans (Mello and Fire, 1995).
determine whether pPt hspPro was also expressed in the parasitic life cycle. Infective larvae from each of the three transgenic lines described above were used to infect one possum per line (after a sample of the infecting dose was reserved for genotyping by PCR). The parasite eggs passed in the faeces were collected and cultured to obtain free-living worms while adult parasites were recovered at necropsy. We
3.3. Expression and inheritance in parasitic worms The Pt hsp-1 gene from which the promoter for the pPt hspPro transgene plasmid was derived is constitutively expressed in all stages of the free-living and parasitic life cycles (Newton-Howes et al., 2006), so it was of interest to
Fig. 3. Inheritance of the transgene from single pair matings. Southern blots of PCR products, probed with digoxigenin-labelled green fluorescent protein. Upper panel: two single pair broods (1a and 1b), each of eight F1worms. Lower panel: three single pair broods (3a–c), each of F1 worms. The C’ve controls are PCR products amplified from plasmid pPt hspPro.
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W.N. Grant et al. / International Journal for Parasitology 36 (2006) 475–483 Table 1 Frequency of transgene in a one line of parasitic adult worms and their freeliving progency by PCR and by histochemical staining for b-galactosidase activity
Fig. 4. Expression and inheritance of the reporter in transgenic parasitic worms. (a) b-Galactosidase stain of the posterior of an adult female parasite; (b) mean proportions of transgenic worms by PCR in three transgenic lines, A, B and C. Solid black, iL3s; light grey, male parasitic adults; mid-grey, free-living F1 males. Bars are 1 SD from the mean.
observed b-galactosidase expression in infective larvae and in the gut of adult parasites, as shown in Fig. 4(a), although the staining in any given adult parasitic worm was less intense than in free-living worms and fewer gut cells stained. We also genotyped, by PCR, a sample of the iL3s that were used to initiate the infections, a sample of the adult male parasites recovered at necropsy and a sample of their first generation male free-living progeny cultured from the faeces of infected possums. The proportion of each population, for three independent lines, that were transgenic is shown in Fig. 4(b). No consistent pattern of transgene frequency was observed when the lines are compared, but for each of the lines: (i) transgenic worms were able to establish patent infections; and (ii) the transgene was transmitted efficiently from the parasitic adults to their free-living progeny. In one case at least (Fig. 4, transgenic worm line B) the frequency of worms positive by PCR for the presence of the transgene was greater in parasites compared to iL3s, indicating that, in this line, more transgenic than non-transgenic iL3s established as adults. These results clearly demonstrate that the transgene can remain stable in parasitic adults in the mammalian host and be successfully passed to the free-living progeny of those parasites. 3.4. Mosaic expression of the transgene During these studies, we noted that the frequency of transgenic animals observed in different cultures was generally smaller when the scoring was done by b-galactosidase expression (phenotypic assay) compared with PCR (genotypic assay). This suggested that expression of the transgene might
Detection method
Percentage parasitic adults transgenic
Percentae free-living progency transgenic
Single worm PCR b-galactosidace expression
20.4 (n=93) 1.5 (n=600)
16.9 (n=89) 2.6 (n=6000)
be suppressed or silenced in some proportion of the worms that had inherited it. To test this more directly, we compared the proportion of worms positive by PCR with those positive by b-galactosidase staining in adult parasites and their free-living progeny for one of the transgenic lines described in Section 3.3. These data (Table 1) showed that 5- to 10-fold more animals contained the transgene (based on PCR) than expressed b-galactosidase. This is consistent with silencing of the transgene in many parasitic worms and their progeny. The proportion of worms showing silencing varied between lines. For example, the worm population depicted in Fig. 2(a) and (b), which were from free-living worms that had not passed through a parasitic life cycle, showed many fewer silenced worms. It also raises the possibility that the mosaic pattern of expression (i.e. the observation that different numbers of intestinal cells express the transgene in different worms) may be due, at least in part, to silencing as well as mitotic loss of the transgene in somatic cells. 4. Discussion Previous reports in several species of parasitic nematodes of vertebrates have demonstrated that it is possible to obtain transient expression of a transgene in the worm to which it was delivered (Davis et al., 1999; Jackstadt et al., 1999; Higazi et al., 2002; Lok and Massey, 2002) or to primary cell cultures derived from worms (Higazi et al., 2004). Lok and Massey (2002) were able to show that the transgene could also be transiently expressed in early embryos in eggs produced by microinjected free-living females of S. stercoralis but that these embryos invariably arrested development at an early stage. We have shown here for what we believe is the first time, using the model nematode parasite P. trichosuri, that it is possible to generate transgenic worms that express a transgene and then to establish and maintain these as transgenic worm lines. Importantly, we have also shown that the transgene can be heritably passed through either free-living or parasitic life cycles and it should therefore, be possible to maintain the transgene indefinitely. The ability to maintain and express transgenes through the parasitic life cycle is of particular importance, since it now opens the possibility to modulate the expression of parasite genes in the host. This capability makes possible the introduction of new functional genes or the modulation of existing genes to test hypotheses that involve the role of genes in host/parasite interactions.
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We have obtained transgenic progeny following microinjection of either parasitic or free-living adult females. Microinjection of free-living females is technically more challenging, as they appear to be significantly more sensitive to damage during the injection protocol than do C. elegans hermaphrodites. We were unable to accurately assess the tolerance of adult parasitic females to microinjection because they do not survive O36–48 h in tissue culture even when not injected but we were able to recover progeny from injected parasites, suggesting that they at least tolerate injection to some extent. For free-living females, the age of the worms at the time of injection was an important parameter: young adults with less than two or three eggs in utero generally survived more often and were more fertile following microinjection than were older worms. This is perhaps a reflection of the brief reproductive period of free-living females and their short lifespan (Grant et al., 2006). We have varied the p.i. age of parasitic females used for microinjection from young (shortly after the appearance of eggs in the hosts’ faeces) to several weeks after the onset of patency and have not observed any consistent change in the frequency of transgenic progeny. The observation that we obtain incomplete transmission of the transgene and that the transgenes display a mosaic expression pattern suggests that the transgene is inherited as an extrachromosomal array. If this is the case, then incomplete transmission of the transgene is due to the loss of array in the germline during meiosis. The only alternative to extrachromosomal inheritance (i.e. chromosomal inheritance) requires that the transgene is integrated into a chromosomal site and this is difficult to reconcile with the incomplete inheritance we observed. Transgenes generated by gonad microinjection of C. elegans are generally inherited as extrachromosomal arrays (Mello and Fire, 1995; Praitis et al., 2001) and the observations reported here in transgenic P. trichosuri suggest that extrachromosomal arrays may be a general feature of how nematodes handle transgenes. We have also observed that expression of the transgene is mosaic, i.e. that the expression pattern differs between worms, such that from one to all intestinal cells exhibit b-galactosidase expression. Mosaic expression is also a feature of extrachromosomal maintenance of transgenes in C. elegans, where the silencing is due to a combination of somatic loss of the array and specific down regulation of expression due to the repetitive nature of the array (Hsieh et al., 1999). We have not yet tested whether those adults in which the transgene is silenced are able to give rise to progeny in which transgene expression is reactivated, although this seems unlikely. Nor have we quantitatively compared silencing in different, independently derived, transgenes in different transgenic lines, although our qualitative observation was that there was considerable variation between lines in the degree of silencing observed. The inclusion of Sau3A restriction enzyme digested genomic DNA in the microinjection mix was observed to improve significantly the intensity of transgene expression but it did not entirely prevent somatic silencing in worm progeny (data not shown). Similar observations have been made regarding the silencing of transgenes in the germline of C. elegans, which is also
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partially relieved by the inclusion of irrelevant DNA in the injection mix (Kelly et al., 1997). Taken together, these observations favour the hypothesis that the incomplete inheritance and mosaic expression of the transgene are due to extrachromosomal inheritance. Expression of the transgene was driven by the Pt hsp-1 promoter and we observed expression of the b-galactosidase reporter only in gut cells of transgenic P. trichosuri. This is the same pattern of expression we found with this plasmid construct when introduced into transgenic C. elegans (Newton-Howes et al., 2006). Similar gut-specific expression was also seen in P. trichosuri when the C. elegans p212GATA promoter (Britton et al., 1998) replaced the Pt hsp-1 promoter (data not shown). These results show that, in these cases, promoter function is essentially the same in both species. The reporter gene, which we introduced into worms, is a translational fusion between the bacterial lacZ gene encoding b-galactosidase and GFP (Fire et al., 1990). Surprisingly, we have consistently failed to observe GFP expression even though b-galactosidase activity shows that the reporter gene is clearly transcribed and translated. Adult free-living P. trichosuri display strong gut autofluorescence of similar wavelength to GFP, so it is possible that GFP fluorescence is masked in adult worms. However, this does not explain all of our results since b-galactosidase activity, but not GFP fluorescence, was also easily detectable in earlier worm stages (including eggs), which display little or no autofluorescence. We have also observed b-galactosidase activity in the absence of GFP fluorescence in transgenic free-living P. trichosuri from the same GFP/b-galactosidase reporter gene driven by the C. elegans let-858 promoter (not shown), suggesting that the failure of P. trichosuri to express GFP fluorescence may be a general phenomenon. We believe the data we report here comprise the first evidence for transgene inheritance and thus for heritable germline transformation in a parasitic nematode of a mammal. The potential utility of this system as a functional genomic tool with which to investigate P. trichosuri is significant. This research avenue will be facilitated by the recent release of w8000 P. trichosuri ESTs and many additional ESTs from the closely related parasites Strongyloides ratti (Thompson et al., 2005) and S. stercoralis (Mitreva et al., 2004b). The technology reported in this paper also makes possible the expression of RNA containing hairpin loops (Tavernarakis et al., 2000; Johnson et al., 2005) in transgenic P. trichosuri, which should elicit RNAi phenotypes. The hairpin RNA could be put under the control of stage-specific or cell-specific promoters if desired or promoters that can be induced by exogenous agents (e.g. tetracycline) to permit regulated inhibition of specific genes that are expressed any stage including parasitic stages within the host. It should also be possible to ‘knock-in’ foreign genes under the control of selected promoters. For example, one could express secretory host immune modulators or fertility factors during the parasitic stages and measure their influence on host immune responses or fecundity. The capability to produce transgenic P. trichosuri adds to the already powerful attributes of this nematode
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