Transinfection: a method to investigate Wolbachia-host interactions and control arthropod-borne disease

Abstract

The bacterial endosymbiont Wolbachia manipulates arthropod host biology in numerous ways including sex ratio distortion and differential offspring survival. These bacteria infect a vast array of arthropods, some of which pose serious agricultural and human health threats. Wolbachia-mediated phenotypes such as cytoplasmic incompatibility and/or pathogen interference can be utilized for vector and disease control. However, many medically important vectors and important agricultural species are uninfected or are infected with strains of Wolbachia that do not elicit phenotypes desirable for disease or pest control. The ability to transfer strains of Wolbachia into new hosts (transinfection) can create novel Wolbachia-host associations. Transinfection has two primary benefits. First, Wolbachia-host interactions can be examined to tease apart the influence of the host and bacteria on phenotypes. Secondly, desirable phenotypes induced by Wolbachia in a particular insect can be transferred to another recipient host. This can allow for manipulation of insect populations that transmit pathogens or detrimentally affect agriculture. As such, transinfection is a valuable tool to explore Wolbachia biology and control arthropod-borne disease. This review summarizes what is currently known about Wolbachia transinfection methods and applications. We also provide a comprehensive list of published successful and unsuccessful Wolbachia transinfection attempts.

Introduction

Wolbachia are obligate intracellular bacteria that infect arthropods and nematodes. Within arthropods, Wolbachia infect a wide spectrum of insects, and the bacteria’s success has been attributed to their ability to manipulate host reproduction to favor it’s own maternal transmission. It is evident that over evolutionary time, horizontal transfer between species has occurred. Incongruent phylogenies of the host and the bacteria suggest that these horizontal transfers are commonplace (O’Neill et al. 1992; Vavre et al. 1999; Werren et al. 1995). The permissive nature of Wolbachia to transfer horizontally between individuals has been exploited in the laboratory to artificially infect new insect species.

Wolbachia strains have been artificially transferred, both interspecifically and intraspecifically, in many arthropod species. Recipient hosts include naturally infected species that have previously had their Wolbachia infection cleared, infected species that consequentially became superinfected with a new strain of Wolbachia, and species that were naturally uninfected. Artificial transfer of symbionts is not limited to Wolbachia, with other examples including Sodalis symbionts in tsetse fly (Weiss et al. 2006), diverse symbionts of aphids (Russell and Moran 2005; Tsuchida et al. 2005) and Spiroplasma in flies and beetles (Hutchence et al. 2011; Tinsley and Majerus 2007). However, in terms of numbers of studies and diversity of artificial interactions created, transfer of Wolbachia is by far the most developed (see Table 1). This review covers the current knowledge of transfer of Wolbachia between hosts, evaluates different transfer techniques, explores the biology of novel associations and discusses prospects for transinfecting new insect species.

Table 1.

Wolbachia transinfection attempts in arthropods.

Order/class	Recipient species	donor Wolbachia λ	Microinjection technique and Wolbachia source	Transfer result	Phenotype in recipient host †	Reference
Diptera, Drosophilidae	D. melanogaster	wRi	Embryonic, egg cytoplasm	Stable - G9	CI	(Boyle et al. 1993)
	D. simulans	wRi	Embryonic, egg cytoplasm	Stable - G4	CI
	D. simulans	wMel	Embryonic, egg cytoplasm	Stable - G4	CI
	D. simulans	wAlbA, wAlbB	Embryonic homogenized eggs	Stable -G6	CI	(Braig et al. 1994)
	D. simulans	wRi	Embryonic, egg cytoplasm	Stable - G5	CI	(Rousset and de Stordeur 1994)
	D. simulans	wHA	Embryonic, egg cytoplasm	Stable - G5	CI
	D. simulans∞	wHa	Embryonic, egg cytoplasm	90% at G12	CI	(Sinkins et al. 1995)
	D. simulans	wRi	Embryonic, egg cytoplasm	Stable - G11	CI	(Giordano et al. 1995)
	D. simulans	wMau	Embryonic, egg cytoplasm	Stable - G11	none
	D. mauritiana	wRi	Embryonic, egg cytoplasm	Stable - G11	CI
	D. serrata	wRi	Embryonic, egg cytoplasm	Stable - G45	CI	(Clancy and Hoffmann 1997)
	D. simulans	wMel	Embryonic, egg cytoplasm	Stable -10 years φ	CI	(Poinsot et al. 1998)
	D. simulans¶	wRi, wHa	Embryonic, egg cytoplasm	Stable	CI	(Rousset et al. 1999)
	D. simulans	wUni	Embryonic, egg cytoplasm	Lost after G7	T → none	(Van Meer and Stouthamer 1999)
	D. simulans	wHa	Embryonic, egg cytoplasm	Stable - G3	CI	(Poinsot and Mercot 2001)
	D. simulans	wNo	Embryonic, egg cytoplasm	Stable - G40	CI
	D. simulans	wHa, wNo	Embryonic, egg cytoplasm	Stable - G40	CI
	D. simulans	wHa	Embryonic, egg cytoplasm	Stable - G40	CI
	D. simulans	wNo	Embryonic, egg cytoplasm	Stable - G40	CI
	D. simulans	wHa, wNo	Embryonic, egg cytoplasm	Stable - G40	CI
	D. simulans	wMelPop	Embryonic, egg cytoplasm	Stable - G20	CI	(McGraw et al. 2001)
	D. simulans	wMelPop	Embryonic, egg cytoplasm	Stable - G40	CI	(McGraw et al. 2002)
	D. yakuba	wRi	Embryonic, egg cytoplasm	Stable >G200	CI	(Zabalou et al. 2004a)
	D. teissieri	wRi	Embryonic, egg cytoplasm	Stable >G100	CI
	D. santomea	wRi	Embryonic, egg cytoplasm	Stable >G70	CI
	D. simulans	wCer1, wCer2 β	Embryonic, egg cytoplasm	Stable – G20	CI	(Riegler et al. 2004)
	D. simulans	wRi	Embryonic, egg cytoplasm/ homogenized eggs	Stable - G19	CI	(Xi and Dobson 2005)
	D. melanogaster	wMel	Adult, homogenized tissue	Stable - G29	NT	(Frydman et al. 2006)
	D. simulans	wHa	Embryonic, egg cytoplasm	Stable - G10	CI NT	(Dean 2006)
	D. simulans	wHa	Embryonic, egg cytoplasm	Stable - G10	NA, NT
	D. simulans	wHa	Embryonic, egg cytoplasm	Stable - G10	N/A, NT
	D. simulans	wYak	Embryonic, egg cytoplasm	Stable >G200	None → CI	(Zabalou et al. 2008)
	D. simulans	wTei	Embryonic, egg cytoplasm	Stable >G200	None → CI
	D. simulans	wSan	Embryonic, egg cytoplasm	Stable >G200	None → CI
	D. melanogaster	wMelPop	Embryonic, cell culture	Stable - G46	CI	(McMeniman et al. 2008)
Diptera, Tephritidae	C. capitata	wCer1, wCer2 β	Embryonic, egg cytoplasm	Stable - G26	CI	(Zabalou et al. 2004b)
	C. capitata	wCer4	Embryonic, egg cytoplasm	Stable - G26	CI
	C. capitata	wCer2	Embryonic, egg cytoplasm	Stable - G60	CI	(Zabalou et al. 2009)
	B. oleae	wCer2	Embryonic, egg cytoplasm	Stable - G8	CI	(Apostolaki et al. 2011)
Diptera, Culicidae	Ae. albopictus	wAlbA, wAlbB α	Embryonic, egg cytoplasm	Stable - G8	CI	(Xi et al. 2005a)
	Ae. aegypti	wAlbA, wAlbB α	Embryonic, egg cytoplasm	Stable - G13	CI ψ	(Xi et al. 2005b)
	Ae. aegypti	wAlbA, wAlbB	Adult, homogenized tissue	40% at G12	Partial CI	(Ruang-Areerate and Kittayapong 2006)
	Ae. albopictus	wRi	Embryonic, egg cytoplasm	90% at G11	CI	(Xi et al. 2006)
	C. pipiens	wPipMol π	Embryonic, egg cytoplasm	Stable - G11	CI	(Walker et al. 2009)
	Ae. aegypti	wMelPop	Embryonic, egg cytoplasm	Stable - G17	CI ψ	(McMeniman et al. 2009)
	Ae. albopictus	wMelPop	Embryonic, egg cytoplasm	Stable - G10	LS ††	(Suh et al. 2009)
	Ae. albopictus	wPip	Embryonic, egg cytoplasm	Stable - G11	CI	(Calvitti et al. 2010)
	Ae. albopictus	wRi	Embryonic, egg cytoplasm	Stable - G13	CI	(Fu et al. 2010)
	Ae. aegypti	wMel	Embryonic, cell culture	Stable - G8	CI ψ	(Walker et al. 2011)
	Ae. albopictus	wMel	Embryonic, egg cytoplasm	Stable - G8	CI	(Blagrove et al. 2011)
	An. gambiae	wMelPop,	Adult, cell culture	Infection in G0	BMM ψ	(Hughes et al. 2011a)
	An. gambiae	wAlbB	Adult, cell culture	Infection in G0	None ψ
	Ae. polynesiensis	wAlbB	Embryonic, egg cytoplasm	Stable - G8	CI	(Andrews et al. 2012)
	Ae. albopictus	wAlbA	Embryonic, egg cytoplasm	Stable - G8	CI	(Calvitti et al. 2012)
		wAlbB	Embryonic, egg cytoplasm	Stable - G8	CI
	An. stephensi	wAlbB	Embryonic, egg cytoplasm	Stable - G34	CI	(Bian et al. 2013)
Lepidoptera	E. kuehniella	wKue	Embryonic, egg cytoplasm	Stable - G3	CI	(Sasaki and Ishikawa 2000)
	E. kuehniella	wSca	Embryonic, egg cytoplasm	Stable - G21	F → MK	(Fujii et al. 2001)
	E. kuehniella	wCauA, wCauB	Embryonic, egg cytoplasm	Stable - G15	CI →many δ	(Sasaki et al. 2002)
	O. scapulalis	wKue	Embryonic, egg cytoplasm	Stable - G7	MK → CI	(Sakamoto et al. 2005)
	B. mori	wHecFem	Prepupae and pupae, cell culture	Infection in G0		(Kageyama et al. 2008)
	B. mori	wHecCI	Prepupae and pupae, cell culture	Infection in G0
Hymenoptera	T. dendrolimi	T. pretiosum	Prepupae and pupae, insect	Stable - G26 ε		(Grenier et al. 1998)
	T. kaykai	wKay	Co-rearing, live insects	Infection in G0	PI	(Huigens et al. 2004)
	T. deion	wDei	Co-rearing, live insects	Infection in G0	PI
	Tr. atopovirilia	T. atopovirilia	Co-rearing, live insects	No infection
	T. deion	wKay	Co-rearing, live insects	Infection at G0
	T. kaykai	wDei	Co-rearing, live insects	Lost at G3	PI
	T. atopovirilia	T. pretiosum	Co-rearing, live insects	No infection
	T. pretiosum	T. atopovirilia	Co-rearing, live insects	No infection
	T. kaykai	wKay	Pupae, cell culture	Transmission to G1		(Kubota et al. 2005)
Hemiptera	L. striatellus	wRi χ	Adult, hemolymph	30% at F12	CI	(Kang et al. 2003)
	L. striatellus	wMel χ	Adult, hemolymph	Lost at G1
	L. striatellus	wHa χ	Adult, hemolymph	Lost at G1
	L. striatellus	wNo χ	Adult, hemolymph	Lost at G1
	N. lugens	wStri	Nymphal, cell culture	<10% at G40–60	CI	(Kawai et al. 2009)
Coleoptera	T. confusum	Tribollium sp.	Embryonic, egg cytoplasm	Stable - G2	CI	(Chang and Wade 1994)
	T. confusum	Tribollium sp.	Embryonic, egg cytoplasm	Stable - G2	NA	(Chang and Wade 1996)
Isopoda	A. nasatum	wVul	Adult, homogenized ovary	Infected G1 females	F	(Rigaud and Juchault 1995)
	A. vulgare	wElo	Adult, homogenized ovary	Somatic infection in G0
	A. vulgare	wVul	hemolymph transfer, hemolymph	Infection in G0	F
	Isopods §	Isopod Wolbachia *	Adult, homogenized tissue	Infected ~ 400 dpi	F	(Bouchon et al. 1998)
	O. asellus	wAse	Adult, homogenized ovary	Stable - G3	FBB	(Rigaud et al. 1999)
	A. vulgare	wCc	Adult, homogenized tissue	Infection in G0	CI	(Moret et al. 2001)
	P. scaber	wVul	Adult, homogenized ovary	No infection (N=19)		(Rigaud 2001)
	P. scaber	wNas	Adult, homogenized ovary t	9% at G1	None
	O. asellus	wVul	Adult, homogenized ovary	7% at G1	None
	O. asellus	wNas	Adult, homogenized ovary	No infection (N=12)
	A. vulgare	wVul	Adult, homogenized ovary	95% at G1	F
	A. vulgare	wNas	Adult, homogenized ovary	88% at G1	F
	A. nasatum	wVul	Adult, homogenized ovary	84% at G1	F
	A. nasatum	wNas	Adult, homogenized ovary	95% at G1	F
	A. vulgare	wVulC	Adult, homogenized ovaries	Stable - G2	NT	(Braquart-Varnier et al. 2008)

Abbreviations: NT- not tested, CI – cytoplasmic incompatibility, F – feminization, PI – parthenogenesis inducing, MK – male killing, FBB – female bias broods, BMM – blood meal mortality, dpi – days post injection;

^λTwo Wolbachia strains listed indicates both strains were microinjected, if no Wolbachia strain is listed, the arthropod host is named;

^†affect of transinfection of Wolbachia on reproduction in recipient host

^*Five Wolbachia strains from Armadillidium nasatum, Armadillidium vulgare, Cylisticus convexus, Chaetophiloscia elongata, Porcellionides pruinosus were used;

^§Up to ten isopods recipients: Armadillidium nasatuma, Armadillo officinalisa, Porcellio scaber, Porcellio gallicus Porcellio dilatatus dilatatusa, Porcellio dilatatus petit, i Porcellionides pruinosus, Cylisticus convexus;

^¶recipient host was infected with wHa and wNo;

^∞recipient host was infected with wRi,

^πrecipient host was infected with wPipCol;

^αonly wAlbB infected the host;

^βonly wCer2 infected the host;

^χrecipient host was infected with wStri;

^εdetails about infection from (Pintureau et al. 2000);

^φdetails about infection from (Zabalou et al. 2008);

^δwCauA - MK, wCauB - CI, wCauA and wCauB – MK.

Arrow indicates a change in phenotype from natural (left of arrow) to novel (right of arrow) host;

^††high embryonic mortality in transinfected line masked potential CI effects;

^ψinsect also display pathogen interference.

What is transinfection?

Transinfection is the mechanical transfer of symbionts into a novel host. As opposed to introgression, whereby a Wolbachia-infected line is repeatedly backcrossed with males to homogenize the host background, transinfection allows the creation of diverse novel Wolbachia-host associations and is not constricted by species mating barriers. Two procedures have been successfully used to create arthropods transinfected with Wolbachia: embryo microinjection and adult microinjection. By far the most widespread technique used to develop transinfected lines is embryo microinjection, whereby Wolbachia are injected into the posterior pole of pre-blastoderm embryos using a fine needle and micromanipulator. Embryos are left to develop to adulthood, and the subsequent progeny are screened to determine if germline infection and transmission occurred. As the name suggests, adult microinjection differs in that the recipient is at the adult stage rather than embryo. For this technique to be successful, Wolbachia must traverse through tissues and cross membranes to reach the germline where it can then be transmitted to the next generation. In some instances, Wolbachia has been injected into the pupal or pre-pupal life-stages (Grenier et al. 1998; Kageyama et al. 2008; Kubota et al. 2005), however similar constraints apply in that ultimately, Wolbachia need to reach the germline. Other techniques such as co-rearing of the recipient and donor species (Huigens et al. 2004) and hemolymph transfer (Rigaud and Juchault 1995) have achieved transfer of Wolbachia to a new host to some degree but these approaches are only suitable for a limited number of arthropods and are generally not considered viable or efficient options for transinfection in many insect species.

Comparison of embryonic and adult microinjection

For the majority of cases where the successful establishment of stable transinfected lines has occurred, the embryonic microinjection technique was employed, however both this technique and microinjection of other life stages have their advantages and disadvantages. Embryonic microinjection localizes Wolbachia directly within the developing embryo, compared to adult microinjection where Wolbachia are intrathoracically injected. This direct access to the pole cells means that cells that differentiate into germline or soma are already infected, rather than Wolbachia needing to gain entry to the germline when injected into more developed life stages. Wolbachia have a natural propensity to infect the germline, and in adult Drosophila melanogaster, intrathoracically injected Wolbachia are seen to localize in the stem cell niche within the germline (Frydman et al. 2006). As the soma can be infected, embryonic microinjection can often lead to somatic infection in the transinfected line, which can be important for Wolbachia phenotypes such as pathogen interference (Bian et al. 2010; Moreira et al. 2009; Walker et al. 2011).

When Wolbachia are microinjected into adult insects the bacteria must evade the insect immune response. With naturally occurring associations, Wolbachia seem to neither elicit nor suppress the immune response (Bourtzis et al. 2000; Xi et al. 2008), but when analyzing artificial associations, Wolbachia have been shown to affect immunity (Bian et al. 2010; Hughes et al. 2011b; Kambris et al. 2009; Moreira et al. 2009). After injection of Wolbachia into an arthropod host, there is evidence that the bacteria may manipulate host immunity, perhaps to enhance their own proliferation (Braquart-Varnier et al. 2008; Hughes et al. 2011a). In crustaceans, Wolbachia is present within hemocytes and in hematopoietic organs, and these have been suggested to act as a reservoir to discharge the bacteria into the hemolymph (Chevalier et al. 2011). But whether hemocytes can act as a source of Wolbachia in insects is still to be resolved. Microinjected Anopheles mosquitoes were also seen to contain Wolbachia within circulating hemocytes, however it is uncertain if the bacteria were viable (Hughes et al. 2011a). To overcome diminishing Wolbachia levels due to the host immune response, more brute force approaches can be adopted by injecting more bacteria, a luxury that can be more easily afforded in adult microinjection. Alternatively, host immunity can be suppressed in adults using molecular techniques such as RNA interference (RNAi).

Innate differences between adult and embryo immunity may aid Wolbachia establishment in embryos. In Drosophila melanogaster early embryogenesis, hemocytes are restricted to a region within the head mesoderm and have highly divergent roles in development (Holz et al. 2003; Wood and Jacinto 2007). Another study using Drosophila embryos showed that specialist insect pathogens employ methods to avoid phagocytosis, whereas Escherichia coli were engulfed (Vlisidou et al. 2009). Given the evidence that Wolbachia can avoid or manipulate host immunity, it could be predicted that similar mechanisms are employed by the bacteria to avoid detection in injected embryos. While little is known regarding the interaction of Wolbachia with the host immune system in developing embryos, it is feasible that the host may treat microinjected Wolbachia in a similar fashion to maternally transmitted bacteria, however further work is required to examine Wolbachia-host interaction in embryos to validate this theory.

Another difference between the transinfection techniques is the number of viable G₀ (injected generation) females produced, presumably due to the sensitivity of each life stage to injection. Using adult microinjection, a high number of G₀ can be attained, compared to embryo microinjection where only a small proportion of reproducing adults will arise from injected embryos. These differences can either be viewed in a positive or negative light, and preference for a particular technique may lie where resources allocation is preferred. A larger number of G₀ obtained from adult injection means more time needs to be spent on screening progeny for successful transfers, while the lower number of G₀ from embryonic microinjection means greater effort needs to be allocated in the injection process. For either technique, the progeny of the G₀ need to be screened, as both can lead to somatic infection (which is not necessarily indicative of germline infection). As Wolbachia are maternally inherited, infected males are a dead-end for the bacteria. Obviously for adult injections, females will be injected, but prior to embryo microinjection sex is unknown, therefore approximately half of surviving G₀ insects are of no use for establishment of infected lines. Lastly, embryonic microinjection is a highly specialized technique that requires a substantial initial outlay of funds for equipment and trained personnel, whereas adult microinjection is less technical and cheaper to undertake.

Source of donor Wolbachia

There are several sources where Wolbachia can be obtained for microinjection. When using the embryo microinjection technique, Wolbachia is predominantly extracted from the egg cytoplasm of an infected species then transferred to the recipient species. Wolbachia can be extracted from either the anterior or posterior regions of the donor egg (Xi and Dobson 2005), while homogenized ovaries or eggs can also be used as a source of Wolbachia. Alternatively Wolbachia can be extracted from cell culture systems. Cell culture is advantageous in that a high density of homogeneous bacteria can be obtained for microinjection and cell lines can be used to adapt strains of bacteria to the desired host background. The adaptation of the wMelPop strain of Wolbachia by serial passage in a RML-12 Aedes aegypti cell line was seen as critical for successful transinfection of this strain into the mosquito, Ae. aegypti (McMeniman et al. 2009). After transfer of Aedes–adapted wMelPop back into the native D. melanogaster host, phenotypic shifts were interpreted as genetic adaptation of Wolbachia to the mosquito intracellular environment (McMeniman et al. 2008), however it is unclear if these effects were specifically due to adaptation in Aedes cells or changes to the Wolbachia phenotype resulting from serial passage in cell lines. In contrast, the wAlbB strain was transferred from Ae. albopictus to An. stephensi without adaptation in cell lines (Bian et al. 2013). Although more laborious, adaptation of Wolbachia to the novel host could also be achieved by subsequent microinjection into the recipient species followed by extraction after Wolbachia has replicated within the insect (Frydman 2006). However, this may preferentially select for Wolbachia capable of infecting somatic tissue. The most optimal adaptation system would be a cell line derived from the germline of the recipient species or genus.

Phenotype modification upon transinfection

Phenotypic shifts are common upon transfer of Wolbachia to a novel host, probably due to the maladaptation of the new association. Theory predicts that novel host-parasite relationships are likely to be maladapted, compared to old associations that are more mutualistic (Levin 1996; Turelli 1994). There are several examples where transinfected insects, representing a novel association, display fitness costs. Transfer of wVul from Armadillidium vulgare to Porcellio dilatatus males was lethal (Bouchon et al. 1998), while wMelPop from D. melanogaster had pathogenic effects in Ae. albopictus (Suh et al. 2009). Anopheles stephensi mosquitoes infected with wAlbB have severe reductions in fecundity (Bian et al. 2013). Furthermore, fitness costs were incurred in Drosophila species transinfected with Wolbachia (Clancy and Hoffmann 1997; McGraw et al. 2002). However, in a relatively short time period, shifts from pathogenic to mutualistic associations can occur, demonstrated in both transinfected (McGraw et al. 2002) and natural infections (Weeks et al. 2007).

Selection of transinfected lines

Due to the maladaptation of the novel Wolbachia-host associations, the infection frequency in early generations can fluctuate stochastically. This may be due to incomplete cytoplasmic incompatibility (CI; a form of reproductive manipulation induced by Wolbachia on the host), inefficient vertical transmission and/or pathogenic association between the Wolbachia and host. Therefore, selection plays a critical role in establishing stable transinfected lines. In transinfected D. melanogaster lines, an initial selection period saw infection frequencies rise before a period where relaxed selection led to a drop in infection frequency (McMeniman et al. 2008). A second selection period after approximately G₁₅ led to 100% infection in subsequent generations (McMeniman et al. 2008). After transinfection of a life-shortening strain of Wolbachia from D. melanogaster to Ae. albopictus, selection and outcrossing to wild type males was crucial in preventing the loss of infection (Suh et al. 2009). In contrast, Xi and Dobson (2005) suggest that after extensive screening at G₀ and G₁ stages, it was unusual to lose an infection from a Drosophila transinfected line. However, this may be the case in intraspecific transfers, particularly into Drosophila backgrounds, which are likely to be more amenable to infection. As associations tend to shift towards mutualism over time, selection in the initial phase after transinfection may avoid loss of infection due to instability in infection frequencies and temporary fitness costs due to novel interactions between the host and bacteria.

Applications of transinfected arthropods

Transinfection can be used to examine novel associations and to disentangle the role of the bacteria or the host genotype on the type of reproductive alterations induced. Such examples are prominent within the order Lepidoptera. When the feminizing strain wSca from the moth Ostrinia scapulalis was transinfected into Ephestia kuehniella, it induced male killing, demonstrating the influence of host factors determining reproductive phenotype (Fujii et al. 2001). The reciprocal transfer of wKue, which induces CI in E. kuehniella, also induced CI when transferred into O. scapulalis, suggesting this phenotype is derived from the bacteria (Sakamoto et al. 2005). Transfer of wCauA from Cadra cautella induced male killing in E. kuehniella, while transfer of wCauB induced incomplete CI (Sasaki et al. 2002). Host suppression of Wolbachia-mediated reproductive manipulations and hidden Wolbachia phenotypes have been observed in other natural associations (Hornett et al. 2006; Hornett et al. 2008).

The Drosophila species complex harbor many strains of Wolbachia, with variable levels of CI (McGraw and O’Neill 1999; Merçot and Charlat 2004). The variability within these strains lies with the ability of the bacteria to induce and/or rescue CI, which has led to classification of strains based on their modification (mod) and rescue (resc) propensity (Werren and O’Neill 2004). Four possible variants are possible: mod+/resc+, mod+/resc−, mod−/resc+, and mod−/resc−. The strains wYak, wTei and wSan do not cause CI in their natural hosts (Drosophila yakuba, Drosophila teissieri, and Drosophila santomea, respectively), but when these strains were transferred into D. simulans, they all induced CI (Zabalou et al. 2008). Conversely, when the wRi strain from D. simulans was transferred into D. yakuba, D. teissieri, or D. santomea, this strain induced CI in all three species (Zabalou et al. 2004a). Unexpectedly, the natural infections (wYak, wTei and wSan) rescued the modification of wRi-transfected flies (Zabalou et al. 2004a). Interestingly, when the wTei strain was transfected into D. simulans, it was unable to rescue its own CI modification, suggesting that this association changes wTei into a “suicide strain” (mod+/resc−) (Zabalou et al. 2008). Taken together, these novel associations demonstrate the unique interactions between Wolbachia and the host that influence reproductive phenotypes.

Wolbachia has been suggested as an agent for vector and disease control given its ability to spread into insect populations and inhibit pathogens (for reviews see (Bourtzis 2008; Brownlie and Johnson 2009; Cook and McGraw 2010)). However, many important vector species are uninfected. These uninfected insects can be considered an open niche for Wolbachia modification. Transinfected disease vectors that transmit human or agricultural pathogens with a stable Wolbachia infection include, An. stephensi (Bian et al. 2013), Ae aegypti (McMeniman et al. 2009; Ruang-Areerate and Kittayapong 2006; Walker et al. 2011; Xi et al. 2005b), Ae. albopticus (Blagrove et al. 2011; Calvitti et al. 2010; Fu et al. 2010; Suh et al. 2009; Xi et al. 2006), Ae. polynesiensis (Andrews et al. 2012) Culex pipiens (Walker et al. 2009) Ceratitis capitata (Zabalou et al. 2009; Zabalou et al. 2004b), Nilaparvata lugens (Kawai et al. 2009), and Laodelphax striatellus (Kang et al. 2003). Some of these transinfected insects have been evaluated in semi-field and field conditions and are being implemented as control strategies (Hoffmann et al. 2011; Rasgon 2011; Walker et al. 2011), which is encouraging for the use of Wolbachia in a applied setting. Still, despite many transinfection attempts, Wolbachia infection remains illusive in some important vector species.

Insects recalcitrant to infection

Some hosts are seemingly impervious to Wolbachia infection. The reasons for refractoriness are unknown, but could relate to either bacterial or host factors. It is likely that the number of hosts recalcitrant to infection is under-estimated as predominantly transinfection attempts are made on either model organisms or insects of economical and medical importance. Furthermore (and unfortunately for the scientific community) unsuccessful transinfection attempts are rarely published. Despite Tribolium confusum being naturally infected and amenable to transinfection (Chang and Wade 1994; Chang and Wade 1996), hundreds of attempts to infect T. castaneum with Wolbachia from T. confusum have not yielded an infected line despite using the same methodology (pers com. M.J. Wade). Transinfection attempts by injecting Wolbachia into Bombyx mori larvae lead to somatic infection in G₀, but no stable infection (Kageyama et al. 2008).

For many years transinfection was attempted on mosquitoes within the Anopheles genus given their importance in human health. Due to many failed attempts, and the lack of natural infection in native Anophelines (Kittayapong et al. 2000; Rasgon and Scott 2004; Ricci et al. 2002), there were suggestions that this genus was impervious to Wolbachia infection. However infection of Anopheles cells (Rasgon et al. 2006) and somatic infection of adults (Hughes et al. 2011a; Hughes et al. 2012b; Jin et al. 2009; Kambris et al. 2010) indicated these mosquitoes are capable of harboring Wolbachia, suggesting the inability to create a stable line was due to other barriers. Recently, Bian and colleagues (Bian et al. 2013) developed a stable line using embryonic microinjection in the Indian malaria vector, Anopheles stephensi demonstrating Wolbachia can stably associate in this genus of mosquitoes. However, a stable infection in the major African malaria vector, An. gambiae, which is the main vector of Plasmodium parasites in sub-Saharan Africa, still remains elusive. Promisingly, the tissue distribution of Wolbachia and pathogen inhibitory effects between stably infected An. stephensi (Bian et al. 2013) and somatically infected An. gambiae (Hughes et al. 2011a; Hughes et al. 2012b; Kambris et al. 2010) are similar, suggesting stable infection in the African malaria mosquito is feasible.

While speculative, there are several strategies that could be implemented to achieve infection in this important mosquito species. Although more convenient, the use of laboratory mosquito lines may decrease genetic heterogeneity in the host background. Greater genotypic divergence in both the host and the bacteria may provide a wider base to select for compatible associations. Furthermore, pathogenic Wolbachia strains such as wMelPop, despite having desirable phenotypes for vector control, may not be best suited to infection. Recently, we developed an ex vivo germline assay to determine the infectivity of Wolbachia into ovaries (Hughes et al. 2012a). Wolbachia strains infected native hosts or close relatives at higher titers compared to more divergent hosts, with the wAlbB strain (from Aedes albopictus) more adapted to the Anopheles germline than Wolbachia from flies. This technique may be used to assess the suitability of bacterial strains in a particular host and to determine if adaptation to a new species is occurring. Specifically adapting a host to infection may be achieved by genetic modification of the insect, yielding the hosts more susceptible to Wolbachia. In Aedes mosquitoes, host factors induced by Wolbachia manipulation include the host microRNA profile which subsequently increased bacterial titre (Hussain et al. 2011). The development of new transgenic techniques that allows manipulation of host gene expression at early life stages provide tools that could be used to enhance embryonic microinjection in species where transinfection attempts have previously been unsuccessful (Peng et al. 2011). Further research that identifies barriers to horizontal transfer of Wolbachia between species and novel transinfection techniques would be advantageous to the scientific community and accelerate the creation of new Wolbachia-host associations.

Summary

Diverse Wolbachia-host combinations can be created through transinfection. This allows a tractable method to study Wolbachia-host interactions. Embryonic microinjection is predominantly used to create stable Wolbachia infected lines, however the number of examples that use other techniques is increasing. Novel transinfection techniques may be the key to transfer Wolbachia into hosts that have so far resisted infection. Further developments in this field may allow for the biological control of arthropod-borne diseases.