Next Article in Journal
Associations of Conifer-Infesting Bark Beetles and Fungi in Fennoscandia
Next Article in Special Issue
Generation of Nutrients and Detoxification: Possible Roles of Yeasts in Leaf-Cutting Ant Nests
Previous Article in Journal / Special Issue
Phylogenetic Analysis of Fusarium solani Associated with the Asian Longhorned Beetle, Anoplophora glabripennis

Insects 2012, 3(1), 161-199; doi:10.3390/insects3010161

Review
Insect Sex Determination Manipulated by Their Endosymbionts: Incidences, Mechanisms and Implications
Daisuke Kageyama *, Satoko Narita and Masaya Watanabe
Insect-Microbe Research Unit, National Institute of Agrobiological Sciences (NIAS), Owashi 1-2, Tsukuba, Ibaraki 305-8634, Japan; Email: naritas@affrc.go.jp (S.N.); wtnbm1ab@affrc.go.jp (M.W.)
*
Author to whom correspondence should be addressed; Email: kagymad@affrc.go.jp.
Received: 25 November 2011; in revised form: 14 January 2012 / Accepted: 2 February 2012 /
Published: 10 February 2012

Abstract

: The sex-determining systems of arthropods are surprisingly diverse. Some species have male or female heterogametic sex chromosomes while other species do not have sex chromosomes. Most species are diploids but some species, including wasps, ants, thrips and mites, are haplodiploids (n in males; 2n in females). Many of the sexual aberrations, such as sexual mosaics, sex-specific lethality and conversion of sexuality, can be explained by developmental defects including double fertilization of a binucleate egg, loss of a sex chromosome or perturbation of sex-determining gene expression, which occur accidentally or are induced by certain environmental conditions. However, recent studies have revealed that such sexual aberrations can be caused by various groups of vertically-transmitted endosymbiotic microbes such as bacteria of the genera Wolbachia, Rickettsia, Arsenophonus, Spiroplasma and Cardinium, as well as microsporidian protists. In this review, we first summarize the accumulated data on endosymbiont-induced sexual aberrations, and then discuss how such endosymbionts affect the developmental system of their hosts and what kinds of ecological and evolutionary effects these endosymbionts have on their host populations.
Keywords:
endosymbiont; feminization; insect; male killing; Wolbachia; sex determination

1. Introduction

In arthropods, the fundamental system of sex determination is generally considered to be genetic, and this is supported by the strict concordance in the dimorphism of sexual phenotypes and karyotypes (i.e., sex chromosomes) and the rare occurrence of gynandromorphs (chimeric individuals having tissues with male and female genotypes). However, the sex determination and differentiation of arthropods can be perturbed by the endosymbionts or parasites they harbor. The most striking effects on sexual phenotype are induced by microbes that are transmitted from mothers to offspring (often called maternally transmitted or cytoplasmic parasites). Since males cannot transmit cytoplasmic parasites to their offspring, parasites in the male cytoplasm are essentially dead from the evolutionary viewpoint. To make up for these shortcomings, some parasites adopt a variety of tactical strategies such as killing of males, converting males into females or inducing parthenogenesis.

In this article, we briefly overview the diversity of the sex-determining mechanisms in insects, summarize the incidences of maternally transmitted microbes that affect or may affect sex determination, and discuss the possible mechanism of such phenomena. We also discuss the ecological and evolutionary effects that such microbes potentially have on their host populations.

2. Diversity and Common Features of Insect Sex Determination

In the majority of insects, sex is strictly determined according to the genotype. For example, many of the dipteran insects (flies and mosquitoes) have a male-heterogametic sex chromosome constitution (i.e., XX: female; XY: male) while many of the lepidopteran insects (butterflies and moths) have a female-heterogametic chromosomal constitution (i.e., ZZ: male; ZW: female). On the other hand, many of the hymenopteran insects (ants, bees and wasps) do not have sex chromosomes. Instead, they have a haplodiploid sex-determination system, in which fertilized diploid (2n) eggs become females and unfertilized haploid (n) eggs develop into males [1,2,3]. The molecular mechanisms underlying sex determination and differentiation in the model insect Drosophila melanogaster (Diptera; Drosophilidae) are well understood. At a very early embryonic stage, each cell determines its sex independently, and once determined, the sex of each cell is maintained during later development through a gene expression cascade consisting of Sex-lethal (Sxl), transformer (tra), doublesex (dsx) and other genes, in which sex-specific mRNA splicing plays an important role [4,5,6]. In the honeybee Apis mellifera, the complementary sex determiner (csd) gene affects sex through allelic combination, whereas the feminizer (fem) gene induces sex-specific splicing, producing a functional protein only in females. Comparisons between the sex-determination pathways of A. mellifera, Ceratitis capitata and D. melanogaster suggest that the tra/dsx pathway is conserved among insects and is likely to be ancestral [7] (Figure 1). Sex determination in a cell-autonomous manner is also believed to be widespread among insects on the basis that sexually mosaic individuals often occur in a diverse array of insects [8].

Similar to insects, some of the non-insect arthropods are also considered to have genetically based sex determination. However, they differ from insects because their sexual differentiation is deeply affected by sex hormones that are secreted by particular organs (e.g., the androgenic gland in crustaceans) and circulate in the body together with hemolymph.

Insects 03 00161 g001 1024
Figure 1. The sex determination cascade in Drosophila melanogaster. PeSxl indicates Sxl transcript from the early promoter. PmSxl indicates Sxl from the late promoter. The gray box shows the conserved part of the cascade. (Adapted from [7].)

Click here to enlarge figure

Figure 1. The sex determination cascade in Drosophila melanogaster. PeSxl indicates Sxl transcript from the early promoter. PmSxl indicates Sxl from the late promoter. The gray box shows the conserved part of the cascade. (Adapted from [7].)
Insects 03 00161 g001 1024

3. Sexual Aberrations of Insects

In arthropods, sexually mosaic or gynandromorphic individuals are repeatedly observed in natural and laboratory populations of numerous species [9]. Most of them are considered to arise through accidental, very rare developmental defects such as double fertilization of a binucleate egg, loss of a sex chromosome or upregulation/downregulation of sex-determining genes.

On the other hand, some conditions can stably generate sexual aberrations in certain insects. In some mosquitoes of the genus Aedes, a high temperature condition can cause males to develop into morphological females. When the number of days of exposure to high temperature was lessened, various degrees of intersexes, i.e., individuals having both male and female characters, appeared [10,11,12]. Crosses between geographical races of the gypsy moth Lymantria dispar also resulted in the appearance of intersexes [13,14,15]. In some crossing combinations, genetically male offspring (ZZ) showed mosaic color patterns in their wings, which is clearly distinguishable between sectors of male color and female color. In other combinations, genetically female offspring (ZW) showed homogeneous in their wing color, which is intermediate between male and female [14]. Sexual mosaics were also found in crosses between tetraploid females and diploid males in the psychid moth Solenobia triquetrella [16]. The resulting triploid offspring with two Z chromosomes and three sets of autosomes developed into sexual intergrades owing to the intermediate ratio (2:3) between the Z chromosomes and autosome sets, which is neither typically female (1:2) nor typically male (1:1). Although it was used to be considered that all the aberrations of sexuality were attributed to environmental disturbances or stochastic effects, recent investigations revealed that some of them can be caused by endosymbionts, another organism living inside the body.

4. Involvement of Endosymbiotic Microorganisms

It has been increasingly recognized that various forms of sex aberrations in arthropods can be caused by symbiotic microorganisms. It was first recognized that Wolbachia pipientis (belonging to Alphaproteobacteria) exerts various types of manipulation on the reproductive system of arthropods [17,18,19]. Wolbachia-induced reproductive manipulation can be classified into four major categories: feminization, male killing, parthenogenesis induction and cytoplasmic incompatibility. By using these manipulations, Wolbachia can spread among host populations at the expense of host fitness. Following the succession of discoveries regarding Wolbachia-induced host manipulations, several other bacteria such as Cardinium, Spiroplasma and Arsenophonus, which belong to distantly related taxonomic groups, were also found to manipulate host reproduction in similar ways to Wolbachia (Table 1 and Table 2; Table S1).

Similar to other symbiotic bacteria such as Buchnera [20] and Wigglesworthia [21], Wolbachia resides in the cytoplasm of the host cells and mainly relies on maternal inheritance within arthropod lineages. However, in contrast to mutualistic symbionts, phylogenetic analyses suggest that Wolbachia have moved horizontally between distantly related insect lineages multiple times possibly through host–parasite or predator–prey interactions [22,23,24].

Table Table 1. Endosymbionts that are capable of manipulating host sexuality and reproduction.

Click here to display table

Table 1. Endosymbionts that are capable of manipulating host sexuality and reproduction.
EndosymbiontHostsPhenotype
KingdomPhylumClassSpecies
BacteriaProteobacteriaα-ProteobacteriaWolbachia pipientisinsects, crustaceans, arachnids, nematodesCI, MK, PI, FM
Rickettsia sp.insectsMK, PI
γ-ProteobacteriaArsenophonus nasoniaeinsectsMK
FirmicutesMollicutesSpiroplasma poulsoniiinsectsMK
Spiroplasma ixodetis relativeinsectsMK
Cytophaga-Flavobacterium-BacteroidesBacteroidetesCardinium hertigiiinsects, arachnids, nematodesCI, PI, FM
Flavobacteria relativeinsectsMK
EukaryotesMicrosporidiaDihaplophaseaOctosporea effeminanscrustaceansFM
Thelohania herediteriacrustaceansFM
Nosema granulosiscrustaceansFM
Dictyocoela duebenumcrustaceansFM
Amblyospora spp.insectsMK
Parathelohania legeriinsectsMK
Parathelohania obesainsectsMK
NematodaAdenophoreaGasteromermis sp.insectsFM
VirusesUnknownUnknownUnknowncrustaceansMS
Unknown (RNA virus)insectsMK

Abbreviations: CI, cytoplasmic incompatibility; MK, male killing; PI, parthenogenesis induction; FM, feminization; MS, masculinization.

Table Table 2. A List of endosymbionts and their hosts, where feminization, male killing and parthenogenesis induction were described.

Click here to display table

Table 2. A List of endosymbionts and their hosts, where feminization, male killing and parthenogenesis induction were described.
EndosymbiontClassOrderSpeciesReference
(a) Feminizing bacteria
Wolbachia pipientisInsectaHemipteraZyginidia pullula[28]
LepidopteraEurema hecabe[27]
E. madarina[25,26,129]
MalacostracaIsopodaArmadillidium vulgare[130,131,132]
A. nasatum[132,133,134]
Chaetophiloscia elongata[135]
Porcellionides pruinosus[135,136]
Sphaeroma rugicauda[137]
Cardinium hertigiiInsectaHymenopteraEncarsia hispida[44]
ArachnidaTrombidiformesBrevipalpus phoenicis[40]
B. californicus[42]
(b) Feminizing microsporidia
Octosporea effeminansMalacostracaAmphipodaGammarus duebeni[46]
Thelohania herediteria G. duebeni[138]
Nosema granulosis G. duebeni[48]
Dictyocoela duebenum G. duebeni[52]
(c) Other feminizers
Gasteromermis sp.InsectaEphemeropteraBaetis bicaudatus[53]
f factor (unknown)MalacostracaIsopodaArmadillidium vulgare[33]
(d) Male-killing bacteria
Wolbachia pipientisInsectaColeopteraAdalia bipunctata[139]
Tribolium madens[140]
DipteraDrosophila bifasciata[141]
D. borealis[142]
D. innubila[143]
LepidopteraAcraea encedon[128]
A. encedana[144]
Hypolimnas bolina[69]
Ostrinia furnacalis[64]
O. orientalis[145]
O. scapulalis[63,146]
O. zaguliaevi[145]
ArachnidaPseudoscorpionidaCordylochernes scorpioides[147]
Spiroplasma ixodetis relativesInsectaColeopteraAdalia bipunctata[148,149]
Anisosticta novemdecimpunctata[150]
Harmonia axyridis[151,152]
Menochilius sexmaculatus[153]
HemipteraAcyrthosiphon pisum[77,79]
LepidopteraDanaus chrysippus[154]
Ostrinia zaguliaevi[155]
Spiroplasma poulsonii DipteraDrosophila nebulosa[156]
D. neocardini[157]
D. melanogaster[158]
D. ornatifrons[157]
D. paraguayensis[157]
D. willistoni[159]
Rickettsia spp.InsectaColeopteraAdalia bipunctata[149,160,161]
A. decempunctata[162]
Brachys tesselatus[163]
Propylea japonica[164]
FlavobacteriaInsectaColeopteraAdonia variegate[165]
Coccinula sinensis[166,167]
Coleomegilla maculata[168]
Arsenophonus nasoniaeInsectaHymenopteraNasonia vitripennis[169,170]
(e) Male-killing microsporidia
Parathelohania legeri DipteraAnopheles quadimaculatus[171]
Parathelohania obesa A. quadimaculatus[172]
Amblyospora spp. Aedes spp.[173,174]
Amblyospora spp. Culex spp.[174,175,176,177]
Amblyospora spp. Culiseta spp.[174]
(f) Other male killers
Unknown virusInsectaLepidopteraHomona magnanima[59,178]
Armadillidium vulgare[56]
Porcellio dilatatus[56]
P. laevis[56]
(g) Parthenogenesis-inducing bacteria
Wolbachia pipientisInsectaHymenopteraAphytis diaspidis[179,180]
A. lignaensis[179,181]
Aponanagyrus diversicornis[91]
Asobara japonica[92]
Diplolepsis rosae[182]
Encarsia formosa[90]
Eretmocerus mundus[183]
Gronotoma micromorpha[184]
Muscidifurax uniraptor[185,186]
Telenomus nawai[187]
Trichogramma brevicapilliun[188]
T. chilonis[189]
T. cordubensis[188,189]
T. deion[93,188,189]
T. embryophagum[188,189]
T. evanescens[188,189]
T. kaykai[190]
T. oleae[132,189]
T. platneri[188,189]
T. pretiosum[93,188,189]
InsectaThysanopteraFranklinothrips vespiformis[88]
ArachnidaTrombidiformesBryobia praetiosa[40]
Bryobia sp.[40]
Cardinium hertigiiInsectaHymenopteraEncarsia hispida[44]
E. pergandiella[191]
E. protransvena[191,192]
ArachnidaTrombidiformesBrevipalpus phoenicis[43]
Rickettsia spp.InsectaHymenopteraNeochrysocharis formosa[96,193]
Pnigalio soemius[194]

5. Endosymbiont-Induced Feminization: Examples Are Scarce but May Potentially Be More Common

For maternally transmitted microbes, males are an evolutionary dead end. To circumvent this problem, some microbes convert males into functional females so that they can be transmitted to subsequent generations. In insects, microbe-induced feminization has only been found in butterflies and leafhoppers, wherein Wolbachia is the causal agent of feminization. Among non-insects, woodlice, mites and shrimps are known to be feminized by Wolbachia, Cardinium and microsporidians (e.g., Octosporea), respectively (Table 2).

5.1. Feminization of Butterflies by Wolbachia

In two islands located in the southern part of Japan (Tanegashima Island and Okinawa Island), some females of the butterfly Eurema mandarina (Lepidoptera; Pieridae) are known to have a chromosomal constitution of males (ZZ) instead of females (ZW). This incongruence between chromosomal and phenotypic sex can be explained by feminization of genetic males induced by Wolbachia [25,26]. Two distinct strains of Wolbachia, wCI and wFem, have been found in E. mandarina inhabiting these islands. Females having male chromosomes (ZZ) are consistently infected with both wCI and wFem, while females singly infected with wCI are true females (ZW). Despite having complete male chromosomes, ZZ females are morphologically and behaviorally completely female and fully fertile. Moreover, when such individuals are treated with tetracycline hydrochloride, a bacteriostatic antibiotic, during larval development, they develop as butterflies with intersexual morphology. Before carrying out this experiment, the appearance of intersexes in the treated generation was not expected because perturbation of sex determination was believed to occur during early embryogenesis when the sex-determining genes start to be expressed. However, the appearance of intersexes clearly shows that Wolbachia needs to be present during larval development for complete feminization. The fact that butterflies with more male-like characters were generated after longer antibiotic treatments shows that Wolbachia acts continuously during larval development [26] (Figure 2). The presence of Wolbachia during the embryonic stage seems to be necessary for complete feminization since antibiotic treatment during the whole larval stages also generated intersexes instead of complete males. We recently found that the splicing pattern of the sex-determining gene doublesex (dsx) changes according to the Wolbachia infection status (Narita et al., in preparation). Wolbachia-induced feminization has also been found in Eurema hecabe, a sibling species of E. mandarina. The two Wolbachia strains found in feminized E. hecabe are indistinguishable from wCI and wFem derived from E. mandarina based on multi-locus sequence typing [27].

Insects 03 00161 g002 1024
Figure 2. Effects of the antibiotic on the sexual phenotype of the butterfly Eurema mandarina infected with two strains of Wolbachia [26]. (a) A normal female and an intersex generated by antibiotic treatment. The intersex has both male and female features in external morphology. (b) Ovarioles of normal females. (c) The reproductive organs of the intersex. Both male-specific organs (testes) and female-specific organs (ovarioles) occur in an individual treated with tetracycline hydrochloride during larval stages. (d) Schematic illustration showing the effect of antibiotic treatment of larvae on adult sexual phenotype. The longer the treatment, the more male-like phenotype appears. NT: non-treated. Tet: tetracycline-treated.

Click here to enlarge figure

Figure 2. Effects of the antibiotic on the sexual phenotype of the butterfly Eurema mandarina infected with two strains of Wolbachia [26]. (a) A normal female and an intersex generated by antibiotic treatment. The intersex has both male and female features in external morphology. (b) Ovarioles of normal females. (c) The reproductive organs of the intersex. Both male-specific organs (testes) and female-specific organs (ovarioles) occur in an individual treated with tetracycline hydrochloride during larval stages. (d) Schematic illustration showing the effect of antibiotic treatment of larvae on adult sexual phenotype. The longer the treatment, the more male-like phenotype appears. NT: non-treated. Tet: tetracycline-treated.
Insects 03 00161 g002 1024

5.2. Feminization of Leafhoppers by Wolbachia

A second case of Wolbachia-induced feminization in diploid insects has been found in the leafhopper Zyginidia pullula (Homoptera; Cicadellidae) [28]. In Z. pullula, females have two X chromosomes (2n = 8AA + XX) while males have only one X chromosome (2n = 8AA + X0). This is contrary to the situation of the Eurema butterflies wherein females have one Z chromosome and males have two Z chromosomes. When Wolbachia-infected Z. pullula females collected in northern Italy were mated with males, they produced exclusively female broods. Close inspection of these female broods revealed that about half of them had an intersexual morphology (i.e., showing upper pygofer appendages, a typical male secondary sexual character), while the rest of the broods had the normal female phenotype. While the karyotypes of the normal females were XX, those with the upper pygofer appendages were X0 and were thus feminized males. Administration of tetracycline hydrochloride to Wolbachia-infected females (phenotypically normal female) resulted in a nearly 1:1 sex ratio in the subsequent generation [28]. The incongruence between the phenotypic sex and genomic sex implies the presence of epigenetic modification. Negri et al. [29] compared the DNA methylation patterns between intersexual and normal Z. pullula by performing a methylation-sensitive random PCR and found that females with the male upper pygofer appendages showed a female methylation pattern. On the other hand, some rare feminized males bore testes instead of ovaries. And these individuals showed a male methylation pattern. These findings suggest that Wolbachia induces feminization by disrupting male imprinting.

5.3. Feminization of Woodlice by Wolbachia

Wolbachia can also cause feminization in non-insect arthropods. Wolbachia-induced feminization in the woodlouse Armadillidium vulgare (Isopoda; Armadillidiidae) has a long research history [30]. As known in several insect species, the A. vulgare Wolbachia appear to be quite sensitive to high temperatures. Very young Wolbachia-infected females of A. vulgare reared at 30 °C gradually acquire a male phenotype [31,32]. In addition to Wolbachia, a non-bacterial feminizing factor (f) can also force chromosomal males of A. vulgare to become phenotypic functional females. The f factor is suspected to be a genetic element derived from the Wolbachia genome that becomes inserted into the host nuclear genome [33]. Genes resisting the feminizing effects or the transmission of feminizing elements have been found in natural populations of A. vulgare [34,35], thus illustrating the conflict between the feminizing elements and the rest of the host genome.

Unlike insects, androgenic hormone, a sex hormone secreted from a specific male organ called the androgenic gland, profoundly affects male sexual differentiation in A. vulgare [36,37]. In this respect, sex determination in A. vulgare seems to be more labile compared with that in insects. It is considered that Wolbachia feminize genetic males by disrupting the secretion of androgenic hormone. Before the discovery of feminization in Eurema butterflies, Wolbachia was assumed to be incapable of inducing complete feminization in insects [31]. This assumption appeared reasonable considering the substantial differences in the sex-determining systems between insects and non-insects. The actual occurrence of Wolbachia-induced feminization in both insects and non-insects may imply the presence of some common mechanism of sex determination that is targeted by Wolbachia. Alternatively, the feminization found in insects and non-insects may have distinct underlying mechanisms and only be superficially similar.

5.4. Feminization of Mites and Wasps by Cardinium Bacteria

The false spider mite, Brevipalpus phoenicis (Acarina; Tenuipalpidae), along with two closely related species, Brevipalpus obovatus and Brevipalpus californicus, is known to reproduce by thelytokous parthenogenesis [38]. Surprisingly, B. phoenicis females have a haploid genome, which was confirmed by fluorescence microscopy and variations at nine microsatellite loci [39]. Treatment of adult females with tetracycline hydrochloride led to a drastic increase in the proportion of male offspring (from 5.6% to 50.6%), suggesting that haploid individuals were feminized by a bacterium [40]. This bacterium is now known to be a member of the genus Cardinium [41]. Haploid individuals of the other species, B. obovatus and B. californicus, were also found to be feminized by Cardinium [42,43]. A feminizing effect of Cardinium was also found in the wasp Encarsia hispida (Hymenoptera; Aphelinidae), in which thelytokous parthenogenesis is known [44] (see Section 7.1).

5.5. Feminization of Shrimps by Microsporidian Parasites

Sex determination of the amphipod crustacean Gammarus duebeni (Amphipoda; Gammaridae) is more complex compared with that of insects. Basically, males and females are determined by a balance of a polyfactorial system of allelic sex genes located on several pairs of chromosomes, but the photoperiod can profoundly affect the sex determination [30]. Moreover, based on detailed breeding experiments, cytoplasmic factors were considered to act as female determiners in G. duebeni [45]. Subsequently, it was found that several microsporidian parasites such as Octosporea effeminans, Nosema granulosis and Dictyocoela duebenum living in the host cytoplasm alter male G. duebeni to functional females [46,47,48,49,50,51,52]. Rodgers-Gray et al. [50] demonstrated that Nosema manipulate the sex differentiation of G. duebeni by preventing androgenic gland differentiation, androgenic gland hormone production and consequently male differentiation. This is in agreement with observations of Wolbachia-induced feminization in A. vulgare. Although taxonomically unrelated (eukaryotes and prokaryotes), these feminizers may manipulate their crustacean hosts through a common mechanism.

5.6. Feminization Induced by Non-Microbes

Feminization can also be caused by non-microbes. According to Vance [53], more than 10% of wild-caught adults of the mayfly Baetis bicaudatus (Ephemeroptera; Baetidae) are parasitized by nematode Gasteromermis sp. (Nematoda; Mermithidae). None of the parasitized individuals (n = 126) contained visible eggs, ovaries or testes. Among them, the external morphology of 82 individuals (65%) was indistinguishable from that of normal females. The remaining 44 parasitized individuals showed an array of intersexual morphologies. Moreover, measurement of the DNA contents by flow cytometry suggested that the parasitized individuals having intersexual morphologies (“parasitized intersexes”) were genetically male individuals while parasitized individuals visibly indistinguishable from normal females (“parasitized females”) were composed of both genetically female and genetically male individuals [53]. The behaviors of the mayfly are also changed by the nematode. Unparasitized males form swarms near the river and do not return to the water after they have emerged. Vance [53] found that all 418 swarming individuals were unparasitized males and that the parasitized individuals showed ovipositing behavior, which is never seen in unparasitized males. Laboratory studies demonstrated that parasitized individuals (both parasitized females and parasitized intersexes) became very agitated shortly after emergence as adults. Within 3–6 hours, all the parasitized mayflies had crawled into the water down the side of the rock. The nematodes could then be seen escaping through a puncture wound in the mayfly’s abdomen [53]. Therefore, the Gasteromermis nematode can horizontally transmit to a new host. The mayflies were killed by the emergence of the nematode. The feminization in this case seems to be an adaptive strategy for the Gasteromermis nematodes, which do not have a vertical route of transmission.

Although less conspicuous than the case of the mayfly, perturbation of the development of secondary sexual characters by endosymbionts or parasites has also been found in various insects [54,55], in which the adaptive role of feminization remains unclear.

5.7. Masculinization Induced by Viruses

In woodlice such as Porcellilo dilatatus, Porcellio laevis and A. vulgare, an intersexual trait in genetic females is known to be transmitted from both parents. Viruses are considered to be the causal agent of the masculinization in genetic females because male characters disappeared after heat treatment, male characters appeared after injection of a 0.22-μm-filtered tissue extract and intersexuality was correlated with the presence of cytoplasmic viral particles [56]. The adaptive significance of the masculinizing effect remains unknown.

6. Endosymbiont-Induced Male Killing: An Easily Evolved Trait?

Compared with feminization, male killing is more common in insects (Table 2). Moreover, male killers have been found in taxonomically diverse microorganisms (e.g., bacteria in the genus Spiroplasma, Wolbachia, Rickettsia, Arsenophonus; unicellular eukaryotes in the phylum Microsporidia; and RNA viruses). Therefore, male killing may be easy to evolve [57]. Among them, Wolbachia and Spiroplasma are prevalent male killers among insects. In bacteria-induced male killing, only males are typically killed during embryonic or early larval stages which is called “early” male killing. On the other hand, in male killing induced by microsporidia and RNA viruses, the males are killed during later development (typically during the last stage of larva) which is called “late” male killing [58,59].

6.1. Function of Dosage Compensation is Necessary for Male Killing

Drosophila have two X chromosomes in females and only one X chromosome in males. By hypertranscribing the X-linked genes in males, Drosophila dissolve the imbalance in the gene dosage of the X-linked genes, which is often called dosage compensation. This process requires the formation of the dosage compensation complex (DCC), which consists of MSL-1, MSL-2, MSL-3, MLE and MOF [60].

How do male killers discriminate between males and females? Using classic genetic experiments, Veneti et al. [61] tested the ability of Spiroplasma to kill D. melanogaster males carrying mutations in genes encoding the DCC. Interestingly, Spiroplasma failed to kill males lacking any of the five protein components (MSL-1, MSL-2, MSL-3, MLE and MOF) of the DCC. Therefore, although the direct mechanism of male killing is elusive, their study clearly showed that the presence of the functional DCC is necessary for Spiroplasma to cause male killing in D. melanogaster.

6.2. Male Killing by Lethal Effect of Feminization?

A mechanistic implication of male killing was obtained in our study on Wolbachia-induced male killing in the moth Ostrinia scapulalis (Lepidoptera; Crambidae), namely that males are killed by the feminizing effect of Wolbachia. Similar to other lepidopteran species, the sex chromosome constitution of O. scapulalis is ZZ in males and ZW in females. Wolbachia-infected females produce both ZZ and ZW offspring, but only ZZ offspring die during late embryonic and larval development. Interestingly, when adult females were fed with sucrose containing an antibiotic (tetracycline hydrochloride) prior to oviposition, they produced eggs that developed as intersexes as well as normal males and females [62,63,64] (Figure 3). The forewings of these intersexes showed a clear mosaic pattern of male-color and female-color sectors. Some of the intersexes had the bursa copulatrix, the female organ which accepts spermatophores from males during mating. Surprisingly, none of the tissues of the intersexes, including the bursa copulatrix, had a W chromosome, thus representing the pure male genotype (ZZ). Therefore, it is clear that the bursa copulatrix in these intersexes shows a female phenotype under a male genotype. Based on these findings, we concluded that Wolbachia has a feminizing effect on Ostrinia males but its full expression, which occurs in the natural condition, is lethal for genetic males. A reduction in the Wolbachia density during embryogenesis may result in partially feminized individuals (intersexes), probably due to the attenuated expression of the feminizing effect. The sex-determining gene dsx of these intersexes was shown to exhibit both male and female splicing patterns [65]. Reported very recently was conclusive evidence for the feminization as a mechanism of male killing in Ostrinia. In Wolbachia-infected O. scapulalis, female-specific splicing in dsx was observed in all the embryos including those with male genotype (ZZ) which were destined to die [66].

Insects 03 00161 g003 1024
Figure 3. Effects of antibiotics on sexual phenotype of the moth Ostrinia scapulalis infected with Wolbachia [62]. (a) Intersexual individuals generated by tetracycline treatment. Black and white depict male-like color and female-like color, respectively. (b) The proportion of phenotypic females (white), intersexes (gray) and phenotypic males (black) among individuals whose mothers were treated with tetracycline hydrochloride prior to oviposition.

Click here to enlarge figure

Figure 3. Effects of antibiotics on sexual phenotype of the moth Ostrinia scapulalis infected with Wolbachia [62]. (a) Intersexual individuals generated by tetracycline treatment. Black and white depict male-like color and female-like color, respectively. (b) The proportion of phenotypic females (white), intersexes (gray) and phenotypic males (black) among individuals whose mothers were treated with tetracycline hydrochloride prior to oviposition.
Insects 03 00161 g003 1024

In addition, this case of male killing in Ostrinia showed another aspect when Wolbachia was completely eliminated. While other cases of male killing exhibit a normal 1:1 sex ratio after elimination of the male killers in their mothers, the elimination of Wolbachia from Ostrinia females by treatment with tetracycline hydrochloride during the whole larval stages resulted in the appearance of only males in the subsequent generation. This finding was erroneously interpreted as evidence for the Wolbachia-induced feminization [67,68]. However, later studies clearly showed that half of these offspring having female karyotype (ZW) die during late embryonic and larval development [63,64]. These individuals were recently shown to exhibit male-specific splicing in dsx [66], which implies that a female-determining factor, possibly located on W chromosome is degraded in Ostrinia and the Wolbachia substitutes the sex-determining role.

6.3. Hidden Male Killing

In some cases, male killing is masked or suppressed. Wolbachia-induced male killing was reported in the butterfly Hypolimnas bolina (Lepidoptera; Nymphalidae) [69,70]. In H. bolina, a strong female-biased sex ratio has been maintained for more than 100 years [71]. Surprisingly, in some populations, host resistance to male killing has recently spread and reached fixation [72,73,74]. In these populations, Wolbachia ceased to kill males but was not excluded from the host population. Hornett et al. [75] revealed that the same Wolbachia was inducing cytoplasmic incompatibility in the butterfly. This finding can explain the persistence of Wolbachia in the H. bolina population without inducing male killing.

In Drosophila recens, the sex ratios are not affected by naturally-occurring Wolbachia. This Wolbachia causes strong cytoplasmic incompatibility (see below) in D. recens and the infection frequency of Wolbachia in this species is 98%. On the other hand, none of the individuals are infected with Wolbachia in Drosophila subquinaria. Interestingly, introgression of the D. recens Wolbachia into D. subquinaria by hybridization and backcrossing resulted in the expression of male killing [76]. Furthermore, crossing experiments have demonstrated that the resistance to male killing is dominant, autosomal, multigenic and dependent on the zygotic, not maternal, genotype [76]. Similar to the case of H. bolina, male killing was masked by the fixation of resistant genes in D. recens. If there were no sister species, we would not have been able to reveal the potential for male killing in this insect.

The pea aphid Acyrthosiphon pisum is known to exhibit cyclical parthenogenesis. Previously, infection of Spiroplasma was only investigated under the asexual reproduction mode [77,78]. The effects of various endosymbionts in A. pisum under the sexual reproduction mode were recently analyzed and Spiroplasma was revealed to cause male killing [79]. The fact that Spiroplasma found in A. pisum was monophyletic with other male-killing Spiroplasma in ladybird beetles, butterflies and moths may indicate a common origin of the male-killing ability.

6.4. Is Timing of Male Killing Important?

Why do these microbes kill males? Killing only males during early development (early male killing) is considered advantageous for maternally transmitted microbes for three possible reasons that are not mutually exclusive [80,81]: (i) females can gain extra resources that were to be allocated to their brothers (resource reallocation); (ii) mating with sibling can be avoided (inbreeding avoidance); and (iii) microbes can be transmitted from dead (or dying) males to females by oral intake (horizontal transmission by cannibalism). On the other hand, resource reallocation cannot explain late male killing because males would consume nearly all the resources necessary for their own development before they are killed. Microsporidian parasites that cause late male killing in mosquitoes have intermediate hosts (copepods). Killing the mosquito larvae of large body size (late male killing) is considered to be an optimal strategy for the microsporidians because they can maximize their number and thereby increase their probability of transmission to copepod hosts [58,82].

Thus, the classification of male-killing phenotypes is based not only on the timing of the action, but also on the evolutionary strategies adopted by the symbionts and microbial agents responsible for the phenotypes (i.e., bacteria versus eukaryotes and viruses). Therefore, it has been assumed that early and late male killing are fundamentally different phenomena whose underlying mechanisms are also entirely different [83,84]. In Drosophila, however, a Spiroplasma strain that normally cause early male killing was also found to induce late male killing depending on the maternal host age, in that male-specific mortality of larvae and pupae was more frequently observed in the offspring of young females [85]. Since the lowest Spiroplasma density and occasional male production were also associated with newly emerged females, we proposed a density-dependent hypothesis for the expression of the early and late male-killing phenotypes. Early male killing and late male killing could be regarded as alternative strategies adoptable by microbial reproductive manipulators [85].

The timing of male killing can be artificially changed in the Wolbachia-infected butterfly, H. bolina. Treatment of Wolbachia-infected adult females with tetracycline, a bacteriostatic antibiotic, produced a delay in the timing of male death. On the other hand, treatment of the surviving larvae with rifampicin, a bactericidal antibiotic, rescued the males. Based on these findings, it was hypothesized that Wolbachia possesses the ability to kill males through bacterial activity during larval development [86]. This phenomenon argues against the view that male killing is achieved by specifically targeting an early developmental process within the sex determination pathway. This is in line with the case of Wolbachia-induced feminization in the butterfly E. mandarina, wherein Wolbachia needs to be present during larval development for complete feminization [26].

7. Parthenogenesis Induction: Conversion of Genetic Males to Genetic Females or Feminization Following Diploidization?

Microbe-induced thelytokous parthenogenesis has been reported in haplodiploid arthropods, such as wasps, thrips and mites [19,87,88] (Table 2). On the other hand, there is no conclusive evidence for the microbe-induced parthenogenesis in diplodiploid organisms, although involvement of Wolbachia in thelytokous parthenogenesis in springtails and booklice is suspected based on the presence of Wolbachia [89]. In most of the known cases, microbe-induced parthenogenesis is achieved by altering the male genotype (haploid) to the female genotype (diploid), i.e., restoration of diploidy. In this sense, the phenomenon can also be referred to as conversion of genetic males to genetic females [49]. At least in the wasp E. hispida, however, the restoration of diploidy is not sufficient for female development, and thus diploidization and feminization can be separate processes [44]. Frequently, certain species or populations are fixed for parthenogenesis-inducing microbes. In such cases, selection will not act on host genes involved in male and female sexuality (e.g., mating behaviors, sperm production, or sex pheromone production). Thus, mutations in such genes are expected to accumulate, resulting in eventual degeneration of sexuality and irreversible parthenogenesis. This process appears to be occurring in species of wasps [90,91,92] and thrips [88]. In contrast, sexual functions are not degenerated in Trichogramma deion, wherein parthenogenesis-inducing Wolbachia is not fixed and genetic exchange occurs between sexual and asexual individuals [93].

7.1. Mechanisms of Microbe-Induced Parthenogenesis

There are various mechanisms underlying microbe-induced parthenogenesis. Gamete duplication appears to be common in Wolbachia-infected hymenopteran wasps [93,94]. In wasps such as Trichogramma species and Leptopilina clavipes, meiosis is normal, but during the first mitotic division, the chromosomes fail to segregate in metaphase, resulting in diploidization of the nucleus [93,94]. In Muscidifurax uniraptor, on the other hand, meiosis and the first mitotic division are normal, but during the second mitotic division, diploid females are produced by the fusion of two cell nuclei [95]. Each offspring produced by gamete duplication is a homozygote at all loci and is not a genomic copy of its mother. On the other hand, the genotype of all offspring was indistinguishable from their mothers in Rickettsia-induced parthenogenesis occurring in Neochrysocharis formosa (Hymenoptera; Eulophidae) based on the polymorphisms in a microsatellite locus [96]. Moreover, by excluding the possibility of automictic parthenogenesis with central fusion, which has been observed in some ants [97,98], and using cytogenetic observations, Adachi-Hagimori et al. [96] concluded that apomictic parthenogenesis is the underlying mechanism wherein eggs do not undergo meiosis. Similarly, in the Wolbachia-induced parthenogenesis in the mite Bryobia praetiosa, the genotypes of the mothers and offspring are indistinguishable based on the polymorphisms in three microsatellite loci [40]. Therefore, apomictic parthenogenesis is the likely mechanism for this phenomenon as Weeks et al. [40] assumed, but another possibility of automictic parthenogenesis with central fusion cannot be completely excluded owing to the lack of cytogenetic observations.

It has been naturally assumed that diploidization induced by microbes automatically leads to female development. However, a recent finding indicates that, in the wasp E. hispida, Cardinium-induced parthenogenesis induction does not occur by diploidization alone. Feeding antibiotics to infected adult E. hispida females resulted in uninfected male offspring. By karyotype observations and flow cytometry analyses, Giorgini et al. [44] demonstrated that these males were diploid. This finding indicates that at least in E. hispida, diploidy restoration is necessary but not sufficient for female development. Thus, Cardinium is required to feminize diploid male embryos [44]. In this sense, this example should also be added to the list of microbe-induced feminization. As Giorgini et al. [44] argues, it might be necessary to consider the possibility that the mechanism of Wolbachia-induced parthenogenesis is also comprised of two separate steps, i.e., diploidization and feminization.

8. Other Phenotypes

Here we briefly overview the other conspicuous effects caused by the above endosymbionts (Table S1).

8.1. Cytoplasmic Incompatibility

Cytoplasmic incompatibility is the most common phenotype of Wolbachia and Cardinium infection (Table S1). In diploid organisms, cytoplasmic incompatibility is an embryonic lethality that results in sperm and eggs having different cytoplasmic contents [17,99,100]. The effect arises from changes in the gamete cells caused by cytoplasmic (intracellular) parasites like Wolbachia and Cardinium, which infect a wide range of insect species. Cytoplasmic incompatibility occurs when a Wolbachia-infected male mates with a female that is either uninfected (unidirectional cytoplasmic incompatibility) or infected by another Wolbachia strain (bidirectional cytoplasmic incompatibility). Any other combinations of crosses are compatible. An infected female is compatible with an uninfected male or any infected male of the same Wolbachia strain. On the other hand, an uninfected female is only compatible with an uninfected male.

Cytoplasmic incompatibility in haplodiploid hosts may lead to haploid (male) offspring. Cytoplasmic incompatibility produces distinct phenotypes in three closely related haplodiploid hymenopteran species of the genus Nasonia, namely mortality in Nasonia longicornis and Nasonia giraulti, and conversion to male development in Nasonia vitripennis [101].

8.2. Beneficial Effects of Wolbachia and Spiroplasma on Hosts

In addition to arthropods, some nematodes are also hosts of Wolbachia [102]. In filarial nematodes, Wolbachia does not seem to exhibit selfish behaviors like reproductive manipulations. Instead, Wolbachia is necessary for the host nematodes. Elimination of Wolbachia from filarial nematodes generally results in either death or sterility of the nematodes [103]. Consequently, current strategies for the control of filarial nematode diseases include elimination of Wolbachia via treatment with the antibiotic doxycycline rather than far more toxic anti-nematode medications [102].

In the bedbug Cimex lectularius (Hemiptera; Cimicidae), Wolbachia appears to be an obligate nutritional mutualist. Wolbachia is specifically localized in the bacteriomes, a specialized organ for endosymbiotic bacteria, and vertically transmitted via the somatic stem cell niche of germalia to oocytes. The transmitted Wolbachia infects the incipient symbiotic organ at an early stage of embryogenesis. Elimination of Wolbachia results in retarded growth and sterility of the host insect. However, these deficiencies are rescued by oral supplementation of B vitamins. These findings suggest that Wolbachia provides essential nutrition for this host [104].

In the parasitic wasp Asobara tabida (Hymenoptera; Braconidae), a strain of Wolbachia is necessary for oogenesis [105]. It is also known that Wolbachia rescues the oogenesis defect in a Sex-lethal mutant of D. melanogaster [106].

Increasing attention has been paid to the recent findings that Wolbachia can protect Drosophila against pathogenic RNA viruses such as Drosophila C virus, Cricket Paralysis virus, Flock House virus, Nora virus [107,108,109] and West Nile virus [110], as well as the fungus Beauveria bassiana [111]. Interestingly, in mosquitoes, the presence of transinfected Wolbachia interferes with a wider range of pathogens including nematodes and bacteria [112], Dengue virus and Chikungunya virus [113,114], as well as the avian and rodent malaria parasites Plasmodium gallinaceum [114] and Plasmodium berghei [115]. Biological control of mosquito-borne diseases is at the testing stage [116,117,118].

Spiroplasma also has similar effects on its hosts. Drosophila neotestacea, a fly that feeds on mushrooms, suffers complete loss of fecundity when parasitized by the nematode Howardula aoronymphium (Allantonematidae; Tylenchida). However, flies infected with Spiroplasma have near normal fecundity when parasitized by the nematode [119].

9. Transition of the Biological Systems of Arthropods by Sex-Associated Microbes

Despite the diversity of the sex-determining systems of arthropods and the diversity of microbial agents, the types of reproductive manipulation seem to be limited. For example, male killing occurs in male heterogametic insects, female heterogametic insects and haplodiploid insects. Moreover, the causal agents of male killing include a wide variety of bacteria as well as some eukaryotic organisms. Similarly, feminization occurs in arthropods having a ZW-ZZ system (butterflies, woodlice and shrimps), XX-X0 system (leafhoppers) and haplodiploid system (mayflies), and the causal agents of feminization also include bacteria and eukaryotes. Considering the stability of vertical transmission, these endosymbiotic microbes can be assumed to be sex-determining elements in the cytoplasm that work together with other sex-determining genes in the nucleus. It can also be assumed that the effects of these cytoplasmic sex-determining elements are relatively well conserved among arthropods.

Moreover, the presence of resistance genes against male killers and feminizers has been documented in some arthropods [73,76,120]. These resistance genes against microbial effects might be recognized as sex-determining genes. Although speculative, some of the sex-determining genes, such as Sex-lethal, intersex, daughterless and sisterless in D. melanogaster, could originally have arisen as resistance genes against the effects of the microbes and become fixed within the species. The diversity and complexity of the sex-determining systems of insects might have been generated by an arms race involving repeated invasion and/or mutation of various microbes and other genetic elements, which may not necessarily exist at the present time [2].

Currently, almost nothing is known about the molecular mechanisms of these acquired sex-determining systems. Regardless of the actual mechanisms, however, we can hypothesize a possible link between the microbial effects and the diversity of arthropod sex determination. As shown in Figure 4, one can place the possible effects of endosymbionts in the informational flow of sex-determining signals. To cause feminization, for example, endosymbionts need to simultaneously affect the sex determination in the germline, somatic line and central nervous system, each of which is considered to depend on a distinct expression cascade consisting of different genes. To achieve this, the endosymbionts may interfere with certain elements in the upstream of the sex-determining gene cascade that play pivotal roles, like Sxl in Drosophila, in sex determination in different lineages [4,121]. On the other hand, at least in Ostrinia moths, male killing may involve interactions with downstream genes, which may result in simultaneous expression of sexually antagonistic, male-specific and female-specific, genes and lead to lethality [63,65]. Although it was previously assumed that diploidization is the only effect of the parthenogenesis inducers in haplodiploid insects, recent findings in the E. hispida wasps imply that we need to take into account the possibility that endosymbionts may cause two distinct effects in order, i.e., diploidization and feminization, the latter of which can be integrated into the feminization phenomena found in diploid insects. It is tempting to consider that, although endosymbionts work very simply (e.g., by only producing some proteins), the diversity and complexity of the host biology generate the diverse effects on the phenotype. A straightforward and frequently-used approach to elucidate the mechanisms of microbial effects on their hosts would be to compare the gene expression profiles between infected and uninfected insects. However, despite various attempts to compare the gene expression profiles [122,123,124,125], the mechanisms of reproductive manipulation still remain unclear. We consider that the mechanisms of microbe-induced reproductive manipulation will be clarified by a deeper understanding of insect development and physiology. Moreover, genomic data of endosymbionts and their hosts together with sophisticated mutagenesis techniques may allow reverse genetics approaches to elucidate the precise mechanism of reproductive manipulations.

Insects 03 00161 g004 1024
Figure 4. A parsimonious explanation of male killing (MK), feminization (FM) and parthenogenesis induction (PI). Here, MK, FM and PI are all assumed to be associated with the sex-determining process. The hypothetical flows of the male-determining and female-determining information are shown with blue and red arrows, respectively. Horizontal arrows indicate manipulation by endosymbionts. Note that this explanation is a hypothesis and may not necessarily reflect the actual mechanisms.

Click here to enlarge figure

Figure 4. A parsimonious explanation of male killing (MK), feminization (FM) and parthenogenesis induction (PI). Here, MK, FM and PI are all assumed to be associated with the sex-determining process. The hypothetical flows of the male-determining and female-determining information are shown with blue and red arrows, respectively. Horizontal arrows indicate manipulation by endosymbionts. Note that this explanation is a hypothesis and may not necessarily reflect the actual mechanisms.
Insects 03 00161 g004 1024

10. Population-Level Effects of Endosymbiotic Microbes on Their Hosts

The spread and fixation of male killers and feminizers should lead to the absence of males and the extinction of the sexually reproducing hosts [126]. Moreover, a strongly-biased population sex ratio leads to a drastic reduction in the effective population size, which may inevitably cause inbreeding. Furthermore, inbreeding may cause a reduction in fecundity, which may increase the probability of extinction. In fact, a decrease in heterozygosity, an indication of inbreeding, was shown to increase the risk of extinction of island populations of the butterfly Melitaea cinxia [127]. However, endosymbiont-driven extinction has not been empirically demonstrated thus far.

Instead, in some butterflies, a Wolbachia-induced extreme bias in the population sex ratio was shown to be maintained for substantially long periods of time [71,128]. Moreover, resistance genes against male killers and feminizers have been documented [34,35]. Is this a usual reaction of hosts against the invasion of male killers and feminizers? We should be aware, however, that it is practically difficult to show the occurrence of population extinction caused by the spread and fixation of male killers and feminizers. Under the evolutionary timescale, the time required for sex-ratio distorters to spread among the host population can be too short for resistance genes to evolve. Owing to the short timeframe, the probability that we can observe a gradual decline in a population size is extremely low, even if endosymbiont-driven extinctions occur repeatedly in natural populations.

11. Conclusions

The accumulating data demonstrate that a variety of sexual aberrations in arthropods is imposed by endosymbionts (e.g., bacteria, protists, nematodes, insects and viruses). Although reducing the diverse phenomena to simple fundamental mechanisms may be dangerous and is not completely possible at present, some of the sexual aberrations can be explained by the interference with the sex-determining systems: for instance, male killing can be induced by the lethal effect of Wolbachia-induced feminization in a moth; parthenogenesis can be induced by Cardinium that feminizes haploid males in a mite; and diploid males are transformed by Cardinium into parthenogenetic females in a wasp. Importantly, these endosymbionts may profoundly affect host behavior, ecology and population structure. At present, the molecular mechanisms of any of the sexual aberrations are largely unknown. The deeper understanding of the arthropod physiology and development as well as arthropod and microbial genomes may allow us to elucidate the mechanism of the sexual aberrations, which will benefit both applied sciences (e.g., management of agricultural and medical pests and mass rearing of beneficial organisms) and other areas of biological sciences.

Acknowledgments

We are grateful to Tetsuya Adachi-Hagimori for reading our manuscript and the Grant for Basic Science Research Projects of the Sumitomo Foundation, Grant-in-Aid for Young Scientists (B) and Grant-in-Aid for JSPS fellows for financial support.

References

  1. Bull, J.J. Evolution of Sex Determining Mechanisms; The Benjamin/Cummings Publishing Company, Inc.: Menlo Park, CA, USA, 1983; p. 316. [Google Scholar]
  2. Werren, J.H.; Beukeboom, L.W. Sex determination, sex ratios, and genetic conflict. Ann. Rev. Ecol. Evol. Syst. 1998, 29, 233–261. [Google Scholar] [CrossRef]
  3. Heimpel, G.E.; de Boer, J.G. Sex determination in the hymenoptera. Ann. Rev. Entomol. 2008, 53, 209–230. [Google Scholar] [CrossRef]
  4. Schütt, C.; Nöthiger, R. Structure, function and evolution of sex-determining systems in Dipteran insects. Development 2000, 127, 667–677. [Google Scholar]
  5. Nöthiger, R.; Steinmann-Zwicky, M. A single principle for sex determination in insects. Cold Spring Harb. Symp. Quant. Biol. 1985, 50, 615–621. [Google Scholar] [CrossRef]
  6. Hoy, M.A. Insect Molecular Genetics, 2nd ed.; Academic Press/Elsevier: San Diego, CA, USA, 2003; p. 560. [Google Scholar]
  7. Verhulst, E.C.; van de Zande, L.; Beukeboom, L.W. Insect sex determination: It all evolves around transformer. Curr. Opin. Genet. Dev. 2010, 20, 376–383. [Google Scholar] [CrossRef]
  8. Laugé, G. Sex determination: Genetic and epigenetic factors. In Comprehensive Insect Physiology, Biochemistry and Pharmacology, Vol. 1. Embryogenesis and Reproduction; Kerkut, G.A., Gilbert, L.I., Eds.; Pergamon Press: Oxford, UK, 1985; pp. 295–318. [Google Scholar]
  9. Narita, S.; Pereira, R.A.S.; Kjellberg, F.; Kageyama, D. Gynandromorphs and intersexes: Potential to understand the mechanism of sex determination in arthropods. Terrestrial Arthropod Rev. 2010, 3, 63–96. [Google Scholar] [CrossRef]
  10. Brust, R. Temperature-induced intersexes in Aedes mosquitoes: Comparative study of species from Manitoba. Can. Entomol. 1968, 100, 879–891. [Google Scholar] [CrossRef]
  11. Horsfall, W.R.; Anderson, J.F. Suppression of male characteristics of mosquitoes by thermal means. Science 1961, 133, 1830. [Google Scholar]
  12. Horsfall, W.R.; Anderson, J.F. Thermally induced genital appendages on mosquitoes. Science 1963, 141, 1183. [Google Scholar]
  13. Goldschmidt, R. Lymantria. Bibliogr. Genet. 1934, 11, 1–186. [Google Scholar]
  14. Mosbacher, G.C. Die Intersexualität bei Lymantria dispar L. (Lepidoptera). Z. Morphol. Tiere 1973, 76. [Google Scholar]
  15. Mosbacher, G.C. Sex specific cell differentiation in different types of intersexes of Lymantria dispar L. In Intersexuality in the Animal Kingdom; Reinboth, R., Ed.; Springer: Berlin, Germany, 1975; pp. 146–157. [Google Scholar]
  16. Seiler, J. Sexuality as developmental process. In Genetics Today. Proc. XI Int. Congress of Genetics, The Hague 1963; Geerts, S.J., Ed.; Pergamon: Oxford, UK, 1965; pp. 199–207. [Google Scholar]
  17. Werren, J.H. Biology of Wolbachia. Ann. Rev. Entomol. 1997, 42, 587–609. [Google Scholar] [CrossRef]
  18. Werren, J.H.; Baldo, L.; Clark, M.E. Wolbachia: Master manipulators of invertebrate biology. Nat. Rev. Microbiol. 2008, 6, 741–751. [Google Scholar] [CrossRef]
  19. Stouthamer, R.; Breeuwer, J.A.J.; Hurst, G.D.D. Microbial Manipulator of Arthropod Reproduction. Ann. Rev. Microbiol. 1999, 53, 71–102. [Google Scholar] [CrossRef]
  20. Douglas, A.E. Nutritional interactions in insect-microbial symbioses: Aphids and their symbiotic bacteria Buchnera. Ann. Rev. Entomol. 1998, 43, 17–37. [Google Scholar] [CrossRef]
  21. Weiss, B.L.; Wang, J.; Aksoy, S. Tsetse immune system maturation requires the presence of obligate symbionts in larvae. PLoS Biol. 2011, 9, e1000619. [Google Scholar] [CrossRef]
  22. Baldo, L.; Ayoub, N.A.; Hayashi, C.Y.; Russell, J.A.; Stahlhut, J.K.; Werren, J.H. Insight into the routes of Wolbachia invasion: High levels of horizontal transfer in the spider genus Agelenopsis revealed by Wolbachia strain and mitochondrial DNA diversity. Mol. Ecol. 2008, 17, 557–569. [Google Scholar]
  23. Russell, J.A.; Goldman-Huertas, B.; Moreau, C.S.; Baldo, L.; Stahlhut, J.K.; Werren, J.H.; Pierce, N.E. Specialization and geographic isolation among Wolbachia symbionts from ants and lycaenid butterflies. Evolution 2009, 63, 624–640. [Google Scholar] [CrossRef]
  24. Stahlhut, J.K.; Desjardins, C.A.; Clark, M.E.; Baldo, L.; Russell, J.A.; Werren, J.H.; Jaenike, J. The mushroom habitat as an ecological arena for global exchange of Wolbachia. Mol. Ecol. 2010, 19, 1940–1952. [Google Scholar]
  25. Hiroki, M.; Tagami, Y.; Miura, K.; Kato, Y. Multiple infection with Wolbachia inducing different reproductive manipulations in the butterfly Eurema hecabe. Proc. R. Soc. Lond. B 2004, 271, 1751–1755. [Google Scholar] [CrossRef]
  26. Narita, S.; Kageyama, D.; Nomura, M.; Fukatsu, T. Unexpected mechanism of symbiont-induced reversal of insect sex: Feminizing Wolbachia continuously acts on the butterfly Eurema hecabe during larval development. Appl. Environ. Microbiol. 2007, 73, 4332–4341. [Google Scholar]
  27. Narita, S.; Kageyama, D.; Hiroki, M.; Sanpei, T.; Hashimoto, S.; Kamitoh, T.; Kato, Y. Wolbachia-induced feminisation newly found in Eurema hecabe, a sibling species of Eurema mandarina (Lepidoptera: Pieridae). Ecol. Entomol. 2011, 36, 309–317. [Google Scholar] [CrossRef]
  28. Negri, I.; Pellecchia, M.; Mazzoglio, P.J.; Patetta, A.; Alma, A. Feminizing Wolbachia in Zyginidia pullula (Insecta, Hemiptera), a leafhopper with an XX/X0 sex-determination system. Proc. R. Soc. Lond. B 2006, 273, 2409–2416. [Google Scholar]
  29. Negri, I.; Franchini, A.; Gonella, E.; Daffonchio, D.; Mazzoglio, P.J.; Mandrioli, M.; Alma, A. Unravelling the Wolbachia evolutionary role: The reprogramming of the host genomic imprinting. Proc. R. Soc. Lond. B 2009, 276, 2485–2491. [Google Scholar]
  30. Legrand, J.J.; Legrand-Hamelin, E.; Juchault, P. Sex determination in Crustacea. Biol. Rev. 1987, 62, 439–470. [Google Scholar] [CrossRef]
  31. Rigaud, T. Inherited microorganisms and sex determination of arthropod hosts. In Influential Passengers; O’Neill, S.L., Hoffmann, A.A., Werren, J.H., Eds.; Oxford University Press: Oxford, UK, 1997; pp. 81–101. [Google Scholar]
  32. Thierry, R.; Juchault, P.; Mocquard, J.P. The evolution of sex determination in isopod crustaceans. BioEssays 1997, 19, 409–416. [Google Scholar] [CrossRef]
  33. Juchault, P.; Mocquard, J.P. Transfer of a parasitic sex factor to the nuclear genome of the host: A hypothesis on the evolution of sex-determining mechanisms in the terrestrial Isopod Armadillidium vulgare Latr. J. Evol. Biol. 1993, 6, 511–528. [Google Scholar]
  34. Rigaud, T.; Juchault, P. Genetic control of the vertical transmission of a cytoplasmic sex factor in Armadillidium vulgare Latr. (Crustacea, Oniscidea). Heredity 1992, 68, 47–52. [Google Scholar] [CrossRef]
  35. Rigaud, T.; Juchault, P. Conflict between feminizing sex ratio distorters and an autosomal masculinizing gene in the terrestrial isopod Armadillidium vulgare Latr. Genetics 1993, 133, 247–252. [Google Scholar]
  36. Martin, G.; Sorokine, O.; Moniatte, M.; Bulet, P.; Hetru, C.; Van Dorsselaer, A. The structure of a glycosylated protein hormone responsible for sex determination in the isopod, Armadillidium vulgare. Eur. J. Biochem. 1999, 262, 727–736. [Google Scholar] [CrossRef]
  37. Okuno, A.; Hasegawa, Y.; Ohira, T.; Katakura, Y.; Nagasawa, H. Characterization and cDNA cloning of androgenic gland hormone of the terrestrial isopod Armadillidium vulgare. Biochem. Biophys. Res. Comm. 1999, 264, 419–423. [Google Scholar] [CrossRef]
  38. Helle, W.; Bolland, H.R.; Heitmans, W.R.B. Chromosomes and types of parthenogenesis in the false spider mites (Acari: Tenuipalpidae). Genetica 1980, 54, 45–50. [Google Scholar] [CrossRef]
  39. Weeks, A.R.; Marec, F.; Breeuwer, J.A.J. A mite species that consists entirely of haploid females. Science 2001, 292, 2479–2482. [Google Scholar]
  40. Weeks, A.R.; Breeuwer, J.A.J. Wolbachia-induced parthenogenesis in a genus of phytophagous mites. Proc. R. Soc. Lond. B 2001, 268, 2245–2251. [Google Scholar] [CrossRef]
  41. Zchori-Fein, E.; Perlman, S.J.; Kelly, S.E.; Katzir, N.; Hunter, M.S. Characterization of a “Bacteroidetes” symbiont in Encarsia wasps (Hymenoptera: Aphelinidae): Proposal of “Candidatus Cardinium hertigii.”. Int. J. Syst. Evol. Microbiol. 2004, 54, 961. [Google Scholar] [CrossRef]
  42. Chigira, A.; Miura, K. Detection of “Candidatus Cardinium” bacteria from the haploid host Brevipalpus californicus (Acari: Tenuipalpidae) and effect on the host. Exp. Appl. Acarol. 2005, 37, 107–116. [Google Scholar] [CrossRef]
  43. Groot, T.V.M.; Breeuwer, J.A.J. Cardinium symbionts induce haploid thelytoky in most clones of three closely related Brevipalpus species. Exp. Appl. Acarol. 2006, 39, 257–271. [Google Scholar] [CrossRef]
  44. Giorgini, M.; Monti, M.M.; Caprio, E.; Stouthamer, R.; Hunter, M.S. Feminization and the collapse of haplodiploidy in an asexual parasitoid wasp harboring the bacterial symbiont Cardinium. Heredity 2009, 102, 365–371. [Google Scholar] [CrossRef]
  45. Traut, W. Zur Geschlechtsbestimmung bei Gammarus duebeni und verwandten Arten. Z. wiss. Zool. 1962, 167, 1–72. [Google Scholar]
  46. Bulnheim, H.P.; Vávra, J. Infection by the Microsporidian Octosporea effeminans sp. n., and its sex determining influence in the amphipod Gammarus duebeni. J. Parasitol. 1968, 54, 241–248. [Google Scholar] [CrossRef]
  47. Dunn, A.M.; Rigaud, T. Horizontal transfer of parasitic sex ratio distorters between crustacean hosts. Parasitology 1998, 117, 15–19. [Google Scholar] [CrossRef]
  48. Terry, R.S.; Smith, J.E.; Dunn, A.M. Impact of a Novel, Feminising Microsporidium on its Crustacean Host. J. Euk. Microbiol. 1998, 45, 497–501. [Google Scholar] [CrossRef]
  49. Dunn, A.M.; Hatcher, M.J.; Terry, R.S.; Tofts, C. Evolutionary ecology of vertically transmitted parasites: Transovarial transmission of a microsporidian sex ratio distorter in Gammarus duebeni. Parasitology 1995, 111, S91–S109. [Google Scholar] [CrossRef]
  50. Rodgers-Gray, T.P.; Smith, J.E.; Ashcroft, A.E.; Isaac, R.E.; Dunn, A.M. Mechanisms of parasite-induced sex reversal in Gammarus duebeni. Int. J. Parasitol. 2004, 34, 747–753. [Google Scholar] [CrossRef]
  51. Dunn, A.; Adams, J.; Smith, J. Transovarial transmission and sex-ratio distortion by a microsporidian parasite in a shrimp. J. Invertebr. Pathol. 1993, 61, 248–252. [Google Scholar] [CrossRef]
  52. Terry, R.S.; Smith, J.E.; Sharpe, R.G.; Rigaud, T.; Littlewood, D.T.J.; Ironside, J.E.; Rollinson, D.; Bouchon, D.; MacNeil, C.; Dick, J.T.; Dunn, A.M. Widespread vertical transmission and associated host sex-ratio distortion within the eukaryotic phylum Microspora. Proc. R. Soc. Lond. B 2004, 271, 1783–1789. [Google Scholar]
  53. Vance, S.A. Morphological and Behavioural sex reversal in mermithid-infected mayflies. Proc. R. Soc. Lond. B 1996, 263, 907–912. [Google Scholar] [CrossRef]
  54. Baudoin, M. Host castration as a parasitic strategy. Evolution 1975, 29, 335–352. [Google Scholar] [CrossRef]
  55. Kathirithamby, J. The effects of stylopisation on the sexual development of Javesella dubia (Kirschbaum) (Homoptera: Delphacidae). Biol. J. Linn. Soc. 1978, 10, 163–179. [Google Scholar] [CrossRef]
  56. Juchault, P.; Louis, C.; Martin, G.; Noulin, G. Masculinization of female isopods (Crustacea) correlated with non-Mendelian inheritance of cytoplasmic viruses. Proc. Natl. Acad. Sci. USA 1991, 88, 10460–10464. [Google Scholar]
  57. Hurst, G.D.; Jiggins, F.M. Male-killing bacteria in insects: Mechanisms, incidence, and implications. Emerg. Inf. Dis. 2000, 6, 329–336. [Google Scholar] [CrossRef]
  58. Hurst, L.D. The incidences and evolution of cytoplasmic male killers. Proc. R. Soc. Lond. B 1991, 244, 91. [Google Scholar] [CrossRef]
  59. Nakanishi, K.; Hoshino, M.; Nakai, M.; Kunimi, Y. Novel RNA sequences associated with late male killing in Homona magnanima. Proc. R. Soc. Lond. B 2008, 275, 1249–1254. [Google Scholar] [CrossRef]
  60. Gilfillan, G.D.; Dahlsveen, I.K.; Becker, P.B. Lifting a chromosome: Dosage compensation in Drosophila melanogaster. FEBS Lett. 2004, 567, 8–14. [Google Scholar] [CrossRef]
  61. Veneti, Z.; Bentley, J.K.; Koana, T.; Braig, H.R.; Hurst, G.D.D. A functional dosage compensation complex required for male killing in Drosophila. Science 2005, 307, 1461–1463. [Google Scholar]
  62. Kageyama, D.; Ohno, S.; Hoshizaki, S.; Ishikawa, Y. Sexual mosaics induced by tetracycline treatment in the Wolbachia-infected adzuki bean borer, Ostrinia scapulalis. Genome 2003, 46, 983–989. [Google Scholar] [CrossRef]
  63. Kageyama, D.; Traut, W. (2004) Opposite sex-specific effects of Wolbachia and interference with the sex determination of its host Ostrinia scapulalis. Proc. R. Soc. Lond. B 2004, 271, 251–258. [Google Scholar] [CrossRef]
  64. Sakamoto, H.; Kageyama, D.; Hoshizaki, S.; Ishikawa, Y. Sex-specific death in the Asian corn borer moth (Ostrinia furnacalis) infected with Wolbachia occurs across larval development. Genome 2007, 50, 645–652. [Google Scholar] [CrossRef]
  65. Sugimoto, T.N.; Fujii, T.; Kayukawa, T.; Sakamoto, H.; Ishikawa, Y. Expression of a doublesex homologue is altered in sexual mosaics of Ostrinia scapulalis moths infected with Wolbachia. Insect Biochem. Mol. Biol. 2010, 40, 847–854. [Google Scholar] [CrossRef]
  66. Sugimoto, T.N.; Ishikawa, Y. A male-killing Wolbachia carries a feminizing factor and is associated with degradation of the sex-determining system of its host. Biol. Lett. 2012. [Google Scholar]
  67. Kageyama, D., Hoshizaki. Female-biased sex ratio in the Asian corn borer, Ostrinia furnacalis: Evidence for the occurrence of feminizing bacteria in an insect. Heredity 1998, 81, 311–316. [Google Scholar] [CrossRef]
  68. Kageyama, D.; Nishimura, G.; Hoshizaki, S.; Ishikawa, Y. Feminizing Wolbachia in an insect, Ostrinia furnacalis (Lepidoptera: Crambidae). Heredity 2002, 88, 444–449. [Google Scholar] [CrossRef]
  69. Dyson, E.A.; Kamath, M.K.; Hurst, G.D.D. Wolbachia infection associated with all-female broods in Hypolimnas bolina (Lepidoptera: Nymphalidae): Evidence for horizontal transmission of a butterfly male killer. Heredity 2002, 88, 166–171. [Google Scholar] [CrossRef]
  70. Mitsuhashi, W.; Saiki, T.; Wei, W.; Kawakita, H.; Sato, M. Two novel strains of Wolbachia coexisting in both species of mulberry leafhoppers. Insect Mol. Biol. Biol. 2002, 11, 577–584. [Google Scholar] [CrossRef]
  71. Dyson, E.A.; Hurst, G.D.D. Persistence of an extreme sex-ratio bias in a natural population. Proc. Natl. Acad. Sci. USA 2004, 101, 6520–6523. [Google Scholar]
  72. Hornett, E.A.; Charlat, S.; Duplouy, A.M.R.; Davies, N.; Roderick, G.K.; Wedell, N.; Hurst, G.D. Evolution of male-killer suppression in a natural population. PLoS Biol. 2006, 4, e283. [Google Scholar] [CrossRef]
  73. Charlat, S.; Hornett, E.A.; Fullard, J.H.; Davies, N.; Roderick, G.K.; Wedell, N.; Hurst, G.D. Extraordinary flux in sex ratio. Science 2007, 317, 214. [Google Scholar]
  74. Mitsuhashi, W.; Ikeda, H.; Muraji, M. Fifty-year trend towards suppression of Wolbachia-induced male-killing by its butterfly host, Hypolimnas bolina. J. Insect Sci. 2011, 11, 1–15. [Google Scholar]
  75. Hornett, E.A.; Duplouy, A.M.R.; Davies, N.; Roderick, G.K.; Wedell, N.; Hurst, G.D.; Charlat, S. You can’t keep a good parasite down: Evolution of a male-killer suppressor uncovers cytoplasmic incompatibility. Evolution 2008, 62, 1258–1263. [Google Scholar] [CrossRef]
  76. Jaenike, J. Spontaneous emergence of a new Wolbachia phenotype. Evolution 2007, 61, 2244–2252. [Google Scholar] [CrossRef]
  77. Fukatsu, T.; Tsuchida, T.; Nikoh, N.; Koga, R. Spiroplasma symbiont of the pea aphid, Acyrthosiphon pisum (Insecta: Homoptera). Appl. Environ. Microbiol. 2001, 67, 1284–1291. [Google Scholar] [CrossRef]
  78. Tsuchida, T.; Koga, R.; Shibao, H.; Matsumoto, T.; Fukatsu, T. Diversity and geographic distribution of secondary endosymbiotic bacteria in natural populations of the pea aphid, Acyrthosiphon pisum. Mol. Ecol. 2002, 11, 2123–2135. [Google Scholar] [CrossRef]
  79. Simon, J.C.; Boutin, S.; Tsuchida, T.; Koga, R.; Le Gallic, J.F.; Frantz, A.; Outreman, Y.; Fukatsu, T. Facultative symbiont infections affect aphid reproduction. PLoS One 2011, 6, e21831. [Google Scholar]
  80. Elnagdy, S.; Majerus, M.E.N.; Handley, L.-J.L. The value of an egg: Resource reallocation in ladybirds (Coleoptera: Coccinellidae) infected with male-killing bacteria. J. Evol. Biol. 2011, 24, 2164–2172. [Google Scholar]
  81. Hurst, G.D.D.; Majerus, M.E.N. Why do maternally inherited microorganisms kill males? Heredity 1993, 71, 81–95. [Google Scholar] [CrossRef]
  82. Dunn, A.M.; Smith, J.E. Microsporidian life cycles and diversity: The relationship between virulence and transmission. Microb. Inf. 2001, 3, 381–388. [Google Scholar] [CrossRef]
  83. O’Neill, S.L.; Hoffmann, A.A.; Werren, J.H. Influential Passengers; Oxford University Press: Oxford, UK, 1997; p. 214. [Google Scholar]
  84. Bourtzis, K.; Miller, T. Insect Symbiosis; CRC Press: Boca Raton, FL, USA, 2003; p. 368. [Google Scholar]
  85. Kageyama, D.; Anbutsu, H.; Shimada, M.; Fukatsu, T. Spiroplasma infection causes either early or late male killing in Drosophila, depending on maternal host age. Naturwissenschaften 2007, 94, 333–337. [Google Scholar] [CrossRef]
  86. Charlat, S.; Davies, N.; Roderick, G.K.; Hurst, G.D.D. Disrupting the timing of Wolbachia-induced male-killing. Biol. Lett. 2007, 3, 154–156. [Google Scholar] [CrossRef]
  87. Stouthamer, R. Wolbachia-induced parthenogenesis. In Influential Passengers; O’Neill, S.L., Hoffmann, A.A., Werren, J.H., Eds.; Oxford University Press: Oxford, UK, 1997; pp. 102–124. [Google Scholar]
  88. Arakaki, N.; Miyoshi, T.; Noda, H. Wolbachia-mediated parthenogenesis in the predatory thrips Franklinothrips vespiformis (Thysanoptera: Insecta). Proc. R. Soc. Lond. B 2001, 268, 1011–1016. [Google Scholar] [CrossRef]
  89. Koivisto, R.K.K.; Braig, H.R. Microorganisms and parthenogenesis. Biol. J. Linn. Soc. 2003, 79, 43–58. [Google Scholar] [CrossRef]
  90. Zchori-Fein, E.; Roush, R.T.; Hunter, M.S. Male production induced by antibiotic treatment in Encarsia formosa (Hymenoptera: Aphelinidae), an asexual species. Experientia 1992, 48, 102–105. [Google Scholar] [CrossRef]
  91. Pijls, J.W.A.M.; Steenbergen, H.J.V.A.N.; Van, J.J.M. Asexuality cured: The relations and differences between sexual and asexual Apoanagyrus diversicornis. Heredity 1996, 76, 506–513. [Google Scholar] [CrossRef]
  92. Kremer, N.; Charif, D.; Henri, H.; Bataille, M.; Prévost, G.; Kraaijeveld, K.; Vavre, F. A new case of Wolbachia dependence in the genus Asobara: Evidence for parthenogenesis induction in Asobara japonica. Heredity 2009, 103, 248–256. [Google Scholar] [CrossRef]
  93. Stouthamer, R.; Kazmer, D.J. Cytogenetics of microbe-associated parthenogenesis and its consequences for gene flow in Trichogramma wasps. Heredity 1994, 73, 317–327. [Google Scholar] [CrossRef]
  94. Pannebakker, B.A.; Pijnacker, L.P.; Zwaan, B.J.; Beukeboom, L.W. Cytology of Wolbachia-induced parthenogenesis in Leptopilina clavipes (Hymenoptera: Figitidae). Genome 2004, 47, 299–303. [Google Scholar] [CrossRef]
  95. Gottlieb, Y.; Zchori-Fein, E.; Werren, J.H.; Karr, T.L. Diploidy restoration in Wolbachia-infected Muscidifurax uniraptor (Hymenoptera: Pteromalidae). J. Invertebr. Pathol. 2002, 81, 166–174. [Google Scholar] [CrossRef]
  96. Adachi-Hagimori, T.; Miura, K.; Stouthamer, R. A new cytogenetic mechanism for bacterial endosymbiont-induced parthenogenesis in Hymenoptera. Proc. R. Soc. Lond. B 2008, 275, 2667–2673. [Google Scholar] [CrossRef]
  97. Verma, S.; Ruttner, F. Cytological analysis of thelytokous parthenogenesis in the Cape honeybee (Apis mellifera capensis Escholtz). Apidologie 1983, 17, 47–58. [Google Scholar]
  98. Pearcy, M.; Hardy, O.; Aron, S. Thelytokous parthenogenesis and its consequences on inbreeding in an ant. Heredity 2006, 96, 377–382. [Google Scholar] [CrossRef]
  99. Bourtzis, K.; Braig, H.; Karr, T. Cytoplasmic incompatibility. In Insect Symbiosis; Bourtzis, K., Miller, T., Eds.; CRC Press: Boca Raton, FL, USA, 2003; pp. 217–246. [Google Scholar]
  100. Poinsot, D.; Charlat, S.; Merçot, H. On the mechanism of Wolbachia-induced cytoplasmic incompatibility: Confronting the models with the facts. BioEssays 2003, 25, 259–265. [Google Scholar] [CrossRef]
  101. Tram, U.; Fredrick, K.; Werren, J.; Sullivan, W. Paternal chromosome segregation during the first mitotic division determines Wolbachia-induced cytoplasmic incompatibility phenotype. J. Cell Sci. 2006, 119, 3655–3663. [Google Scholar] [CrossRef]
  102. Taylor, M.J.; Bandi, C.; Hoerauf, A. Wolbachia bacterial endosymbionts of filarial nematodes. Adv. Parasitol. 2005, 60, 245–284. [Google Scholar] [CrossRef]
  103. Hoerauf, A.; Mand, S.; Fischer, K.; Kruppa, T.; Marfo-Debrekyei, Y.; Debrah, A.Y.; Pfarr, K.M.; Adjei, O.; Büttner, D.W. Doxycycline as a novel strategy against bancroftian filariasis-depletion of Wolbachia endosymbionts from Wuchereria bancrofti and stop of microfilaria production. Med. Microbiol. Immunol. 2003, 192, 211–216. [Google Scholar]
  104. Hosokawa, T.; Koga, R.; Kikuchi, Y.; Meng, X.-Y.; Fukatsu, T. Wolbachia as a bacteriocyte-associated nutritional mutualist. Proc. Natl. Acad. Sci. USA 2010, 107, 769–774. [Google Scholar]
  105. Dedeine, F.; Vavre, F.; Fleury, F.; Loppin, B.; Hochberg, M.E.; Bouletreau, M. Removing symbiotic Wolbachia bacteria specifically inhibits oogenesis in a parasitic wasp. Proc. Natl. Acad. Sci. USA 2001, 98, 6247–6252. [Google Scholar]
  106. Starr, D.J.; Cline, T.W. A host-parasite interaction rescues Drosophila oogenesis defects. Nature 2002, 418, 76–79. [Google Scholar]
  107. Hedges, L.M.; Brownlie, J.C.; O’Neill, S.L.; Johnson, K.N. Wolbachia and virus protection in insects. Science 2008, 322, 702. [Google Scholar]
  108. Osborne, S.E.; Leong, Y.S.; O’Neill, S.L.; Johnson, K.N. Variation in antiviral protection mediated by different Wolbachia strains in Drosophila simulans. PLoS Pathog. 2009, 5, e1000656. [Google Scholar] [CrossRef]
  109. Teixeira, L.; Ferreira, A.; Ashburner, M. The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster. PLoS Biol. 2008, 6, e2. [Google Scholar]
  110. Glaser, R.L.; Meola, M.A. The native Wolbachia endosymbionts of Drosophila melanogaster and Culex quinquefasciatus increase host resistance to West Nile virus infection. PLoS One 2010, 5, e11977. [Google Scholar] [CrossRef]
  111. Panteleev, D.I.; Goriacheva, I.I.; Andrianov, B.V.; Reznik, N.L.; Lazebnyĭ, O.E.; Kulikov, A.M. The endosymbiotic bacterium Wolbachia enhances the nonspecific resistance to insect pathogens and alters behavior of Drosophila melanogaster. Genetika 2007, 43, 1277–1280. [Google Scholar]
  112. Kambris, Z.; Cook, P.; Phuc, H.; Sinkins, S. Immune activation by life-shortening Wolbachia and reduced filarial competence in mosquitoes. Science 2009, 326, 134–136. [Google Scholar]
  113. Bian, G.; Xu, Y.; Lu, P.; Xie, Y.; Xi, Z. The endosymbiotic bacterium Wolbachia induces resistance to dengue virus in Aedes aegypti. PLoS Pathog. 2010, 6, e1000833. [Google Scholar] [CrossRef]
  114. Moreira, L.A.; Iturbe-Ormaetxe, I.; Jeffery, J.A.; Lu, G.; Pyke, A.T.; Hedges, L.M.; Rocha, B.C.; Hall-Mendelin, S.; Day, A.; Riegler, M.; Hugo, L.E.; Johnson, K.N.; Kay, B.H.; McGraw, E.A.; van den Hurk, A.F.; Ryan, P.A.; O'Neill, S.L. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell 2009, 139, 1268–1278. [Google Scholar]
  115. Kambris, Z.; Blagborough, A.M.; Pinto, S.B.; Blagrove, M.S.C.; Godfray, H.C.J.; Sinden, R.E.; Sinkins, S.P. Wolbachia stimulates immune gene expression and inhibits plasmodium development in Anopheles gambiae. PLoS Pathog. 2010, 6, e1001143. [Google Scholar] [CrossRef]
  116. Hoffmann, A.A.; Montgomery, B.L.; Popovici, J.; Iturbe-Ormaetxe, I.; Johnson, P.H.; Muzzi, F.; Greenfield, M.; Durkan, M.; Leong, Y.S.; Dong, Y.; Cook, H.; Axford, J.; Callahan, A.G.; Kenny, N.; Omodei, C.; McGraw, E.A.; Ryan, P.A.; Ritchie, S.A.; Turelli, M.; O'Neill, S.L. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature 2011, 476, 454–457. [Google Scholar]
  117. Walker, T.; Johnson, P.H.; Moreira, L.A.; Iturbe-Ormaetxe, I.; Frentiu, F.D.; McMeniman, C.J.; Leong, Y.S.; Dong, Y.; Axford, J.; Kriesner, P.; Lloyd, A.L.; Ritchie, S.A.; O'Neill, S.L.; Hoffmann, A.A. The wMel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations. Nature 2011, 476, 450–453. [Google Scholar]
  118. Iturbe-Ormaetxe, I.; Walker, T.; O’ Neill, S.L. Wolbachia and the biological control of mosquito-borne disease. EMBO Rep. 2011, 12, 508–518. [Google Scholar] [CrossRef]
  119. Jaenike, J.; Unckless, R.; Cockburn, S.N.; Boelio, L.M.; Perlman, S.J. Adaptation via symbiosis: Recent spread of a Drosophila defensive symbiont. Science 2010, 329, 212–215. [Google Scholar]
  120. Rigaud, T.; Juchault, P.; Mocquard, J.-P. The evolution of sex determination in isopod crustaceans. BioEssays 1997, 19, 409–416. [Google Scholar] [CrossRef]
  121. Hashiyama, K.; Hayashi, Y.; Kobayashi, S. Drosophila Sex lethal gene initiates female development in germline progenitors. Science 2011, 333, 885–888. [Google Scholar]
  122. Clark, M.E.; Heath, B.D.; Anderson, C.L., Karr. Induced paternal effects mimic cytoplasmic incompatibility in Drosophila. Genetics 2006, 173, 727. [Google Scholar] [CrossRef]
  123. Xi, Z.; Gavotte, L.; Xie, Y.; Dobson, S. Genome-wide analysis of the interaction between the endosymbiotic bacterium Wolbachia and its Drosophila host. BMC Genomics 2008, 9, 1. [Google Scholar] [CrossRef]
  124. Nakamura, Y.; Gotoh, T.; Imanishi, S.; Mita, K.; Kurtti, T.J.; Noda, H. Differentially expressed genes in silkworm cell cultures in response to infection by Wolbachia and Cardinium endosymbionts. Insect Mol. Biol. 2011, 20, 279–289. [Google Scholar] [CrossRef]
  125. Hughes, G.L.; Ren, X.; Ramirez, J.L.; Sakamoto, J.M.; Bailey, J.A.; Jedlicka, A.E.; Rasgon, J.L. Wolbachia infections in Anopheles gambiae cells: Transcriptomic characterization of a novel host-symbiont interaction. PLoS Pathog. 2011, 7, e1001296. [Google Scholar] [CrossRef]
  126. Hatcher, M.J.; Taneyhill, D.E.; Dunn, A.M.; Tofts, C. Population dynamics under parasitic sex ratio distortion. Theor. Pop. Biol. 1999, 56, 11–28. [Google Scholar] [CrossRef]
  127. Saccheri, I.; Kuussaari, M.; Kankare, M.; Vikman, P.; Hanski, I. Inbreeding and extinction in a butterfly metapopulation. Nature 1998, 45, 1996–1999. [Google Scholar]
  128. Jiggins, F.M.; Hurst, G.D.; Majerus, M.E. Sex-ratio-distorting Wolbachia causes sex-role reversal in its butterfly host. Proc. R. Soc. Lond. B 2000, 267, 69–73. [Google Scholar] [CrossRef]
  129. Narita, S.; Nomura, M.; Kageyama, D. Naturally occurring single and double infection with Wolbachia strains in the butterfly Eurema hecabe: Transmission efficiencies and population density dynamics of each Wolbachia strain. FEMS Microbiol. Ecol. 2007, 61, 235–245. [Google Scholar] [CrossRef]
  130. Martin, G.; Juchault, P.; Legrand, J. Mise en évidence d’un microorganisme intracytoplasmique symbiote de l’oniscoïde Armadillidium vulgare Latr., dont la présence accompagne l’intersexualité ou la féminisation totale des mâles génétiques de la lignée thélygène. C. R. Acad. Sci. 1973, 276, 2313–2316. [Google Scholar]
  131. Rigaud, T.; Souty-Grosset, C.; Raimond, R.; Mocquard, J.P.; Juchault, P. Feminizing endocytobiosis in the terrestrial crustacean Armadillidium vulgare Latr. (Isopoda): Recent acquisitions. Endocytobiosis Cell Res. 1991, 7, 259–273. [Google Scholar]
  132. Rousset, F.; Bouchon, D.; Pintureau, B.; Juchault, P.; Solignac, M. Wolbachia endosymbionts responsible for various alterations of sexuality in arthropods. Proc. R. Soc. Lond. B 1992, 250, 91–98. [Google Scholar] [CrossRef]
  133. Juchault, P.; Legrand, J.J. Analyse génétique et physiologique de la détermination du sexe dans une population du crustacé Oniscoïde—Armadillidium nasatum Budde-Lund. Arch. Zool. Exp. Gén. 1979, 120, 25–43. [Google Scholar]
  134. Juchault, P.; Legrand, J.J. Sex determination and monogeny in terrestrial isopods Armadillidium vulgare (Latreille, 1804) and Armadillidium nasatum (Budde-Lund, 1885). Monogr. Monit. Zool. Italian. 1989, 4, 359–375. [Google Scholar]
  135. Juchault, P.; Frelon, M.; Bouchon, D.; Rigaud, T. New evidence for feminizing bacteria in terrestrial isopods: Evolutionary implications. C. R. Acad. Sci. 1994, 317, 225–230. [Google Scholar]
  136. Rigaud, T.; Antoine, D.; Marcadé, I.; Juchault, P. The effect of temperature on sex ratio in the isopod Porcellionides pruinosus: Environmental sex determination or a by-product of cytoplasmic sex determination? Evol. Ecol. 1997, 11, 205–215. [Google Scholar] [CrossRef]
  137. Martin, G.; Gruppe, S.G.; Laulier, M.; Bouchon, D.; Rigurd, D.; Juchault, P. (1994) Evidence for Wolbachia spp. in the estuarine isopod Sphaeroma rugicauda (Crustacea): A likely cytoplasmic sex ratio distorter. Endocytobiosis Cell Res. 1994, 10, 215–225. [Google Scholar]
  138. Bulnheim, H.-P. Interaction between genetic, external and parasitic factors in sex determination of the crustacean amphipod Gammarus duebeni. Helgoländer Wissenschaftliche Meeresuntersuchungen 1978, 31, 1–33. [Google Scholar] [CrossRef]
  139. Hurst, G.D.D.; Jiggins, F.M.; Schulenburg, J.H.; Bertrand, D.; West, S.A.; et al. Male-killing Wolbachia in two species of insect. Proc. R. Soc. Lond. B 1999, 266, 735–740. [Google Scholar]
  140. Fialho, R.F.; Stevens, L. Male-killing Wolbachia in a flour beetle. Proc. R. Soc. Lond. B 2000, 267, 1469–1473. [Google Scholar] [CrossRef]
  141. Hurst, G.D.; Johnson, A.P.; Schulenburg, J.H.; Fuyama, Y. Male-killing Wolbachia in Drosophila: A temperature-sensitive trait with a threshold bacterial density. Genetics 2000, 156, 699–709. [Google Scholar]
  142. Sheeley, S.L.; McAllister, B.F. Mobile male-killer: Similar Wolbachia strains kill males of divergent Drosophila hosts. Heredity 2009, 102, 286–292. [Google Scholar] [CrossRef]
  143. Jaenike, J.; Dyer, K.A.; Reed, L.K. Within-population structure of competition and the dynamics of male-killing Wolbachia. Evol. Ecol. Res. 2003, 5, 1023–1036. [Google Scholar]
  144. Jiggins, F.M.; Hurst, G.D.D.; Dolman, C.E.; Majerus, M.E.N. High-prevalence male-killing Wolbachia in the butterfly Acraea encedana. J. Evol. Biol. 2000, 13, 495–501. [Google Scholar]
  145. Kageyama, D.; Nishimura, G.; Ohno, S.; Takanashi, T.; Hoshizaki, S.; Ishikawa, Y. Wolbachia infection and an all-female trait in Ostrinia orientalis and Ostrinia zaguliaevi. Entomol. Exp. Appl. 2004, 111, 79–83. [Google Scholar] [CrossRef]
  146. Kageyama, D.; Nishimura, G.; Hoshizaki, S.; Ishikawa, Y. Two kinds of sex ratio distorters in a moth, Ostrinia scapulalis. Genome 2003, 46, 974–982. [Google Scholar] [CrossRef]
  147. Zeh, D.W.; Zeh, J.A.; Bonilla, M.M. Wolbachia, sex ratio bias and apparent male killing in the harlequin beetle riding pseudoscorpion. Heredity 2005, 95, 41–49. [Google Scholar] [CrossRef]
  148. Hurst, G.D.; Schulenburg, J.H.; Majerus, T.M.; Bertrand, D.; Zakharov, I.A.; Baungaard, J.; Völkl, W.; Stouthamer, R.; Majerus, M.E. Invasion of one insect species, Adalia bipunctata, by two different male-killing bacteria. Insect Mol. Biol. 1999, 8, 133–139. [Google Scholar]
  149. Majerus, M.E.; Schulenburg, J.H.; Zakharov, I.A. Multiple causes of male-killing in a single sample of the two-spot ladybird, Adalia bipunctata (Coleoptera: Coccinellidae) from Moscow. Heredity 2000, 84, 605–609. [Google Scholar] [CrossRef]
  150. Tinsley, M.C.; Majerus, M.E.N. A new male-killing parasitism: Spiroplasma bacteria infect the ladybird beetle Anisosticta novemdecimpunctata (Coleoptera: Coccinellidae). Parasitology 2006, 132, 757–765. [Google Scholar] [CrossRef]
  151. Matsuka, M.; Hashi, H.; Okada, I. Abnormal sex ratio found in the lady beetle Harmonia axyridis Pallas (Coleoptera: Coccinellidae). Appl. Entomol. Zool. 1975, 10, 84–89. [Google Scholar]
  152. Majerus, T.M.; Schulenburg, J.H.; Majerus, M.E.; Hurst, G.D. Molecular identification of a male-killing agent in the ladybird Harmonia axyridis (Pallas) (Coleoptera: Coccinellidae). Insect Mol. Biol. 1999, 8, 551–555. [Google Scholar] [CrossRef]
  153. Niijima, K.; Nakajima, K. Abnormal sex-ratio in Menochilus sexmaculatus (Fabricius). Bull. of the Faculty of Agriculture-Tamagawa Univ (Japan) 1981, 21, 59–67. [Google Scholar]
  154. Jiggins, F.M.; Hurst, G.D.; Jiggins, C.D.; Schulenburg, J.H.; Majerus, M.E. (2000) The butterfly Danaus chrysippus is infected by a male-killing Spiroplasma bacterium. Parasitology 2000, 120, 439–446. [Google Scholar] [CrossRef]
  155. Tabata, J.; Hattori, Y.; Sakamoto, H.; Yukuhiro, F.; Fujii, T.; Kugimiya, S.; Mochizuki, A.; Ishikawa, Y.; Kageyama, D. Male killing and incomplete inheritance of a novel spiroplasma in the moth Ostrinia zaguliaevi. Microb. Ecol. 2011, 61, 254–263. [Google Scholar] [CrossRef]
  156. Bentley, J.K.; Veneti, Z.; Heraty, J.; Hurst, G.D.D. The pathology of embryo death caused by the male-killing Spiroplasma bacterium in Drosophila nebulosa. BMC Biol. 2007, 5, 9. [Google Scholar] [CrossRef]
  157. Montenegro, H.; Hatadani, L.M.; Medeiros, H.F.; Klaczko, L.B. Male killing in three species of the tripunctata radiation of Drosophila (Diptera: Drosophilidae). J. Zool. Syst. Evol. Res. 2006, 44, 130–135. [Google Scholar] [CrossRef]
  158. Pool, J.E.; Wong, A.; Aquadro, C.F. Finding of male-killing Spiroplasma infecting Drosophila melanogaster in Africa implies transatlantic migration of this endosymbiont. Heredity 2006, 97, 27–32. [Google Scholar] [CrossRef]
  159. Williamson, D.L.; Sakaguchi, B.; Hackett, K.J.; Whitcomb, R.F.; Tully, J.G.; Carle, P.; Bové, J.M.; Adams, J.R.; Konai, M.; Henegar, R.B. Spiroplasma poulsonii sp. nov., a new species associated with male-lethality in Drosophila willistoni, a neotropical species of fruit fly. Int. J. Syst. Bacteriol. 1999, 49, 611–618. [Google Scholar] [CrossRef]
  160. Hurst, G.D.D.; Majerus, M.E.N.; Walker, L.E. The importance of cytoplasmic male killing elements in natural populations of the two spot ladybird, Adalia bipunctata (Linnaeus) (Goleoptera: Goccinellidae). Biol. J. Linn. Soc. 1993, 49, 195–202. [Google Scholar]
  161. Werren, J.H.; Hurst, G.D.; Zhang, W.; Breeuwer, J.A.; Stouthamer, R.; Majerus, M.E. Rickettsial relative associated with male killing in the ladybird beetle (Adalia bipunctata). J. Bacteriol. 1994, 176, 388–394. [Google Scholar]
  162. Schulenburg, J.H.; Habig, M.; Sloggett, J.J.; Webberley, K.M.; Bertrand, D.; Hurst, G.D.; Majerus, M.E. Incidence of male-killing Rickettsia spp. (alpha-proteobacteria) in the ten-spot ladybird beetle Adalia decempunctata L. (Coleoptera: Coccinellidae). Appl. Environ. Microbiol. 2001, 67, 270–277. [Google Scholar]
  163. Lawson, E.T.; Mousseau, T.A.; Klaper, R.; Hunter, M.D.; Werren, J.H. Rickettsia associated with male-killing in a buprestid beetle. Heredity 2001, 86, 497–505. [Google Scholar] [CrossRef]
  164. Majerus, T.M.; Majerus, M.E. Discovery and identification of a male-killing agent in the Japanese ladybird Propylea japonica (Coleoptera: Coccinellidae). BMC Evol. Biol. 2010, 10, 37. [Google Scholar] [CrossRef]
  165. Hurst, G.D.; Bandi, C.; Sacchi, L.; Cochrane, A.G.; Bertrand, D.; Karaca, I.; Majerus, M.E. Adonia variegata (Coleoptera: Coccinellidae) bears maternally inherited flavobacteria that kill males only. Parasitology 1999, 118, 125–134. [Google Scholar] [CrossRef]
  166. Majerus, T.M.O. The Evolutionary Genetics of Male killing in the Coccinellidae. PhD thesis, Department of Genetics, University of Cambridge, Cambridge, UK, 2001. [Google Scholar]
  167. Majerus, M.E.N.; Majerus, T.M.O. Female-biased sex ratio due to male-killing in the Japanese ladybird Coccinula sinensis. Ecol. Entomol. 2000, 25, 234–238. [Google Scholar] [CrossRef]
  168. Hurst, G.D.D.; Hammarton, T.C.; Bandi, C.; Majerus, T.M.O.; Bertrand, D.; Majerus, M.E.N. The diversity of inherited parasites of insects: The male-killing agent of the ladybird beetle Coleomegilla maculata is a member of the Flavobacteria. Genet. Res. 1997, 70, 1–6. [Google Scholar] [CrossRef]
  169. Werren, J.H.; Skinner, S.W.; Huger, A.M. Male-killing bacteria in a parasitic wasp. Science 1986, 231, 990–992. [Google Scholar]
  170. Ferree, P.M.; Avery, A.; Azpurua, J.; Wilkes, T.; Werren, J.H. A bacterium targets maternally inherited centrosomes to kill males in Nasonia. Curr. Biol. 2008, 18, 1409–1414. [Google Scholar]
  171. Hazard, E.I.; Weiser, J. Spores of Thelohania in adult female Anopheles: Development and transovarial transmission, and redescriptions of T. legeri Hesse and T. obesa Kudo. J. Euk. Microbiol. 1968, 15, 817–823. [Google Scholar] [CrossRef]
  172. Avery, S. Horizontal transmission of (Protozoa: Microsporida) to (Diptera: Culicidae). J. Invertebr. Pathol. 1989, 53, 424–426. [Google Scholar] [CrossRef]
  173. Andreadis, T.G. Life cycle, epizootiology, and horizontal transmission of (Microspora: Amblyosporidae) in a univoltine mosquito. J. Invertebr. Pathol. 1985, 46, 31–46. [Google Scholar] [CrossRef]
  174. Kellen, W.; Chapman, H.; Clark, T.; Lindegren, J. Host-parasite relationships of some from mosquitoes (Nosematidae: Microsporidia). J. Invertebr. Pathol. 1965, 7, 161–166. [Google Scholar] [CrossRef]
  175. Kellen, W.R.; Lipa, J.J. Thelohania californica n. sp. a microsporidian parasite of Culex tarsalis Coquillet. J. Insect. Pathol. 1960, 2, 1–12. [Google Scholar]
  176. Kellen, W.R.; Wills, W. The transovarian transmission of Thelohania californica Kellen and Lipa in Culex tarsalis Coquillet. J. Insect Pathol. 1962, 4, 321–326. [Google Scholar]
  177. Andreadis, T.G.; Hall, D.W. Development, ultrastructure, and mode of transmission of Amblyospora sp. (Microspora) in the mosquito. J. Protozool. 1979, 26, 444–452. [Google Scholar]
  178. Morimoto, S.; Nakai, M.; Ono, A.; Kunimi, Y. Late male-killing phenomenon found in a Japanese population of the oriental tea tortrix, Homona magnanima (Lepidoptera: Tortricidae). Heredity 2001, 87, 435–440. [Google Scholar] [CrossRef]
  179. Zchori-Fein, E.; Faktor, O.; Zeidan, M.; Gottlieb, Y.; Czosnek, H.; Rosen, D. Parthenogenesis-inducing microorganisms in Aphytis (Hymenoptera: Aphelinidae). Insect Mol. Biol. 1995, 4, 173–178. [Google Scholar] [CrossRef]
  180. Gottlieb, Y.; Zchori-Fein, E.; Faktor, O.; Rosen, D. Phylogenetic analysis of parthenogenesis-inducing Wolbachia in the genus Aphytis (Hymenoptera: Aphelinidae). Insect Mol. Biol. 1998, 7, 393–396. [Google Scholar]
  181. Zchori-Fein, E.; Rosen, D.; Roush, R. Microorganisms associated with thelytoky in Aphytis lingnansensis compere (Hymenoptera: Aphelinidae). Int. J. Insect. Morphol. Embryol. 1994, 23, 169–172. [Google Scholar] [CrossRef]
  182. Schilthuizen, M.; Stouthamer, R. Distribution of Wolbachia among the guild associated with the parthenogenetic gall wasp Diplolepis rosae. Heredity 1998, 81, 270–274. [Google Scholar] [CrossRef]
  183. Barro, P.J.; Hart, P.J. Antibiotic curing of parthenogenesis in Eretmocerus mundus (Australian parthenogenic form). Entomol. Exp. Appl. 2001, 99, 225–230. [Google Scholar]
  184. Arakaki, N.; Oishi, T.; Noda, H. Parthenogenesis Induced by Wolbachia in Gronotoma micromorpha (Hymenoptera: Eucoilidae). Entomol. Sci. 2001, 4, 9–15. [Google Scholar]
  185. Stouthamer, R.; Breeuwert, J.A.; Luck, R.F.; Werren, J.H. Molecular identification of microorganisms associated with parthenogenesis. Nature 1993, 361, 66–68. [Google Scholar] [CrossRef]
  186. Stouthamer, R.; Lukoe, S.; Mak, F. Influence of parthenogenesis Wolbachia on host fitness. Nor. J. Agric. Sci. (Suppl.) 1994, 6, 117–122. [Google Scholar]
  187. Arakaki, N.; Noda, H.; Yamagishi, K. Wolbachia-induced parthenogenesis in the egg parasitoid Telenomus nawai. Entomol. Exp. Appl. 2000, 96, 177–184. [Google Scholar]
  188. Stouthamer, R.; Werren, J.H. Microbes associated with parthenogenesis in wasps of the genus Trichogramma. J. Invertebr. Pathol. 1993, 61, 6–9. [Google Scholar] [CrossRef]
  189. Stouthamer, R.; Luck, R.F.; Hamilton, W.D. Antibiotics cause parthenogenetic Trichogramma (Hymenoptera/Trichogrammatidae) to revert to sex. Proc. Natl. Acad. Sci. USA 1990, 87, 2424–2427. [Google Scholar]
  190. Schilthuizen, M.; Honda, J.; Stouthamer, R. Parthenogenesis-inducing Wolbachia in Trichogramma kaykai (Hymenoptera: Trichogrammatidae) originates from a single infection. Ann. Entomol. Soc. Am. 1998, 91, 410–414. [Google Scholar]
  191. Zchori-Fein, E.; Gottlieb, Y.; Kelly, S.E.; Brown, J.K.; Wilson, J.M.; Karr, T.L.; Hunter, M.S. A newly discovered bacterium associated with parthenogenesis and a change in host selection behavior in parasitoid wasps. Proc. Natl. Acad. Sci. USA 2001, 98, 12555–12560. [Google Scholar]
  192. Giorgini, M. Induction of males in thelytokous populations of Encarsia meritoria and Encarsia protransvena: A systematic tool. BioControl 2001, 46, 427–438. [Google Scholar] [CrossRef]
  193. Hagimori, T.; Abe, Y.; Date, S.; Miura, K. The first finding of a Rickettsia bacterium associated with parthenogenesis induction among insects. Curr. Microbiol. 2006, 52, 97–101. [Google Scholar]
  194. Giorgini, M.; Bernardo, U.; Monti, M.M.; Nappo, A.G.; Gebiola, M. Rickettsia symbionts cause parthenogenetic reproduction in the parasitoid wasp Pnigalio soemius (Hymenoptera: Eulophidae). Appl. Environ. Microbiol. 2010, 76, 2589–2599. [Google Scholar] [CrossRef]
  195. Kondo, N.; Ijichi, N.; Shimada, M.; Fukatsu, T. Prevailing triple infection with Wolbachia in Callosobruchus chinensis (Coleoptera: Bruchidae). Mol. Ecol. 2002, 11, 167–180. [Google Scholar] [CrossRef]
  196. Keller, G.P.; Windsor, D.M.; Saucedo, J.M.; Werren, J.H. Reproductive effects and geographical distributions of two Wolbachia strains infecting the Neotropical beetle, Chelymorpha alternans Boh. (Chrysomelidae, Cassidinae). Mol. Ecol. 2004, 13, 2405–2420. [Google Scholar] [CrossRef]
  197. Giordano, R.; Jackson, J.J.; Robertson, H.M. The role of Wolbachia bacteria in reproductive incompatibilities and hybrid zones of Diabrotica beetles and Gryllus crickets. Proc. Natl. Acad. Sci. USA 1997, 94, 11439–11444. [Google Scholar] [CrossRef]
  198. Blickenstaff, C.C. Partial Inter sterility of Eastern and Western U.S. Strains of the Alfalfa Weevil. Ann. Entomol. Soc. Am. 1965, 58, 523–526. [Google Scholar]
  199. Hsiao, C.; Hsiao, T. Rickettsia as the cause of cytoplasmic incompatibility in the alfalfa weevil, Hypera postica. J. Insect Pathol. 1985, 45, 244–246. [Google Scholar] [CrossRef]
  200. O’Neill, S.L.; Giordano, R.; Colbert, A.M.; Karr, T.L.; Robertson, H.M. 16S rRNA phylogenetic analysis of the bacterial endosymbionts associated with cytoplasmic incompatibility in insects. Proc. Natl. Acad. Sci. USA 1992, 89, 2699–2702. [Google Scholar]
  201. Fialho, R.; Stevens, L. Wolbachia Infections in the Flour Beetle Tribolium confusum: Evidence for a Common Incompatibility Type across Strains. J. Insect Pathol. 1996, 67, 195–197. [Google Scholar] [CrossRef]
  202. Fialho, R.F.; Stevens, L. Molecular evidence for single Wolbachia infections among geographic strains of the flour beetle Tribolium confusum. Proc. R. Soc. Lond. B 1997, 264, 1065–1068. [Google Scholar]
  203. Macdonald, W. Mosquito genetics in relation to filarial infections. Symposia of the British Society of Parasitology 1976, 14, 1–24. [Google Scholar]
  204. Wright, J.D.; Barr, A.R. The ultrastructure and symbiotic relationships of Wolbachia of mosquitoes of the Aedes scutellaris group. J. Ultra. Res. 1980, 72, 52–64. [Google Scholar] [CrossRef]
  205. Wright, J.; Wang, B. Observations on Wolbachiae in mosquitoes. J. Insect Pathol. 1980, 35, 200–208. [Google Scholar] [CrossRef]
  206. Trpis, M.; Perrone, J.; Reissig, M.; Parker, K. Control of cytoplasmic incompatibility in the Aedes scutellaris complex. J. Heredity 1981, 72, 313–317. [Google Scholar]
  207. Tesfa-Yohannes, T.M.; Rozeboom, L.E. Experimental crossing of Aedes (S.) polynesiensis Marks and A. scutellaris malayensis Colless (Diptera: Culicidae). J. Med. Entomol. 1974, 11, 323–331. [Google Scholar]
  208. Beckett, E.B.; Boothroyd, B.; Macdonald, W.W. A light and electron microscope study of rickettsia-like organisms in the ovaries of mosquitoes of the Aedes scutellaris group. Ann. Trop. Med. Parasitol. 1978, 72, 277–283. [Google Scholar]
  209. Meek, S.R. Occurrence of rickettsia-like symbionts among species of the Aedes scutellaris group (Diptera: Culicidae). Ann. Trop. Med. Parasitol. 1984, 78, 377–381. [Google Scholar]
  210. Meek, S.; Macdonald, W. Crossing relationships among seven members of the group of Aedes scutellaris (Walker)(Diptera: Culicidae). Bull. Entomol. Res. 1984, 74, 65–78. [Google Scholar] [CrossRef]
  211. Smith-White, S.; Woodhill, A. The nature and significance of non-reciprocal fertility in Aedes scutellaris and other mosquitoes. Proc. Linn. Soc. N. S. W. 1954, 79, 163–176. [Google Scholar]
  212. Jamnongluk, W.; Kittayapong, P.; Baisley, K.J.; O’Neill, S.L. Wolbachia infection and expression of cytoplasmic incompatibility in Armigeres subalbatus (Diptera: Culicidae). J. Med. Entomol. 2000, 37, 53–57. [Google Scholar] [CrossRef]
  213. Laven, H. Speciation and evolution in Culex pipiens. In Genetics of insect vectors of disease; Wright, J., Pal, P., Eds.; Elsevier: Amsterdam, The Netherlands, 1967; pp. 251–275. [Google Scholar]
  214. Barr, A.R. Cytoplasmic incompatibility in natural populations of a mosquito, Culex pipiens L. Nature 1980, 283, 71–72. [Google Scholar] [CrossRef]
  215. Magnin, M.; Pasteur, N.; Raymond, M. Multiple incompatibilities within populations of Culex pipiens L. in southern France. Genetica 1987, 74, 125–130. [Google Scholar] [CrossRef]
  216. Irving-Bell, R.J. Cytoplasmic incompatibility within and between Culex molestus and Cx. quinquefasciatus (Diptera: Culicidae). J. Med. Entomol. 1983, 20, 44–48. [Google Scholar]
  217. O’Neill, S.L.; Paterson, H.E. Crossing type variability associated with cytoplasmic incompatibility in Australian populations of the mosquito Culex quinquefasciatus Say. Med. Vet. Entomol. 1992, 6, 209–216. [Google Scholar] [CrossRef]
  218. Bourtzis, K.; Nirgianaki, A.; Markakis, G.; Savakis, C. Wolbachia infection and cytoplasmic incompatibility in Drosophila species. Genetics 1996, 144, 1063–1073. [Google Scholar]
  219. Hoffmann, A.A.; Turelli, M. Unidirectional incompatibility in Drosophila simulans: Inheritance, geographic variation and fitness effects. Genetics 1988, 119, 435–444. [Google Scholar]
  220. Holden, P.R.; Brookfield, J.F.; Jones, P. Cloning and characterization of an ftsZ homologue from a bacterial symbiont of Drosophila melanogaster. Mol. Genet. Genomics 1993, 240, 213–220. [Google Scholar] [CrossRef]
  221. Werren, J.H.; Jaenike, J. Wolbachia and cytoplasmic incompatibility in mycophagous Drosophila and their relatives. Heredity 1995, 75, 320–326. [Google Scholar] [CrossRef]
  222. Giordano, R.; O’Neill, S.L.; Robertson, H.M. Wolbachia infections and the expression of cytoplasmic incompatibility in Drosophila sechellia and D. mauritiana. Genetics 1995, 140, 1307–1317. [Google Scholar]
  223. Hoffmann, A.A.; Turelli, M.; Harshman, L.G. Factors affecting the distribution of cytoplasmic incompatibility in Drosophila simulans. Genetics 1990, 126, 933–948. [Google Scholar]
  224. Tagami, Y.; Doi, M.; Sugiyama, K.; Tatara, A.; Saito, T. Wolbachia-induced cytoplasmic incompatibility in Liriomyza trifolii and its possible use as a tool in insect pest control. Biol. Cont. 2006, 38, 205–209. [Google Scholar] [CrossRef]
  225. Kassem, H.A.; Hassan, A.N.; Abdel-Hamid, I.; Osman, G.; El Khalab, E.M.; Madkour, M.A. Wolbachia infection and the expression of cytoplasmic incompatibility in sandflies (Diptera: Psychodidae) from Egypt. Ann. Trop. Med. Parasitol. 2003, 97, 639–644. [Google Scholar] [CrossRef]
  226. Boller, E.F.; Russ, K.; Vallo, V.; Bush, G.L. Incompatible races of European cherry fruit fly, Rhagoletis cerasi (Diptera: Tephritidae), their origin and potential use in biological control. Entomol. Exp. Appl. 1976, 20, 237–247. [Google Scholar] [CrossRef]
  227. Riegler, M.; Stauffer, C. Wolbachia infections and superinfections in cytoplasmically incompatible populations of the European cherry fruit fly Rhagoletis cerasi (Diptera, Tephritidae). Mol. Ecol. 2002, 11, 2425–2434. [Google Scholar] [CrossRef]
  228. Vasquez, C.J.; Stouthamer, R.; Jeong, G.; Morse, J.G. Discovery of a CI-inducing Wolbachia and its associated fitness costs in the biological control agent Aphytis melinus DeBach (Hymenoptera: Aphelinidae). Biol. Cont. 2011, 58, 192–198. [Google Scholar] [CrossRef]
  229. Dedeine, F.; Vavre, F.; Shoemaker, D.D.; Boulétreau, M. Intra-individual coexistence of a Wolbachia strain required for host oogenesis with two strains inducing cytoplasmic incompatibility in the wasp Asobara tabida. Evolution 2004, 58, 2167–2174. [Google Scholar]
  230. Mochiah, M.B.; Ngi-Song, A.J.; Overholt, W.A.; Stouthamer, R. Wolbachia infection in Cotesia sesamiae (Hymenoptera: Braconidae) causes cytoplasmic incompatibility: Implications for biological control. Biol. Cont. 2002, 25, 74–80. [Google Scholar] [CrossRef]
  231. White, J.A.; Kelly, S.E.; Perlman, S.J.; Hunter, M.S. Cytoplasmic incompatibility in the parasitic wasp Encarsia inaron: Disentangling the roles of Cardinium and Wolbachia symbionts. Heredity 2009, 102, 483–489. [Google Scholar] [CrossRef]
  232. Noda, H. Cytoplasmic incompatibility in a rice planthopper. J. Heredity 1984, 75, 345–348. [Google Scholar]
  233. Rousset, F.; Bouchon, D.; Pintureau, B.; Juchault, P.; Solignac, M. Wolbachia endosymbionts responsible for various alterations of sexuality in arthropods. Proc. R. Soc. Lond. B 1992, 250, 91–98. [Google Scholar] [CrossRef]
  234. Hoshizaki, S.; Shimada, T. PCR-based detection of Wolbachia, cytoplasmic incompatibility microorganisms, infected in natural populations of Laodelphax striatellus (Homoptera: Delphacidae) in central Japan: Has the distribution of Wolbachia spread recently? Insect Mo. Biol. 1995, 4, 237–243. [Google Scholar]
  235. Vavre, F.; Fleury, F.; Varaldi, J.; Fouillet, P.; Boulétreau, M. Evidence for female mortality in Wolbachia-mediated cytoplasmic incompatibility in haplodiploid insects: Epidemiologic and evolutionary consequences. Evolution 2000, 54, 191–200. [Google Scholar]
  236. Vavre, F.; Dedeine, F.; Quillon, M.; Fouillet, P.; Fleury, F.; Boulétreau, M. Within-species diversity of Wolbachia-induced cytoplasmic incompatibility in haplodiploid insects. Evolution 2001, 55, 1710–1714. [Google Scholar]
  237. Mouton, L.; Henri, H.; Boulétreau, M.; Vavre, F. Multiple infections and diversity of cytoplasmic incompatibility in a haplodiploid species. Heredity 2005, 94, 187–192. [Google Scholar] [CrossRef]
  238. Machtelinckx, T.; van Leeuwen, T.; Vanholme, B.; Gehesquière, B.; Dermauw, W.; Vandekerkhove, B.; Gheysen, G.; De Clercq, P. Wolbachia induces strong cytoplasmic incompatibility in the predatory bug Macrolophus pygmaeus. Insect Mol. Biol. 2009, 18, 373–381. [Google Scholar] [CrossRef]
  239. Breeuwer, J.A.; Werren, J.H. Microorganisms associated with chromosome destruction and reproductive isolation between two insect species. Nature 1990, 346, 558–560. [Google Scholar] [CrossRef]
  240. Breeuwer, J.A.; Stouthamer, R.; Barns, S.M.; Pelletier, D.A.; Weisburg, W.G.; Werren, J.H. Phylogeny of cytoplasmic incompatibility micro-organisms in the parasitoid wasp genus Nasonia (Hymenoptera: Pteromalidae) based on 16S ribosomal DNA sequences. Insect Mol. Biol. 1992, 1, 25–36. [Google Scholar] [CrossRef]
  241. Bordenstein, S.R.; O’Hara, F.P.; Werren, J.H. Wolbachia-induced incompatibility precedes other hybrid incompatibilities in Nasonia. Nature 2001, 409, 707–710. [Google Scholar] [CrossRef]
  242. Saul, G.B. An analysis of non-reciprocal cross incompatibility in Mormoniella vitripennis (Walker). Zeitschrift für Vererbungslehre 1961, 92, 28–33. [Google Scholar] [CrossRef]
  243. Watanabe, M.; Miura, K.; Hunter, M.S.; Wajnberg, E. Superinfection of cytoplasmic incompatibility-inducing Wolbachia is not additive in Orius strigicollis (Hemiptera: Anthocoridae). Heredity 2011, 106, 642–648. [Google Scholar] [CrossRef]
  244. Narita, S.; Shimajiri, Y.; Nomura, M. Strong cytoplasmic incompatibility and high vertical transmission rate can explain the high frequencies of Wolbachia infection in Japanese populations of Colias erate poliographus (Lepidoptera: Pieridae). Bull. Entomol. Res. 2009, 99, 385–391. [Google Scholar] [CrossRef]
  245. Sasaki, T.; Ishikawa, H. Wolbachia Infections and Cytoplasmic Incompatibility in the Almond Moth and the Mediterranean Flour Moth. Zool. Sci. 1999, 16, 739–744. [Google Scholar] [CrossRef]
  246. Narita, S.; Nomura, M.; Kato, Y.; Yata, O.; Kageyama, D. Molecular phylogeography of two sibling species of Eurema butterflies. Genetica 2007, 131, 241–253. [Google Scholar] [CrossRef]
  247. Hiroki, M.; Ishii, Y.; Kato, Y. Variation in the prevalence of cytoplasmic incompatibility-inducing Wolbachia in the butterfly Eurema hecabe across the Japanese archipelago. Evol. Ecol. Res. 2005, 7, 931–942. [Google Scholar]
  248. Narita, S.; Nomura, M.; Kato, Y.; Fukatsu, T. Genetic structure of sibling butterfly species affected by Wolbachia infection sweep: Evolutionary and biogeographical implications. Mol. Ecol. 2006, 15, 1095–1108. [Google Scholar] [CrossRef]
  249. Kamoda, S.; Masui, S.; Ishikawa, H.; Sasaki, T. Wolbachia infection and cytoplasmic incompatibility in the cricket Teleogryllus taiwanemma. J. Exp. Biol. 2000, 203, 2503–2509. [Google Scholar]
  250. Moret, Y.; Juchault, P.; Rigaud, T. Wolbachia endosymbiont responsible for cytoplasmic incompatibility in a terrestrial crustacean: Effects in natural and foreign hosts. Heredity 2001, 86, 325–332. [Google Scholar] [CrossRef]
  251. Legrand, J.; Martin, G.; Artault, J. Corrélation entre la présence d’un symbiote bactérien dans les ovocytes de Porcellio dilatatus petiti, et la stérilité du croisement P. d. petiti mâle × P. d. dilatatus femelle. Inst. Pasteur. Tunis. 1978, 55, 507–514. [Google Scholar]
  252. Gotoh, T.; Noda, H.; Hong, X-Y. Wolbachia distribution and cytoplasmic incompatibility based on a survey of 42 spider mite species (Acari: Tetranychidae) in Japan. Heredity 2003, 91, 208–216. [Google Scholar] [CrossRef]
  253. Gotoh, T.; Noda, H.; Fujita, T.; Iwadate, K.; Higo, Y.; Saito, S.; Ohtsuka, S. Wolbachia and nuclear-nuclear interactions contribute to reproductive incompatibility in the spider mite Panonychus mori (Acari: Tetranychidae). Heredity 2005, 94, 237–246. [Google Scholar] [CrossRef]
  254. Tsagkarakou, A.; Guillemaud, T.; Rousset, F.; Navajas, M. Molecular identification of a Wolbachia endosymbiont in a Tetranychus urticae strain (Acari: Tetranychidae). Insect Mol. Biol. 1996, 5, 217–221. [Google Scholar] [CrossRef]
  255. van Opijnen, T.; Breeuwer, J.A. High temperatures eliminate Wolbachia, a cytoplasmic incompatibility inducing endosymbiont, from the two-spotted spider mite. Exp. Appl. Acarol. 1999, 23, 871–881. [Google Scholar] [CrossRef]
  256. Gotoh, T.; Sugasawa, J.; Noda, H.; Kitashima, Y. Wolbachia-induced cytoplasmic incompatibility in Japanese populations of Tetranychus urticae (Acari: Tetranychidae). Exp. Appl. Acarol. 2007, 42, 1–16. [Google Scholar] [CrossRef]
  257. Hunter, M.S.; Perlman, S.J.; Kelly, S.E. A bacterial symbiont in the Bacteroidetes induces cytoplasmic incompatibility in the parasitoid wasp Encarsia pergandiella. Proc. R. Soc. Lond. B 2003, 270, 2185–2190. [Google Scholar] [CrossRef]
  258. de Luna, C.J.; Moro, C.V.; Guy, J.H.; Zenner, L.; Sparagano, O.A.E. Endosymbiotic bacteria living inside the poultry red mite (Dermanyssus gallinae). Exp. Appl. Acarol. 2009, 48, 105–113. [Google Scholar] [CrossRef]
  259. Ros, V.I.D.; Breeuwer, J.A.J. The effects of, and interactions between, Cardinium and Wolbachia in the doubly infected spider mite Bryobia sarothamni. Heredity 2009, 102, 413–422. [Google Scholar] [CrossRef]
  260. Xie, R.; Zhou, L.; Zhao, Z.; Hong, X.; Xiao-Yue, H. Male age influences the strength of Cardinium-induced cytoplasmic incompatibility expression in the carmine spider mite Tetranychus cinnabarinus. Appl. Entomol. Zool. 2010, 45, 417–423. [Google Scholar] [CrossRef]
  261. Kambris, Z.; Blagborough, A.M.; Pinto, S.B.; Blagrove, M.S.C.; Godfray, H.C.J.; Sinden, R.E.; Sinkins, S.P. Wolbachia stimulates immune gene expression and inhibits plasmodium development in Anopheles gambiae. PLoS Pathog. 2010, 6, e1001143. [Google Scholar]
  262. Moreira, L.A.; Iturbe-Ormaetxe, I.; Jeffery, J.A.; Lu, G.; Pyke, A.T.; Hedges, L.M.; Rocha, B.C.; Hall-Mendelin, S.; Day, A.; Riegler, M.; et al. A Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya, and Plasmodium. Cell 2009, 139, 1268–1278. [Google Scholar] [CrossRef]
  263. Kambris, Z.; Cook, P.E.; Phuc, H.K.; Sinkins, S.P. Immune activation by life-shortening Wolbachia and reduced filarial competence in mosquitoes. Science 2009, 326, 134–136. [Google Scholar]
  264. Bian, G.; Xu, Y.; Lu, P.; Xie, Y.; Xi, Z. The endosymbiotic bacterium Wolbachia induces resistance to dengue virus in Aedes aegypti. PLoS Pathog. 2010, 6, e1000833. [Google Scholar] [CrossRef]
  265. Glaser, R.L.; Meola, M.A. The native Wolbachia endosymbionts of Drosophila melanogaster and Culex quinquefasciatus increase host resistance to West Nile virus infection. PloS One 2010, 5, e11977. [Google Scholar] [CrossRef]
  266. Fytrou, A.; Schofield, P.G.; Kraaijeveld, A.R.; Hubbard, S.F. Wolbachia infection suppresses both host defence and parasitoid counter-defence. Proc. R. Soc. Lond. B 2006, 273, 791–796. [Google Scholar] [CrossRef]
  267. Osborne, S.E.; Leong, Y.S.; O’Neill, S.L.; Johnson, K.N. Variation in antiviral protection mediated by different Wolbachia strains in Drosophila simulans. PLoS Pathog. 2009, 5, e1000656. [Google Scholar] [CrossRef]
  268. Panteleev, D.I.; Goriacheva, I.I.; Andrianov, B.V.; Reznik, N.L.; Lazebny, O.E.; Kulikov, A.M. The endosymbiotic bacterium Wolbachia enhances the nonspecific resistance to insect pathogens and alters behavior of Drosophila melanogaster. Genetika 2007, 43, 1277–1280. [Google Scholar]
  269. Hedges, L.M.; Brownlie, J.C.; O’Neill, S.L.; Johnson, K.N. Wolbachia and virus protection in insects. Science 2008, 322, 702. [Google Scholar]
  270. Teixeira, L.; Ferreira, A.; Ashburner, M. The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster. PLoS Biol. 2008, 6, e2. [Google Scholar]
  271. Walker, T.; Johnson, P.H.; Moreira, L.A.; Iturbe-Ormaetxe, I.; Frentiu, F.D.; McMeniman, C.J.; Leong, Y.S.; Dong, Y.; Axford, J.; Kriesner, P.; et al. The wMel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations. Nature 2011, 476, 450–453. [Google Scholar]
  272. Jaenike, J.; Unckless, R.; Cockburn, S.N.; Boelio, L.M.; Perlman, S.J. Adaptation via symbiosis: Recent spread of a Drosophila defensive symbiont. Science 2010, 329, 212–215. [Google Scholar]
  273. Hosokawa, T.; Koga, R.; Kikuchi, Y.; Meng, X-Y.; Fukatsu, T. Wolbachia as a bacteriocyte-associated nutritional mutualist. Proc. Natl. Acad. Sci. USA 2010, 107, 769–774. [Google Scholar]
  274. Supali, T.; Djuardi, Y.; Pfarr, K.M.; Wibowo, H.; Taylor, M.J.; Hoerauf, A.; Houwing-Duistermaat, J.J.; Yazdanbakhsh, M.; Sartono, E. Doxycycline treatment of Brugia malayi-infected persons reduces microfilaremia and adverse reactions after diethylcarbamazine and albendazole treatment. Clin. Infect. Dis. 2008, 46, 1385–1393. [Google Scholar]
  275. Rao, R.; Well, G.J. In vitro effects of antibiotics on Brugia malayi worm survival and reproduction. J. Parasitol. 2002, 88, 605–611. [Google Scholar]
  276. Bandi, C.; Trees, A.J.; Brattig, N.W. Wolbachia in filarial nematodes: Evolutionary aspects and implications for the pathogenesis and treatment of filarial diseases. Vet. Parasitol. 2001, 98, 215–238. [Google Scholar] [CrossRef]
  277. Bandi, C.; McCall, J.W.; Genchi, C.; Corona, S.; Venco, L.; Sacchi, L. Effects of tetracycline on the filarial worms Brugia pahangi and Dirofilaria immitis and their bacterial endosymbionts Wolbachia. Int. J. Parasitol. 1999, 29, 357–364. [Google Scholar] [CrossRef]
  278. Genchi, C.; Sacchi, L.; Bandi, C.; Venco, L. Preliminary results on the effect of tetracycline on the embryogenesis and symbiotic bacteria (Wolbachia) of Dirofilaria immitis. An update and discussion. Parassitologia 1998, 40, 247–249. [Google Scholar]
  279. Hoerauf, A.; Nissen-Pähle, K.; Schmetz, C.; Henkle-Dührsen, K.; Blaxter, M.L.; Büttner, D.W.; Gallin, M.Y.; Al-Qaoud, K.M.; Lucius, R.; Fleischer, B. Tetracycline therapy targets intracellular bacteria in the filarial nematode Litomosoides sigmodontis and results in filarial infertility. J. Clin. Invest. 1999, 103, 11–18. [Google Scholar] [CrossRef]
  280. Hoerauf, A.; Volkmann, L.; Nissen-Paehle, K.; Schmetz, C.; Autenrieth, I.; Büttner, D.W.; Fleischer, B. Targeting of Wolbachia endobacteria in Litomosoides sigmodontis: Comparison of tetracyclines with chloramphenicol, macrolides and ciprofloxacin. Trop. Med. Int. Health 2000, 5, 275–279. [Google Scholar] [CrossRef]
  281. Townson, S.; Hutton, D.; Siemienska, J.; Hollick, L.; Scanlon, T.; Tagboto, S.K.; Taylor, M.J. Antibiotics and Wolbachia in filarial nematodes: Antifilarial activity of rifampicin, oxytetracycline and chloramphenicol against Onchocerca gutturosa, Onchocerca lienalis and Brugia pahangi. Ann. Trop. Med. Parasitol. 2000, 94, 801–816. [Google Scholar] [CrossRef]
  282. Langworthy, N.G.; Renz, A.; Mackenstedt, U.; Henkle-Dührsen, K.; de Bronsvoort, M.B.; Tanya, V.N.; Donnelly, M.J.; Trees, A.J. Macrofilaricidal activity of tetracycline against the filarial nematode Onchocerca ochengi: Elimination of Wolbachia precedes worm death and suggests a dependent relationship. Proc. R. Soc. Lond. B 2000, 267, 1063–1069. [Google Scholar] [CrossRef]
  283. Hoerauf, A.; Mand, S.; Adjei, O.; Fleischer, B.; Büttner, D.W. Depletion of Wolbachia endobacteria in Onchocerca volvulus by doxycycline and microfilaridermia after ivermectin treatment. Lancet 2001, 357, 1415–1416. [Google Scholar]
  284. Hoerauf, A.; Mand, S.; Fischer, K.; Kruppa, T.; Marfo-Debrekyei, Y.; Debrah, A.Y.; Pfarr, K.M.; Adjei, O.; Büttner, D.W. Doxycycline as a novel strategy against bancroftian filariasis-depletion of Wolbachia endosymbionts from Wuchereria bancrofti and stop of microfilaria production. Med. Microbiol. Immunol. 2003, 192, 211–216. [Google Scholar] [CrossRef]
  285. Zchori-Fein, E.; Borad, C.; Harari, A.R. Oogenesis in the date stone beetle, Coccotrypes dactyliperda, depends on symbiotic bacteria. Physiol. Entomol. 2006, 31, 164–169. [Google Scholar] [CrossRef]
  286. Son, Y.; Luckhart, S.; Zhang, X.; Lieber, M.J.; Lewis, E.E. Effects and implications of antibiotic treatment on Wolbachia-infected vine weevil (Coleoptera: Curculionidae). Agr. Forest. Entomol. 2008, 10, 147–155. [Google Scholar] [CrossRef]
  287. Timmermans, M.J.T.N.; Ellers, J. Wolbachia endosymbiont is essential for egg hatching in a parthenogenetic arthropod. Evol. Ecol. 2008, 23, 931–942. [Google Scholar] [CrossRef]
  288. Starr, D.J.; Cline, T.W. A host parasite interaction rescues Drosophila oogenesis defects. Nature 2002, 418, 76–79. [Google Scholar]

Supplementary Materilas

Table Table S1. Other phenotypes induced by Wolbachia, Spiroplasma and Cardinium.

Click here to display table

Table S1. Other phenotypes induced by Wolbachia, Spiroplasma and Cardinium.
PhenotypeEndosymbiontClassOrderSpeciesReference
CIWolbachiaInsectaColeopteraCallosobruchus chinensis[195]
CIWolbachiaInsectaColeopteraChelymorpha alternans[196]
CIWolbachiaInsectaColeopteraD. virgifera[197]
CIWolbachiaInsectaColeopteraD. virgifera virgifera[197]
CIWolbachiaInsectaColeopteraD. virgifera zeae[197]
CIWolbachiaInsectaColeopteraHypera postica[198,199,200]
CIWolbachiaInsectaColeopteraTribolium confusum[201,202]
CIWolbachiaInsectaDipteraAedes albopictus[200]
CIWolbachiaInsectaDipteraA. cooki [209,204]
CIWolbachiaInsectaDipteraA. kesseli[205,206]
CIWolbachiaInsectaDipteraA. malayensis[207,208]
CIWolbachiaInsectaDipteraA. polynesiensis[207,208]
CIWolbachiaInsectaDipteraA. pseudoscutellaris[209,210]
CIWolbachiaInsectaDipteraA. s. scutellaris[209,211]
CIWolbachiaInsectaDipteraArmigeres subalbatus[212]
CIWolbachiaInsectaDipteraCulex pipiens[213,214,215]
CIWolbachiaInsectaDipteraC. pipiens quinquefasciatus[216,217]
CIWolbachiaInsectaDipteraDrosophila auraria[218]
CIWolbachiaInsectaDipteraD. melanogaster[219,220]
CIWolbachiaInsectaDipteraD. recens[221]
CIWolbachiaInsectaDipteraD. sechellia[222]
CIWolbachiaInsectaDipteraD. simulans[219,224]
CIWolbachiaInsectaDipteraLiriomyza trifolii[224]
CIWolbachiaInsectaDipteraPhlebotomus papatasi[225]
CIWolbachiaInsectaDipteraRhagoletis cerasi[226,227]
CIWolbachiaInsectaHymenopteraAphytis melinus DeBach [228]
CIWolbachiaInsectaHymenopteraAsobara tabida[229]
CIWolbachiaInsectaHymenopteraCotesia sesamiae[230]
CIWolbachiaInsectaHymenopteraEncarsia inaron[231]
CIWolbachiaInsectaHemipteraLaodelphax striatellus[232,233,234]
CIWolbachiaInsectaHemipteraLeptopilina heterotoma[235,236,237]
CIWolbachiaInsectaHemipteraMacrolophus pygmaeus[238]
CIWolbachiaInsectaHemipteraNasonia giraulti[239,240]
CIWolbachiaInsectaHemipteraN. longicornis[240,241]
CIWolbachiaInsectaHemipteraN. vitripennis[240,242]
CIWolbachiaInsectaHemipteraOrius strigicollis[243]
CIWolbachiaInsectaLepidopteraColias erate poliographus[244]
CIWolbachiaInsectaLepidopteraEphestia cautella[245]
CIWolbachiaInsectaLepidopteraE. kuehniella[246]
CIWolbachiaInsectaLepidopteraEurema hecabe[247]
CIWolbachiaInsectaLepidopteraE. madarina[248,249]
CIWolbachiaInsectaOrthopteraGryllus assimilis[197]
CIWolbachiaInsectaOrthopteraG. integer[197]
CIWolbachiaInsectaOrthopteraG. ovisopis[197]
CIWolbachiaInsectaOrthopteraG. pennsylvanicus[197]
CIWolbachiaInsectaOrthopteraG. rubens[197]
CIWolbachiaInsectaOrthopteraTeleogryllus taiwanemma [249]
CIWolbachiaMalacostracaIsopodaCylisticus convexus[250]
CIWolbachiaMalacostracaIsopodaPorcellio dilatatus[233,251]
CIWolbachiaArachnidaTrombidiformesOligonychus gotohi[252]
CIWolbachiaArachnidaTrombidiformesPanonychus mori[252,253]
CIWolbachiaArachnidaTrombidiformesTetranychus urticae[253,254]
CICardiniumInsectaHymenopteraEncarsia pergandiella[257]
CICardiniumArachnidaMesostigmataDermanyssus gallinae[258]
CICardiniumArachnidaTrombidiformesBryobia sarothamni[289]
CICardiniumArachnidaTrombidiformesEotetranychus suginamensis[256]
CICardiniumArachnidaTrombidiformesTetranychus cinnabarinus[260]
RP aWolbachiaInsectaDipteraAnopheles gambiae[261]
RP bWolbachiaInsectaDipteraAedes aegypti[262,263,264]
RP cWolbachiaInsectaDipteraCulex quinquefasciatus[265]
RP dWolbachiaInsectaDipteraDrosophila simulans[266,267]
RP eWolbachiaInsectaDipteraD. melanogaster[265,268,269,270]
RP fSpiroplasmaInsectaDipteraD. hydei[271]
RP gSpiroplasmaInsectaDipteraD. neotestacea[272]
MWolbachiaInsectaHemipteraCimex lectularius[273]
MWolbachiaSecernenteaSpiruridaBrugia malayi [274-271]
MWolbachiaSecernenteaSpiruridaB. pahangi[277]
MWolbachiaSecernenteaSpiruridaDirofilaria immitis[277,278]
MWolbachiaSecernenteaSpiruridaLitomosoides sigmodontis[279,280]
MWolbachiaSecernenteaSpiruridaOnchocerca gutturosa [281]
MWolbachiaSecernenteaSpiruridaO. lienalis[281]
MWolbachiaSecernenteaSpiruridaO. ochengi[282]
MWolbachiaSecernenteaSpiruridaO. volvulus[283]
MWolbachiaSecernenteaSpiruridaWuchereria bancroft[284]
OWolbachiaInsectaHymenopteraAsobara tabida[236]
OWolbachiaInsectaColeopteraCoccotrypes dactyliperda [285]
OWolbachiaInsectaColeopteraOtiorhynchus sulcatus[286]
OWolbachiaInsectaCollembolaFolsomia candida[287]
OWolbachiaInsectaDipteraDrosophila melanogaster (Sxl)[288]

CI, cytoplasmic incompatibility; RP, resistance to pathogen; M mutualism; O; oogenesis.a Confer resistance to Plasmodium berghei and P. falciparumb Confer resistance to Plasmodium gallinaceum, Dengue, Chikungunya and Brugia pahangic Confer resistance to West Nile virusd Confer resistance to Drosophila C virus, Flock House virus, Denguee Confer resistance to Drosophila C virus, Nora virus, Flock House virus, West Nile virus and Blauveria bassianaf Confer resistance to the parasitic wasp Leptopilina heterotomag Confer resistance to the nematode Howardula aoronymphium

Insects EISSN 2075-4450 Published by MDPI AG, Basel, Switzerland RSS E-Mail Table of Contents Alert