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New Insights into the Evolution of Wolbachia Infections in Filarial Nematodes Inferred from a Large Range of Screened Species

  • Emanuele Ferri ,

    Contributed equally to this work with: Emanuele Ferri, Odile Bain

    Affiliation Dipartimento di Biotecnologie e Bioscienze, Università degli Studi di Milano Bicocca, Milano, Italy

  • Odile Bain ,

    Contributed equally to this work with: Emanuele Ferri, Odile Bain

    Affiliation Parasitologie Comparée UMR 7205 CNRS & UMR 7245 CNRS, Muséum National d'Histoire Naturelle, Paris, France

  • Michela Barbuto,

    Affiliation Dipartimento di Biotecnologie e Bioscienze, Università degli Studi di Milano Bicocca, Milano, Italy

  • Coralie Martin,

    Affiliation Parasitologie Comparée UMR 7205 CNRS & UMR 7245 CNRS, Muséum National d'Histoire Naturelle, Paris, France

  • Nathan Lo,

    Affiliation School of Biological Sciences, University of Sydney, Sidney, Australia

  • Shigehiko Uni,

    Affiliation Department of Medical Zoology, Osaka City University Medical School, Osaka, Japan

  • Frederic Landmann,

    Affiliation Department of Molecular, Cell and Developmental Biology, University of California Santa Cruz, Santa Cruz, California, United States of America

  • Sara G. Baccei,

    Affiliation Dipartimento di Biotecnologie e Bioscienze, Università degli Studi di Milano Bicocca, Milano, Italy

  • Ricardo Guerrero,

    Affiliation Instituto de Zoologia Tropical, Universidad Central de Venezuela, Caracas, Venezuela

  • Sueli de Souza Lima,

    Affiliation Departamento de Zoologia, Universidade Federal de Juiz de Fora, Minas Geraes, Brasil

  • Claudio Bandi,

    Affiliation Dipartimento di Patologia Animale, Igiene e Sanità Pubblica Veterinaria, Università degli Studi di Milano, Milano, Italy

  • Samuel Wanji,

    Affiliation Research Foundation in Tropical Diseases and Environment, Buea, Cameroun

  • Moustapha Diagne,

    Affiliation Département de Biologie Animale, Université Cheikh Anta Diop de Dakar, Dakar, Sénégal

  • Maurizio Casiraghi

    Affiliation Dipartimento di Biotecnologie e Bioscienze, Università degli Studi di Milano Bicocca, Milano, Italy



Wolbachia are intriguing symbiotic endobacteria with a peculiar host range that includes arthropods and a single nematode family, the Onchocercidae encompassing agents of filariases. This raises the question of the origin of infection in filariae. Wolbachia infect the female germline and the hypodermis. Some evidences lead to the theory that Wolbachia act as mutualist and coevolved with filariae from one infection event: their removal sterilizes female filariae; all the specimens of a positive species are infected; Wolbachia are vertically inherited; a few species lost the symbiont. However, most data on Wolbachia and filaria relationships derive from studies on few species of Onchocercinae and Dirofilariinae, from mammals.

Methodology/Principal Findings

We investigated the Wolbachia distribution testing 35 filarial species, including 28 species and 7 genera and/or subgenera newly screened, using PCR, immunohistochemical staining, whole mount fluorescent analysis, and cocladogenesis analysis. (i) Among the newly screened Onchocercinae from mammals eight species harbour Wolbachia but for some of them, bacteria are absent in the hypodermis, or in variable density. (ii) Wolbachia are not detected in the pathological model Monanema martini and in 8, upon 9, species of Cercopithifilaria. (iii) Supergroup F Wolbachia is identified in two newly screened Mansonella species and in Cercopithifilaria japonica. (iv) Type F Wolbachia infect the intestinal cells and somatic female genital tract. (v) Among Oswaldofilariinae, Waltonellinae and Splendidofilariinae, from saurian, anuran and bird respectively, Wolbachia are not detected.


The absence of Wolbachia in 63% of onchocercids, notably in the ancestral Oswaldofilariinae estimated 140 mya old, the diverse tissues or specimens distribution, and a recent lateral transfer in supergroup F Wolbachia, modify the current view on the role and evolution of the endosymbiont and their hosts. Further genomic analyses on some of the newly sampled species are welcomed to decipher the open questions.


The alpha proteobacteria Wolbachia (Rickettsiales) are present in two distinct zoological groups: the arthropods, where they are widespread [1], and the nematodes, where they are restricted to a single but notable family of parasites, the Onchocercidae [2], [3]. They encompass the agents of human onchocerciasis and lymphatic filariases [4]. The zoological host range of Wolbachia raised a fundamental question on the origin of infection in the filarial nematodes [5], [6]. Investigations performed during the past fifteen years on Wolbachia in filarial and arthropod hosts has led to establish a rather clear and complex picture of the taxonomic status of the bacterium, its distribution and phylogeny [7]. Several distinct bacterial lineages have been called supergroups [8], and, at this date, they are all attributed to the only valid recognized species Wolbachia pipientis. The taxonomy of this species is quite uncertain, and in the scientific literature the genus name Wolbachia has been widely used as a specific name. This is taxomically incorrect, but common in microbiology (where species concept is usually complicated) and in the present work we will follow this trend until new data will be made available for a proper taxonomic restructuring [9], [10]. The supergroups are in majority distinct in arthropods and filariae: A, B, E, H, I, K are found in the arthropods; C, D and J in the nematodes [6], [8], [11]. However, the supergroup F is a relevant and very well supported exception, encompassing arthropod and filarial hosts (i.e. some insects such as termites and the human filariae of the genus Mansonella, [12][15]). Moreover, a newly discovered Wolbachia harboured by a plant parasitic nematode might represent a further supergroup [16], while the supergroup G [17] has been decommissioned due to the high probability of being characterised on the basis of an event of recombination [18].

Whereas the bacteria are mainly parasites in arthropods, usually acting as manipulators of reproduction [19][21], they are mutualistic in filariae [4], [22]. These mechanisms may be diverse, considering that Wolbachia is not only present in the germline but also in a somatic tissue, the hypodermis (lateral chords) of both females and males [23][25]. The biological studies and the Wolbachia genome projects [26] allowed us to suppose that the bacteria may be essential in the biosynthesis of some molecules necessary for filarial host fertility and viability, such as heme, riboflavin or nucleotide synthesis. Biosynthetic pathways are currently analyzed to determine the components of the symbiotic relationships [27][32]. To date, the mutualistic partnership is targeted in treatments against filariases using antibiotics [33].

The spiruroid ancestors of filariae that have been screened so far are devoid of endobacteria [5], [34]. The presence/absence of Wolbachia mapped on a filarial nematodes phylogenetic tree suggests that the bacteria may have possibly been acquired as a single event in the lineage leading to the onchocercid nematodes, followed by host-parasite co-evolution, assuming that Wolbachia in filariae was strictly vertically transmitted to the offspring through the infected female germline [3], [35]. Analysis of supergroup F is changing this view, due to the presence of Wolbachia from both filariae and some insects.

Another potential discrepancy with respect to the suggestion of coevolution is provided by observations of the absence of Wolbachia in two filarial species within the onchocercid lineage: the human parasite Loa loa and the rodent parasite Acanthocheilonema viteae [3], [36]. It has been suggested that for these host species, the endobacteria had been present but subsequently were lost during further evolution [5]. As a corollary, the loss of Wolbachia led to a further hypothesis that the bacterial genes essential to the host fitness might have been successfully transferred and expressed into the host genome. Although still subject of discussion [21], some support for this hypothesis derives from studies on lateral gene transfer, as shown with several insect and filarial hosts [37]. Remnants of Wolbachia-like gene sequences have been identified in the filarial host genomes of the endobacteria-free L. loa and A. viteae, with some of the transferred genes being transcribed [38]. The elimination of the bacteria might be an adaptive advantage because their antigens are inflammation inducers and contribute to filarial pathologies and immunological responses [39]. However, a recent study suggests that the bacteria might act as a decoy target for polynuclear neutrophils, preventing harmful effect of eosinophils on filariae [40]. Furthermore, a strain of Wolbachia that over-replicates in Aedes aegypti inhibits the development of Brugia malayi larvae and switches on a few important immune system genes [41][43]. Thus the limitation of the filarial infection may either be due to immune activation of the invertebrate host or/and the bacteria may outcompetes filariae for some metabolites.

In our previous study [5], it appeared that the number of endobacteria-free filarial species had been underestimated and that several species without Wolbachia detected were parasitic in lizards and frogs. Thus it was suggested that these filariae from reptiles and anurans diversified before the first bacterial invasion on the onchocercid lineage which had been tentatively dated to 110 mya [3], [6]. Indeed the origin of the Oswaldofilariinae, parasitic in crocodiles and squamates, was hypothetically dated from the late Jurassic, at the beginning of Gondwanian dislocation, 140 mya [44], [45]. However representatives of this subfamily had not yet been screened.

Until the present study, Wolbachia screening had been done in only about 10% of the 93 genera currently recognized in the Onchocercidae [46], [47]. This is not surprising since the recovery of filariae from connective tissues, their main localization, is not easy. Our investigation was resolutely rooted in biodiversity, expecting that the exploration of a broader range of filarial species would contribute to decipher the history of the Wolbachia-filaria symbiosis. The recovery of materials from wild animals from several biomes was undertaken. The first oswaldofilarine, the first splendidofilarine (a parasite of birds), and several onchocercid genera from mammals have now been screened through PCR, as well as classic immunohistochemical staining and whole mount fluorescent analysis [21].

This study confirms that Wolbachia are not detected until now in the filarioid species parasitic in amphibians and reptiles. Several other features have emerged from this study: i) lateral hypodermal localization of Wolbachia is not obligatory in bacteria-positive filarial species; ii) new somatic tissue localizations of the bacteria are observed; iii) the number of Wolbachia-free filarial species is greater than expected among filariae of mammals; (iv) lastly, one secondary event of Wolbachia infection, also well supported by a formal cocladogenesis, likely took place in filariae in supergroup F.


The screening for Wolbachia was performed on 35 species (Table 1; specimens detailed in Figure S1 and Tables S1, S2), of which 28 are here examined for the first time and one recently by us [48]. These were the first representatives of Oswaldofilariinae, Piratuba scaffi from a lizard, and of Splendidofilariinae, Aproctella sp. 1 from passeriforms; two more species, Ochoterenella sp. 1 and O. royi, in Waltonellinae, a subfamily restricted to anurans; five genera of Onchocercinae parasitic in mammals, a species of Monanema, Mo. martini, parasitic in African murids and used during a decade as a model for onchocerciasis because of its skin-dwelling microfilariae [49]; several species of Cercopithifilaria, six from ruminants, one from a bear (all from Japan), and one species from an African porcupine; Loxodontofilaria, with the recently described Lo. caprini, recovered from a Japanese caprine bovid [50]; in the genus Mansonella, two of the six subgenera, Tetrapetalonema and Cutifilaria, with a species each, M. (T.) atelensis amazonae from a monkey, and M. (Cu.) perforata from a cervid; and the recently studied species of Litomosa from a South African bat (reported in [48].

Table 1. The 35 species of filariae included in this study, their hosts and collection place.

The histoimmunostainings are presented according to the following genera: Litomosoides and Litomosa (Figure 1), Onchocerca and Loxodontofilaria (Figure 2), Cercopithifilaria japonica and Mansonella (Figure 3), other species of Cercopithifilaria (Figure 4). Whole mount fluorescent analysis is presented on Figure 5.

Figure 1. Wolbachia immunostaining in the genera Litomosoides (L.) and Litomosa (Li.).

Transverse sections of gravid females of three species. A–D. Litomosoides sigmodontis, laboratory strain. A. Cuticle, lateral crests and muscles thin; hypodermal lateral chords Wb+; female germline Wb+, as seen in the rachis of the distal region of ovaries. B. Less distal region of an ovary Wb+, uterus with ova/eggs Wb+ and spermatozoa Wb–. C. Uteri with many divided eggs Wb+. D. Uteri with many microfilariae Wb+. E. Litomosoides yutajensis: cuticle and lateral hypodermis thin; lateral crests and muscles obvious; hypodermal lateral chords Wb–, germ line Wb–, as seen with intra-uterine divided eggs. F. Detail. G. Litomosa chiropterorum: muscles, lateral cuticular crests and hypodermis well distinct; hypodermal lateral chords and germ line Wb–. H. Detail of uteri with microfilarioid eggs Wb–. I, Intestine; O, Ovary; R, Rachis of ovary; U, Uterus; c, cuticle; h, hypodermal lateral chords demarcated by white stars; m, muscles; *, lateral plan; white arrow, oocytes; full arrow, ovulae. Scale bars: F & H, 10 µm; others, 25 µm. Sections were stained with a rabbit polyclonal antiserum against Wolbachia Surface Protein (WSP) of Brugia pahangi Wolbachia (Wol-Bp-WSP, dilution 1∶2000).

Figure 2. Wolbachia immunostaining in the genera Onchocerca (O.) and Loxodontofilaria (Lo).

Transverse sections of females of four species of filariae with Wb+ germ line, at two magnifications. A. Onchocerca dewittei japonica: cuticle thick, lateral crest not marked, hypodermal chords well delineated and Wb–, large section of uterus and small sections of ovaries Wb+. B: Detail of lateral hypodermal chord Wb– and ovary Wb+. C: Detail of intra-uterine ova and ovary Wb+. D. Onchocerca eberhardi: thick cuticle with few ridges, thin muscles and thinner Wb+ lateral hypodermis; uteri with a few sections of divided eggs Wb+. E. Detail. F. Onchocerca skrjabini: cuticle very thick, with particular crescent-like layer forming the lateral crest, very thin lateral hypodermis Wb+ and empty uteri. G. Detail. H. Loxodontofilaria caprini: cuticle thick, well-developed muscle cells, thinner lateral chords Wb–; two uteri, one empty, one with Wb+ ova. I. Detail. I, Intestine; O, Ovary; U, Uterus; c, cuticle; h, hypodermal lateral chords delimitated by white stars; m, muscles; *, lateral plan; thin arrow, detail of Wolbachia. Scale bars: A,D,H, 50 µm; B,C,E,G,I, 25 µm. For staining, see Fig. 1.

Figure 3. Wolbachia immunostaining in the genus Cercopithifilaria (C.).

Transverse sections of females of four species Wb–, at two magnifications. A. Cercopithifilaria crassa: muscles and lateral cuticular crests thick; lateral narrow thick hypodermis and distal ovaries Wb–. B. Detail. C. Cercopithifilaria longa: thin lateral chords Wb–, two uteri with microfilariae Wb–. D. Detail. E. Cercopithifilaria minuta: thin lateral chords Wb –, thick muscles, two uteri with divided eggs Wb–. F. Detail. G. Cercopithifilaria shohoi: lateral crest rather thick, thin hypodermal chord Wb–, small section of ovary with oocytes Wb– and large section of uterus with ova Wb–. H. Detail. I, Intestine; O, Ovary; U, Uterus; c, cuticle; h, hypodermal lateral chords delimitated by white stars; m, muscles; *, lateral plan; white arrow, oocytes. Scale bars: A,C,E,G, 25 µm; B,D,F,H, 10 µm. For staining, see Fig. 1.

Figure 4. Immunostaining of Wolbachia in females of two species.

A. Cercopithifilaria japonica: wide rather flat lateral crests, hypodermis thin and Wb+, muscles thicker, germ line Wb+. as seen with the intra-uterine ovulae (on left); Wb– spermatozoa in this uterus and in the smaller more distal section (on right). B. Detail. C. Mansonella (Cutifilaria) perforata: lateral chords wide, flat and Wb–, muscle cells much thicker, germ line Wb+ as seen with 3 divided eggs in one of the uteri, intestine Wb+. D. Detail. E. Other section, intestine Wb+, eggs Wb+ and hypodermal lateral chord Wb–. F. Detail. I, Intestine; O, Ovary; R, Rachis of ovary; U, Uterus; c, cuticle; h, hypodermal lateral chords delimitated by white stars; m, muscles; *, lateral plan; thin arrow, detail of Wolbachia. Scales: A,C,E, bar = 25 µm; B,D,F, bar = 10 µm. For staining, see Fig.1.

Figure 5. Whole mount fluorescent analysis of Wolbachia in females of four species stained with propidium iodide for DNA (in red) and phalloidin for actin (in green).

A. Hypodermal lateral chord of a specimen of Onchocerca dewittei japonica without Wolbachia. B. Idem, a second specimen with a few Wolbachia (arrow). C. Epithelial somatic gonad with Wolbachia in Mansonella (Cutifilaria) perforata. D. In the same specimen, intestinal wall cells (in) with Wolbachia and above, lateral chord (ch) without Wolbachia. E. Wolbachia in epithelial somatic gonad of Cercopithifilaria japonica. F. Metaphase in a Cercopithifilaria crassa zygote, showing the absence of Wolbachia in the embryo. Polar bodies (PB) on the left. *: host cell nuclei; arrow: Wolbachia. Scale bar = 20 µm.

The results (Table 1 and 2; Figures 17) can be summarised as following:

Figure 6. Phylogeny of Wolbachia based on 16S rDNA gene sequences.

Names at the terminal nodes are those of the host species (with the exceptions of the outgroup species). A–J are the supergroups names according to [1], [3], [8], [14]. The tree has been obtained by Bayesian inference of phylogeny, using MrBayes 2.01; numbers at the nodes are the posterior probability values. Species in bold are those newly found positive in the screening for Wolbachia performed in this study. Accession numbers are given for the sequences present in the databases.

Figure 7. Cocladogenesis of Wolbachia and filarial nematodes based on representatives of the species studied.

For these species sequences for nematode coxI and 12S rDNA (phylogeny on the left) and bacterial 16S rDNA (phylogeny on the right) were available. Dotted lines have been added to emphasize major discrepancies between filarial and Wolbachia trees. For further details, see paragraph 7 in the Results section.

Table 2. Synthetic results of the screening for Wolbachia with PCR, immuno-histo staining and whole mount fluorescent analysis.

1. The presence or absence of Wolbachia were confirmed in species previously screened (Tables 2 and S2). Species previously studied and as expected confirmed positive are: Onchocerca volvulus (female worm from a human nodule, Cameroon), Dirofilaria repens (from an Italian patient), Litomosoides sigmodontis (2 females recovered from a wild Sigmodon hispidus, Venezuela) and a less commonly studied species, Dipetalonema gracile (a female recovered from Cebus olivaceus, collected in Yutaje, Venezuela, like in [5] (Tables 2 and S2). The species confirmed negative were Loa loa (two batches of infective larvae recovered from Chrysops vectors, Cameroon) and the single Litomosoides species that did not harbour Wolbachia, L. yutajensis (3 females, from the same species, Pteronotus parnellii, and locality than in [5]; Figures 1E,F).

2. Eight newly screened filarial species harbored Wolbachia but, for some of them, not all specimens are positive (Tables 2 and S2). The species in which Wolbachia was detected are the following: Onchocerca dewittei japonica, from Sus scrofa leucomystax (Figures 2A–C); O. eberhardi, from Cervus nippon (Figures 2D,E); O. skrjabini, from Cervus nippon (Figures 2F,G) and Capricornis crispus; O. suzukii from Capricornis crispus; Loxodontofilaria caprini, from Capricornis crispus (Figures 2H,I); Cercopithifilaria japonica, from Ursus thibetanus (Figures 3A,B); Mansonella (Cutifilaria) perforata, from Cervus nippon (Figures 3C–F); M. (Tetrapetalonema) atelensis amazonae, from Cebus olivaceus (Tables 2 and S2).

However, in four infected filarial species, Wolbachia were not detected in each specimen. These species were Lo. caprini, M. (Cu.) perforata, O. d. japonica and O. suzukii. The prevalence of Wolbachia varied from 50% to 66% in these species (9, 4, 8 and 4 samples, respectively, see Tables 2 and S2).

3. In Wolbachia positive filarial species lateral hypodermal chords might be infected, might not be infected, or weakly infected. The presence of the bacteria in the female germline was not constantly associated to their presence in the lateral hypodermal chords. Among the Wolbachia positive species in which the tissue distribution of Wolbachia was studied, the lateral chords harboured Wolbachia in L. sigmodontis (Figures 1A–D), O. eberhardi (Figures 2D,E) and O. skrjabini (Figures 2F,G), C. japonica (Figures 3A,B). On the contrary, Wolbachia were not observed in the lateral chords of Lo. caprini (Figures 2H,I) and M. (Cu.) perforata (Figures 3E,F). In O. d. japonica, the bacteria were not detected in lateral chords of sectioned worms (Figures 2A–C) and in one of the two whole mounted worms (Figure 5A), but a few bacteria were observed in the second specimen (Figure 5B).

4. Rare or novel tissue Wolbachia localizations were observed in Mansonella (Cutifilaria) perforata and Cercopithifilaria japonica. In M. (Cu.) perforata the somatic gonad was found positive once on sectioned worms and this was confirmed on a whole mounted worm; the epithelium lining the gonad harboured the bacteria, but not the external muscle layer (Figure 5C). Moreover, bacteria were constantly found in the cells of the intestine wall, in both sectioned and whole mount material (Figures 3C–F; 5D). Wolbachia were also detected in the somatic gonad of C. japonica, in whole mounted worms, but not in sectioned worms (Figure 5E); in this species, Wolbachia were not found in the intestinal cells.

5. Four types of Wolbachia were identified in the whole study. Interestingly, no events of recombination were found in 16S rDNA, dnaA, ftsZ and groEL genes of Wolbachia from the sequences of the filarial nematodes studied. According to the Wolbachia supergroups, four types of Wolbachia were identified in this study: bacteria harboured by Onchocerca, Dirofilaria and Loxodontofilaria species were assigned to supergroup C; bacteria harboured by L. sigmodontis and Litomosoides taylori were assigned to supergroup D; bacteria harboured by the species of the two subgenera of Mansonella (Tetrapetalonema and Cutifilaria) studied, as well as those harboured by Cercopithifilaria japonica were assigned to supergroup F; bacteria harboured by Dipetalonema gracile were assigned to supergroup J (Figure 6).

The main Wolbachia clusters shown in the 16S rDNA phylogenetic tree of figure 6 were ((Litomosa+Litomosoides) + (Wuchereria+Brugia)); (Dirofilaria+ (Onchocerca+Loxodontofilaria)); (Cercopithifilaria+Mansonella+Microcerotermes+Rhinocyllus+Kalotermes + Microcerotermes). Wolbachia from representatives of supergroups A, B and E formed separated and supported clusters; Wolbachia from Ctenocephalides and Dipetalonema formed two separate lineages, that were recently assigned to supergroups I and J respectively [14].

6. Wolbachia was not detected in more than half of the newly screened species (Tables 2 and S2). The Wolbachia negative species are: in Oswaldofilariinae, Piratuba scaffi; in Waltonellinae, Ochoterenella royi and Ochoterenella sp.1; in Setariinae, the five Setaria spp.; in Onchocercinae, 8 (upon nine) Cercopithifilaria species (Figures 4A–H, 5F), Litomosa chiropterorum (Figures 1G,H), and Monanema martini; in Splendidofilariinae, Aproctella sp. 1.

7. Inconsistency between phylogenies of Wolbachia and filarial hosts was evidenced in the cocladogenetic analysis. The topologies of the trees of the infected filariae and their respective Wolbachia were found to have a significant level of similarity for all of the four tree metrics tested in Component, based on comparisons of the Wolbachia tree with 1000 random host trees (p<0.03 in all cases). The lnL scores of the filarial and Wolbachia ML trees based on the filarial dataset were −6189.35 and −6207.14 respectively. A Shimodaira-Hasegawa test showed that the Wolbachia topology was significantly less likely (p = 0.035) than the filaria topology. The lnL scores of the Wolbachia and filarial ML trees based on the Wolbachia dataset were −10757.38 and −10950.85 respectively. A Shimodaira-Hasegawa test showed that the filaria tree was significantly less likely than the Wolbachia tree (p<0.001). Thus the null hypothesis that the host and parasite have strictly co-speciated was rejected. The major inconsistency was due to the newly screened Cercopithifilaria japonica and Mansonella (Cutifilaria) perforata (Figure 7).

Another point of inconsistency in the phylogenies of hosts and symbionts regards the positioning of L. sigmodontis: while Wolbachia from this filarial is placed as the sister group of endosymbionts from lymphatic filariae (B. malayi, B. pahangi and W. bancrofti), the filaria itself is placed as a deeper branch in the filarial tree (Figure 7). Indeed, based on morphological adult and larval criteria, Litomosoides is closer to lymphatic filariae than to Mansonella [45].


As expected, the screening of a broader and more diversified set of species samples, by a combination of PCR and gene sequencing, immunohistological staining and whole mount fluorescent analysis, revealed new information about Wolbachia biology and evolution: novel tissue localizations, strong evidence for recent transfers between unrelated filarial species, and a larger number of species that do not harbour Wolbachia. In addition, the occurrence of Wolbachia in some members of a species and its absence in others raises questions about the evolution of its obligate requirement [51].

1. New filarial tissues infected with Wolbachia

It is clear that tissues other than the female germline and the hypodermal lateral chords may be infected with Wolbachia. One of these infected tissues is the somatic gonad (epithelial layer), once briefly reported previously [52]. This Wolbachia localization was evidenced in Mansonella (Cutifilaria) perforata and Cercopithifilaria japonica (Table 2; Figures 5C,E). Interestingly, they are both members of the supergroup F of Wolbachia.

The real novelty is the Wolbachia tissue localization in the intestinal wall; it was only observed in M. (Cu.) perforata, but in all sectioned and whole mount samples (Figures 3C–F, 5D).

These divergent localizations suggest a more complex and diversified relationship between the bacteria and filariae. They also raise the question of how and when Wolbachia bacteria reach the appropriate filarial host tissues. It is likely that it is an early event, since it was shown in Brugia malayi an asymmetric distribution of bacteria in the egg followed by a preferential segregation in defined blastomeres [21].

2. Recent capture of Wolbachia type F suggested by Cercopithifilaria japonica, parasite of the Japanese bear

The species screened in this study generally confirm the previously identified types of Wolbachia in the Onchocercidae (Figure 6; see for instance [9]). The newly screened Loxodontofilaria is placed among the species of Onchocerca [53]. In addition, endosymbionts from this filaria belong to Wolbachia supergroup C (Figure 6). There is a major phylogenetic congruence discrepancy between Wolbachia and their hosts and it occurs in the genus Cercopithifilaria and the supergroup F of Wolbachia. One African and seven Japanese species of Cercopithifilaria have no Wolbachia, while one species in Japan is Wolbachia positive. The filarial hosts belong to a well-supported genus, Cercopithifilaria as evidenced by adult morphology [45], [54], larval morphology [55], 12S rDNA gene sequences [53], [56], the transmission by hard ticks and the skin-dwelling microfilariae [45]. A parsimonious interpretation of the Wolbachia screening is that a single acquisition event took place in C. japonica. This hypothesis is supported by the co-cladogenetic analyses (Figure 7).

The supergroup F is intriguing as it is presently the only Wolbachia type infecting both insects and onchocercid nematodes [7]. The Wolbachia supergroup F contains the species of Mansonella studied so far: M. (M.) ozzardi and M. (Esslingeria) perstans [13], [15], and in this study, M. (Tetrapetalonema) atelensis amazonae, M. (Cutifilaria) perforata. Cercopithifilaria japonica in supergroup F suggests a transversal transmission event, likely recent due to limited occurrence among the species of this genus. C. japonica is a parasite of the Japanese bear in which it coexists with a species of Mansonella of the subgenus Mansonella [57]. This Mansonella species, M. (M.) akitensis [58], has not been screened for Wolbachia but it likely harbours the type F Wolbachia. The bacterial host switching might have occurred between the two filarial parasites of the bear, perhaps via an oral infection route. Indeed filariae, despite their apparent small mouth, can ingest particles from their environment, such as red blood cells [59] and larger bodies, such as microfilariae released in the coelomic cavities of the filarial host [60].

3. Absence of Wolbachia

The filarial species in which Wolbachia were not detected appeared more numerous than it was thought, based upon previous observations. In [34], the percentage of negative species was 10.5% (2 negative among 19), in [5] with a larger sample, it was 37%. In this study, it is twice more elevated, 63%. It has to be emphasized that the negative results were not due to DNA degradations or bad extractions because all of the Wolbachia PCR negative samples gave positive amplifications using filarial nematode specific primers. However, we must take into account the fact that, in a few species, Wolbachia were not detected in all the specimens. This can partly be explained in species which do not harbour Wolbachia in the lateral chords, or at very low density, as Onchocerca dewittei japonica (Figures 5A,B) and Loxodontofilaria caprini (Figures 2H,I). In M. (Cu.) perforata, the bacteria are in the intestine wall of the female worms (Figures 3C,D, 5D) and any part of female body would be Wolbachia positive, but this is not the case (Tables 2, S1 and S2). Thus, in M. (Cu.) perforata, presence/absence of Wolbachia may occur, as suggested by [15] for another Mansonella species, M. (Esslingeria) perstans from humans. It is interesting to note that in both of these cases, the Wolbachia supergroup is F. Further, research performed on deeply studied filariae, such as Brugia malayi, have also shown that the amount of Wolbachia carried by a worm may vary greatly over time and be stage-dependent [21]. This dynamic probably has little impact when considering developing larval stages, because these are transient and the chance of recovering them in the wild is extremely low. More interesting is the observation that the female worms recovered in the wild are not all fully gravid in some species (Figure 2F). This was not the case of filariae from frogs, lizards, bats, birds, but was the case from some parasites of Japanese ungulates. It is worth to note that we paid great attention to the part of worm sampled to ensure that the germline was screened in all species.

4. Systematic position of Wolbachia negative filariae

It is clear that there is a need to increase the number of screened specimens for more solid results. However, in several cases, the global results are impressive and the distribution of the Wolbachia negative species does not appear random. As a matter of fact, at present, the species parasitic in frogs and lizards are negative for Wolbachia if they are Waltonellinae (three species of Ochoterenella), Oswaldofilariinae (one species of Piratuba), or Dirofilariinae (two species of Foleyella). It was also shown here that the first screened species parasitic in birds, an Aproctella in the Splendidofilariinae, was Wolbachia negative (7 females screened with PCR, from two passeriform species, totalising 7 specimens hosts). Wolbachia were not detected also in several species of Onchocercinae from mammals. The first is Litomosa chiropterorum, from an African bat (8 specimens screened from 8 Miniopterus schreibersi), an unexpected observation since the single Litomosa species screened previously, Li. westi from a North American rodent Geomyoidea, was Wolbachia positive [5].

The second species is Monanema martini (8 females screened from 8 murid specimens). The absence of the bacteria is an important feature, considering that this filaria was used, in the past, as a model of onchocerciasis, a true limit due to the important role played by the bacteria in the pathology of the disease [39], [61][63]. The absence of Wolbachia could explain partly the weak ocular pathology induced in the murid host [49]. Severe or mild ocular onchocerciases have been related to a strain-dependent variation of density of Wolbachia per filarial genome [64].

Other species in which Wolbachia were not detected belong to the genus Cercopithifilaria; 8 out of 9 species were Wolbachia-negative (Table 2; Figures 4; 5F).

Two hypotheses might be taken into account to explain the absence of Wolbachia. In the first case, Wolbachia could be considered to be present and then were subsequently lost [38]. This could have occurred before the mutually dependent symbiosis between symbiont and host developed. The second hypothesis is that the bacteria were not yet acquired within that filarial group.

The first hypothesis seems to possibly explain some of the observed cases, such as Litomosoides yutajensis (5 samples: 2 males previously and now 3 more females), a species without Wolbachia, in contrast with 5 congeneric species infected with Wolbachia. In the cases of Acantocheilonema viteae and Onchocerca flexuosa [38], the absence of Wolbachia appears to be a secondary loss, because some genes of the bacteria were incorporated in the filarial genomes. However, it is not clear whether this loss occurred before symbiosis was established or not. This is of interest because if loss was after symbiotic establishment, perhaps some of the genes incorporated into the host genome were those required by the host now provided by Wolbachia, but the extent of this event is still to be understood.

The hypothesis that the Wolbachia negative species might never have been infected, might be considered for the filarial groups that are supposed to be “ancient” such as the Oswaldofilariinae [44], [45], [65]. Estimation of dates of divergence has been proposed for some groups of nematodes based on molecular phylogenetic analyses [66]. In filariae, data from morphology, biology, geographic distribution, host range and palaeontology led to the proposal that the Oswaldofilariinae emerged during the late Jurassic, at the beginning of the Gondwanian break up, 140 mya [44], [45]. This is before the hypothesized ancestral acquisition of a Wolbachia by an onchocercid [3]; it follows that the absence of Wolbachia in Oswaldofilariinae could be primitive.

Foleyella is another parasite of saurians, which appears to have no Wolbachia; it is presently placed in the Dirofilariinae, which includes the Wolbachia positive Dirofilaria, but this systematic position needs to be revised because the characters of the infective larvae are very distinct [55]. A solid phylogeny of Onchocercidae linking traditional and molecular data is needed and warrants further investigations.

The subfamily Setariinae, parasitic in ungulates and in which Wolbachia were not detected, as first shown by [67], also deserves a comment. Based on larval morphology, it has been hypothesized that it evolved separately from the other onchocercids and derived from a group of spirurid Habronematinae [65]. Until now no spirurids have been found infected with Wolbachia [5], [34].

Did the bacterial infection never occur, or was the useful part of Wolbachia genome incorporated in the host genome and subsequently the bacteria eliminated, to reply to some adverse constraint? Further genomic analyses will solve this question concerning the absence of Wolbachia in ancestral filariae.

At present, the features observed on tissue or specimen distribution, and a very probable recent lateral transfer, suggest complex evolutionary dynamics of interactions between the symbionts, their host filariae and the nematode hosts. In the sampling studied, which will be further enlarged, it may become possible to use some defined species to decipher these questions.

Materials and Methods

Specimens and species

Specimens of filariae were recovered during dissections of the vertebrate hosts captured in the wild from different geographic areas [50], [54], [68][72].

All experiments, procedures and ethical issues were conformed to the competent national ethical bodies: Venezuelian animals were captured according to Licencia con Fines Cientificos N° 2192 dated June 18, 2007 and Contrato Marco Acceso Recursos Genéticos N° 33/2007 both granted by Ministerio del Ambiente de la Republica Bolivariana de Venezuela. Japanese serows, sika deer, bears, and wild boars were killed by hunters who have an individual permit to kill wild animals in accordance with the conservation and control policies of the Ministry of the Environment of Japan. Italian samples were collected by veterinarians and physicians and no permits were necessary. African rodents and agama do not belong to protected species and were obtained from local hunters. African mountain reedbuck, plain zebra, gemsbok and porcupine were also obtained from local hunters. Bats from South African bats have been collected for previous studies [48] in which no permits were necessary. Brazilian birds were donated by the Brazilian Institute of the Environment and Renewable Resources (IBAMA), Region of Juiz de Fora, Minas Gerais for the laboratory of Taxonomia and Ecology of Helminths of the Department of the Zoology.

Many of the filariae from large mammals were extracted from the subconnective tissue, dermis, or tendons of limbs. Host animals kept at 4°C were sent to the laboratory after they were killed. Afterwards the parts of the animals were dissected to collect living filarioids for Wolbachia study. From frogs, lizards, birds, bats and rodents filariae were generally recovered immediately after host death. Samples used for positive and negative PCR controls were laboratory strains.

Species identification of the specimens was done with morphological studies performed by several of us (OB, SU, RG, SL, MD). Some species are not yet named, but all are under study, and morphological analysis and sequencing of coxI gene in this study and in [53] showed that they represent distinct molecular entities. Co-infection of a host specimen by several congeneric or non-congeneric filarial species was rather frequent in large mammals, but in a few cases, the same filarial species was recovered from two host species. The supraspecific levels of taxonomy followed the systematic works of [46], [73],[74] and more recent studies for some taxa: [65], [71], [75] for the subgeneric divisions of Mansonella Faust, 1929; [54], [56] for the genus Cercopithifilaria Eberhard, 1980 (created as a subgenus); and [54] for the genus Loxodontofilaria Berghe & Gillain, 1939.

Abbreviations used in the text are: A. for Aproctella; C. for Cercopithifilaria; Cu. for Cutifilaria; D. for Dirofilaria; Di. for Dipetalonema; F. for Foleyella; L. for Litomosoides; Li. for Litomosa; Lo. for Loxondontofilaria; M. for Mansonella; Mo. for Monanema; O. for Onchocerca; Oc. for Ochoterenella; P. for Piratuba; S. for Setaria; T. for Tetrapetalonema. The material screened in this study is detailed in Figure S1, Tables S1 and S2.

Filarial worms were fixed and kept in absolute alcohol at 4°C for PCR analyses, in 4% PFA (paraformaldehyde) for overnight at 4°C for immunohistochemical staining and whole mount fluorescent analysis. In many cases, filarial specimens were cut into anterior (a), median (m) and posterior (p) parts, which were fixed for the different analysis approaches. Since Wolbachia are transmitted by female filariae, almost all of the studies were based on female worms; male specimens were rarely examined, and we analysed this sex alone in a single case, Di. gracile (Table S3).

Molecular screening for Wolbachia on filarial nematodes

PCR screening for Wolbachia was conducted according to [5], [12], using general Wolbachia primers for 16S rDNA (99f and 994r [76]), originally designed to work on Wolbachia from the supergroups A and B, and primers for 16S rDNA (16SWolbF and 16SWolbR3), originally designed to work on Wolbachia from the supergroups A–D [12], but whose target sites are also conserved in Wolbachia from supergroups E and F [8].

PCRs were performed in a 20 µl final volume under the following conditions: 1x buffer (containing 1.5 mM MgCl2, Eppendorf™), 0.2 mM of each dNTP, 1 µM of each primer, and 0.5 U of Taq DNA Polymerase (Eppendorf™). The thermal profile used was: 94°C 45 sec, 52°C 45 sec, and 72°C 90 sec for 40 cycles.

When the PCRs were negative under the above PCR conditions, a nested-PCR approach was implemented in order to improve the sensitivity of the PCR screening [5], [77]. The first PCR was performed using the general eubacterial primer 27F [78] combined with 16SWolbR3; PCR volumes and conditions were as above. Five µl were visualised on a 1.5% w/v agarose gel and one µl of the first PCR was diluted 1/10 and 1/100 in water, and then both used as templates in a second PCR, performed using primers W-EF and W-ER [70]. W-ER and W-EF recognize sites that are conserved in supergroups E–F and that are internal to the primers used in the first PCR. PCR conditions for this amplification were as described in [79].

Of samples remaining negative after the two PCRs approaches described above, PCRs with primers 16SWolbF and 16SWolbR3 were performed varying the following parameters: MgCl2 concentrations at 2.5, 4 and 6 mM and annealing temperatures of 52°C +/−5°C.

DNA preparations from filarial species harbouring Wolbachia (D. immitis and Brugia pahangi) and from a Wolbachia-infected strain of mosquitoes (Culex pipiens) [5] were included in the screening as positive controls. DNA preparations from a filarial species not harbouring Wolbachia (A. viteae) [5] were included as negative controls.

Of the samples positive for PCR screening homologous to Wolbachia 16S rDNA, dnaA, ftsZ and groEL were also amplified using the primers described in [5], [80] under the following conditions: 1x Eppendorf buffer including 1.5 mM MgCl2, 0.2 µM of each dNTP, 1 µM each of forward and reverse primers, and 0.5 units MasterTaq (Eppendorf). The thermal profiles we used were: (1) dnaA, 94°C 45 sec, 52°C 45 sec, and 72°C 90 sec, for 40 cycles; (2) groEL, 94°C 45 sec, 60°C 45 sec, 72°C 80 sec, for 5 cycles, and 94°C 45 sec, 55°C 45 sec, and 72°C 80 sec, for 34 cycles; (3) ftsZ, 94°C 30 sec, 60°C 45 sec, 72°C 90 sec, for 5 cycles, and 94°C 30 sec, 57°C 45 sec, and 72°C 90 sec, for 34 cycles.

Amplifications were performed in 20–50 µl volumes.

In all cases, in order to ascertain the DNA conditions before Wolbachia screenings and to confirm morphological identification, coxI amplification was performed as described in [12].

Sequencing conditions

coxI sequencing was performed as described in [12]. dnaA, ftsZ and groEL sequencing were performed as described in [5], [80]. From the Wolbachia PCR positive samples, almost the full length of the 16S rDNA gene of Wolbachia was sequenced using primers 27F and 16SWolbR3. The amplifications obtained (about 1400 bp) were gel-purified (using the QIAquick® PCR Purification Kit, Qiagen) and directly sequenced using ABI technology. The sequences obtained have been deposited in the EMBL Data Library. The dnaA, groEL and ftsZ sequences were not obtained from all the taxa included in this study mainly caused by the scarcity of certain specimens, and amplification/sequencing problems for some of the species examined. Where DNA of the host was amplified, the PCR product was purified as above and directly sequenced using ABI technology.

A list of the sequences including accession numbers is available in Table S2.

Immunohistochemical staining of worm sections

Immunohistochemical staining was performed according to [24], [48]. A rabbit polyclonal antiserum raised against the WSP of the Wolbachia from B. pahangi (designed by [81]) has been used to stain samples of 13 filarial species (see Tables S2 and S3). After paraffin inclusion, 4 µm sections were obtained and placed on Silane coated glass slides (3-aminpropyltriethoxysilane) and then kept at 63°C overnight, to avoid sections detaching from slides. To verify that the 63°C overnight did not alter the specific epitopes, Wolbachia positive L. sigmodontis laboratory strain were incubated at both 63°C and room temperature overnight and then stained: no significant differences in the staining were observed. Negative controls were carried out by omitting the primary antibody.

Fixed female worms were divided in three main parts, posterior (p), median (c) and anterior (a) in order to observe the different regions of the genital tract. Transverse sections were made at different levels of each part, and few of them were stained with hemalun-eosin for anatomical identification. Lateral cuticular internal crests were identified to orient the worm section; hypodermal chords extend above and on the side of the crests. The filarial species used for histology are opisthodelphic and the initial part of the ovaries is in the posterior part of the worm; the distal region of the ovary is composed of a cytoplasmic axis, the rachis [82], [83], and an outer cytoplasmic layer with the nuclei of oogonia and oocytes (in the text we will refer to these states as oocytes, and germline to describe the whole production of the gonad). An epithelial layer and an outer muscle layer surround the gonad, both referred to as somatic gonad in the text [25]. Uteri occupy almost the whole body and are found in median and anterior part of worms. The different uterine contents are ovulae, spermatozoa, divided eggs and microfilariae. Eggs were identified as aborted by hemalun-eosin staining when divided eggs were eosinophilic and nuclei not discernible. The laboratory strain of Litomosoides sigmodontis, which has been shown in several studies to harbour Wolbachia [84][86] was used as a positive reference for Wolbachia immunostaining.

Whole mount fluorescent analysis

Worms were cut with a razor blade to expose the different tissues to RNAse A (15 mg/mL, Sigma) in rotating tubes overnight at 4C. They were rinsed in PBS, and incubated with a fluorochrome-conjugated Phalloidin (atto−488 Phalloidin, Fluka, at 10 nanomolar) overnight to stain F-actin, followed by a Propidium Iodide (Molecular Probes, 1.0 mg/mL solution) incubation for DNA staining for 20 minutes in PBS (1∶50) and a 5 minutes wash. Tissues were mounted in Vectashield (Vector Laboratories) [25]. The species analyzed were C. crassa, C. japonica, Lo. caprini, M. (Cu.) perforata and O. d. japonica (Table S1).

Phylogenetic reconstruction

The bacterial 16S rDNA, dnaA, groEL and ftsZ sequences and filarial 12S rDNA and coxI sequences generated were aligned with the sequences available in the databases (for the ribosomal genes according to their secondary structures) using ClustalX2 [87].

The alignments were analysed using Maximum Likelihood (ML) and Bayesian Inference of phylogeny (BI) methods. The appropriate model of sequence evolution for ML and BI was estimated via likelihood ratio test using Modeltest 3.7 [88]: the model selected for the filarial concatenated dataset was HKY+G, while the model for the Wolbachia concatenated dataset was GTR+I+G. Phylogenetic analyses were performed using PAUP* 4.0 b10 [89] and MrBayes [90]. In addition, a phylogenetic tree was inferred with a GTR+Γ4 nucleotide substitution model in a Bayesian framework using MrBayes version 3.0 [90]. Two independent runs were performed, each using 1 million steps with four chains sampling every 100 steps. The first 10% of the trees were removed and posterior probabilities were calculated from these post-burnin trees.

Test for recombination

Four Wolbachia alignments (relative to the genes 16S rDNA, ftsZ, groEL and dnaA) were screened for presence of recombination events by using a set of nonparametric detection programs: RDP, GENECONV, Bootscan, MaxChi, Chimaera, SisScan and LARD [91][97]. The first six programs search for putative recombination breakpoints in a set of aligned DNA sequences and are implemented in RDP3 software package, whilst LARD checks signals detected by other methods [92]. Sequences were auto-masked for optimal recombination detection. General recombination settings were as follows: sequences were considered linear and the highest acceptable P-Value was set to 0.01 and overlapping signals were disentangled.

Method specific options were as follow: MaxChi and Chimaera were run with a variable window size; Bootscan and SisScan were forced for exploratory screening. Successive phases of refining analysis and manual tests were performed where needed.

Cocladogenesis analysis

To test for congruence, phylogenies for the hosts (based on a concatenated dataset of coxI and 12S rDNA) and for the bacterium (based on a concatenated dataset of 16S rDNA, ftsZ, dnaA and groEL) were compared with two methods. The first tested whether there was a greater than random correspondence between reconstructed nodes for host and symbiont. This was performed in Component (R. Page, University of Glasgow, UK) using the “compare tree with” function, with 1,000 randomized trees and the four available tree-comparison metrics (partition; triplets; quartets, nearest neighbour interchange). The second test examined the null hypothesis that the endosymbionts have undergone cocladogenesis with their hosts. ML trees for each dataset were first estimated using the successive approximation method [89]. Then, the scores for each of these ML trees based on the host dataset were compared using the [98] test. This was repeated using the parasite dataset. A single uninfected filarial (T. callipaeda) and a non-filarial Wolbachia (from C. quinquefasciatus) were used as outgroups.

Supporting Information

Figure S1.

Position of the genera screened in the present study indicated on a schematic representation of a key of the onchocercid subfamilies, based on morphological characters (following [46]). Total number of genera per subfamily listed. *Genus screened for the first time. ** Two subgenera in Mansonella.


Table S1.

Details of material studied with PCR, immunostaining assays (IHS) and whole mount fluorescent analysis (fluo). The scheme follows the classification as in Tables 1 and 2. Species, genus and subgenus, subfamily are in bold characters when newly screened; specimens ids are in bold characters when female worms; m = male; f = female; a = anterior part; c = central part; p = posterior part; *150 infective larvae; +  = Wolbachia positive specimens.


Table S2.

Results of Wolbachia screening based on PCR, immunostaining assays and whole mount fluorescent analysis in 35 filarial nematodes. Taxa are presented in alphabetical order.


Table S3.

Wolbachia distribution in the tissues of 13 onchocercid species. +: stained; −: not stained; NA: not available because the body structure is not present. *staining shown on Figures 14.



The authors would like to thank Andrea Galimberti and the personnel at ZooPlantLab of University of Milan-Bicocca for their help.

We are also grateful to Prof. H. Takaoka and Dr. M. Fukuda, Oita University, Japan for their contribution in providing animal hosts and Mr. T. Kenko, the Central Laboratory, Osaka City University Medical School, Osaka, Japan for histologic preparation of the parasites.

Author Contributions

Conceived and designed the experiments: MC OB. Performed the experiments: EF OB MB CM SU FL SGB RG SdSL SW MD MC. Analyzed the data: EF OB MB CM NL SU FL RG SdSL CB MC. Contributed reagents/materials/analysis tools: EF OB MB CM NL SU FL SGB RG CB SW MD. Wrote the paper: EF OB FL CB MC.


  1. 1. Werren JH, Zhang W, Guo LR (1995) Evolution and phylogeny of W. pipientis: reproductive parasites of arthropods. Proc R Soc Lond B 261: 55–71.
  2. 2. Sironi M, Bandi C, Sacchi L, Di Sacco B, Damiani G, et al. (1995) Molecular evidence for a close relative of the arthropod endosymbiont W. pipientis in a filarial worm. Mol Biochem Parasitol 74: 223–227.
  3. 3. Bandi C, Anderson TJC, Genchi C, Blaxter ML (1998) Phylogeny of W. pipientis in filarial nematodes. Proc R Soc Lond B 265: 2407–2413.
  4. 4. Taylor M J, Bandi C, Hoerauf A (2005) W. pipientis bacterial endosymbionts of filarial nematodes. Adv Parasitol 60: 245–284.
  5. 5. Casiraghi M, Bain O, Guerrero R, Martin C, Pocacqua V, et al. (2004) Mapping the presence of W. pipientis pipientis on the phylogeny of filarial nematodes: evidence for symbiont loss during evolution. Int J Parasitol 34: 191–203.
  6. 6. Fenn K, Conlon C, Jones M, Quail MA, Holroyd NE, et al. (2006) Phylogenetic relationships of the W. pipientis of nematodes and arthropods. PLoS Pathog 2(10): e94.
  7. 7. Hoerauf A, Rao RU (2008) Wolbachia: a bug's life in another bug, Issues in Infectious diseases. Es H. Zeichhardt, B. Mahy.
  8. 8. Lo N, Casiraghi M, Salati E, Mazzocchi C, Bandi C (2002) How many W. pipientis supergroups exist? Mol Biol Evol 19: 341–346.
  9. 9. Lo N, Paraskevopoulos C, Bourtzis K, O'Neill SL, Werren JH, et al. (2007) Taxonomic status of the intracellular bacterium Wolbachia pipientis. Int J Syst Evol Microbiol Mar 57: 654–657.
  10. 10. Wieatanaratanabutr I, Kittayapong P, Caubet Y, Bouchon D (2009) Molecular phylogeny of Wolbachia strains in arthropod hosts based on groEL-homologuous gene sequence. Zoo Sci 28(2): 171–177.
  11. 11. Fischer P, Schmetz C, Bandi C, Bonow I, Mand S, et al. (2002) Tunga penetrans: molecular identification of Wolbachia endobacteria and their recognition by antibodies against proteins of endobacteria from filarial parasites. Exp Parasitol 102: 201–211.
  12. 12. Casiraghi M, Favia G, Cancrini G, Bartoloni A, Bandi C (2001) Molecular identification of W. pipientis from the filarial nematode Mansonella ozzardi. Parasitol Res 87: 417–420.
  13. 13. Casiraghi M, Bordenstein SR, Baldo L, Lo N, Beninati T, et al. (2005) Phylogeny of W. pipientis pipientis based on gltA, groEL and ftsZ gene sequences: clustering of arthropod and nematode symbionts in the F supergroup, and evidence for further diversity in the W. pipientis tree. Microbiology 151: 4015–4022.
  14. 14. Ros VL, Fleming VM, Feil EJ, Breeuwer JA (2009) How diverse is the genus Wolbachia? Multiple-gene sequencing reveals a putatively new Wolbachia supergroup recovered from spider mites (Acari: Tetranychidae). Appl Environ Microbiol 75: 1036–1043.
  15. 15. Keiser PB, Coulibaly Y, Kubofcik J, Diallo AA, Klion AD, et al. (2008) Molecular identification of Wolbachia from the filarial nematode Mansonella perstans. Mol Biochem Parasitol 160: 123–128.
  16. 16. Haegeman A, Vanholme B, Jacob J, Vandekerckhove TT, Claeys M, et al. (2009) An endosymbiotic bacterium in a plant-parasitic nematode: Member of a new Wolbachia supergroup. Int J Parasitol 39: 1045–1054.
  17. 17. Rowley SM, Raven RJ, McGraw EA (2004) W. pipientis in Australian spiders. Curr Microbiol 49: 208–214.
  18. 18. Baldo L, Werren JH (2007) Revisiting W. pipientis supergroup typing based on WSP: spurious lineages and discordance with MLST. Cur Microbiol 55: 81–87.
  19. 19. Werren JH (1997) Biology of W. pipientis. Ann Rev Entomol 42: 587–609.
  20. 20. Bandi C, Dunn AM, Hurst GD, Rigaud , T (2001) Inherited microorganisms, sex-specific virulence and reproductive parasitism. Trends Parasitol 17: 88–94.
  21. 21. Fenn K, Blaxter M (2006) Wolbachia genomes: revealing the biology of parasitism and mutualism. Trends Parasit 22: 60–65.
  22. 22. Bordenstein SR, Paraskevopoulos C, Dunning Hotopp JC, Sapountzis P, Lo N, et al. (2009) Parasitism and mutualism in Wolbachia: what the phylogenomic trees can and cannot say. Mol Biol Evol 26: 231–241.
  23. 23. Brattig NW, Büttner DW, Hoerauf A (2001) Neutrophil accumulation around Onchocerca worms and chemotaxis of neutrophils are dependent on Wolbachia endobacteria. Microbes & Infection 3: 439–436.
  24. 24. Kramer LH, Passeri B, Corona S, Simoncini L, Casiraghi M (2003) Immunohistochemical/immunogold detection and distribution of the endosymbiont W. pipientis of Dirofilaria immitis and Brugia pahangi using a polyclonal antiserum raised against WSP (W. pipientis surface protein). Parasitol Res 89: 381–386.
  25. 25. Landmann F, Foster JM, Slatko B, Sullivan W (2010) Asymmetric Wolbachia segregation during early Brugia malayi embryogenesis determines its distribution in adult host tissues. PLoS Negl Trop Dis 4(7): e758.
  26. 26. Foster J, Ganatra M, Kamal I, Ware J, Makarova K, et al. (2005) The Wolbachia genome of Brugia malayi: endosymbiont evolution within a human pathogenic nematode. PLoS Biol 3: e121.
  27. 27. Heider U, Blaxter M, Hoerauf A, Pfarr K (2006) Differential display of genes expressed in the filarial nematode Litomosoides sigmodontis reveals a putative phosphate permease up-regulated after depletion of Wolbachia endobacteria. J Med Microbiol 296: 287–299.
  28. 28. Raverdy S, Foster JM, Roopenian E, Carlow CK (2008) The Wolbachia endosymbiont of Brugia malayi has an active pyruvate phosphate dikinase. Mol Biochem Parasitol 160: 163–166.
  29. 29. Foster J M, Raverdy S, Ganatra M B, Colussi P A, Taron C H, et al. (2009) The W. pipientis endosymbiont of Brugia malayi has an active phosphoglycerate mutase: a candidate target for anti-filarial therapies. Parasitol Res 104: 1047–1052.
  30. 30. Holmal AG, Davis PJ, Foster JM, Carlow CK, Kumar S (2009) Computational prediction of essential genes in an unculturable endosymbiotic bacterium, Wolbachia of Brugia malayi. BMC Microbiol 28: 243.
  31. 31. Johnston KL, Wu B, Guimarães A, Ford L, Slatko BE, et al. (2010) Lipoprotein biosynthesis as a target for anti-Wolbachia treatment of filarial nematodes. Parasit Vectors 3: 99.
  32. 32. Strübing U, Lucius R, Hoerauf A, Pfarr KM (2010) Mitochondrial genes for heme-dependent respiratory chain complexes are up-regulated after depletion of Wolbachia from filarial nematodes. Int J Parasitol 40: 1193–1202.
  33. 33. Hoerauf A (2008) Filariasis: new drugs and new opportunities for lymphatic filariasis and onchocerciasis. Curr Opin Infect Dis 21: 673–81.
  34. 34. Bordenstein SR, Fitch DH, Werren JH (2003) Absence of Wolbachia in nonfilariid nematodes. J Nematol 35: 266–270.
  35. 35. Casiraghi M, McCall JW, Simoncini L, Kramer LH, Sacchi L, et al. (2002) Tetracycline treatment and sex-ratio distortion: a role for W. pipientis in the moulting of filarial nematodes? Int J Parasitol 32: 1457–1468.
  36. 36. Büttner D W, Wanji S, Bazzocchi C, Bain O, Fischer P (2003) Obligatory symbiotic W. pipientis endobacteria are absent from Loa loa. Filaria J 2: 10.
  37. 37. Dunning Hotopp JC, Clark ME, Oliveira DC, Foster JM, et al. (2007) Widespread lateral gene transfer from intracellular bacteria to multicellular eukaryotes. Science 317: 1753–1756.
  38. 38. McNulty SN, Foster JM, Mitreva M, Dunning Hotopp JC, Martin J, et al. (2010) Endosymbiont DNA in endobacteria-free filarial nematodes indicates ancient horizontal genetic transfer. PLoS One 5(6): e11029.
  39. 39. Turner JD, Langley RS, Johnston KL, Gentil K, Ford L, et al. (2009) Wolbachia lipoprotein stimulates innate and adaptive immunity through toll-like receptors 2 and 6 to induce disease manifestations of filariasis. J Biol Chem 284: 22364–22378.
  40. 40. Hansen RD, Trees AJ, Bah GS, Hetzel U, Martin C, et al. (2010) A worm's best friend: recruitment of neutrophils by Wolbachia confounds eosinophil degranulation against the filarial nematode Onchocerca ochengi. Proc Biol Sci. Epub ahead of print.
  41. 41. Kambris Z, Cook PE, Phuc HK, Sinkins SP (2009) Immune Activation by life-shortening Wolbachia and reduced filarial competence in mosquitoes. Science 326: 134–136.
  42. 42. Kambris Z, Blagborough AM, Pinto SB, Blagrove MSC, Godfray HCJ, et al. (2010) Wolbachia stimulates immune gene expression and inhibits plasmodium development in Anopheles gambiae. PLoS Pathog 6: e1001143.
  43. 43. Hughes GL, Ren X, Ramirez JL, Sakamoto JM, Bailey JA, et al. (2011) Wolbachia infections in Anopheles gambiae cells: transcriptomic characterization of a novel host-symbiont interaction. PLoS Pathog 7: e1001296.
  44. 44. Chabaud AG, Bain O (1994) The evolutionary expansion of the Spirurida. Int J Parasitol 24: 1179–1201.
  45. 45. Bain O, Casiraghi M, Martin C, Uni S (2008) The Nematoda Filarioidea: critical analysis linking molecular and traditional approaches. Parasite 15: 342–348.
  46. 46. Anderson RC, Bain O Anderson RC, Chabaud AG, Willmott S, editors. (2009) Spirurida. Diplotriaenoidea, Aproctoidea and Filarioidea. Archival Volume.CAB International, Wallingford U K: 391–448.
  47. 47. Gibbons LM (2010) Keys to the Nematode Parasites of Vertebrates. Supplementary volume. CAB International, Wallingford U K: 416.
  48. 48. Junker K, Barbuto M, Casiraghi M, Martin C, Boomker J, et al. (2009) Litomosa chiropterorum Ortlepp, 1932 (Nematode: Filarioidea) from a South African miniopterid: redescription, Wolbachia screening and phylogenetic relationships with Litomosoides. Parasite 16: 43–50.
  49. 49. Vuong PN, Wanji S, Sakka L, Klager S, Bain O (1991) The murid filarial Monanema martini: a model for onchocerciasis. Part I Description of lesions. Ann Parasitol Hum Comp 66: 109–120.
  50. 50. Uni S, Bain O, Agatsuma T, Katsumi A, Baba M, et al. (2006) New filaria nematode from Japanese serows (Naemorhedus crispus: Bovidae) close to parasites from elephants. Parasite 13: 193–200.
  51. 51. Keiser PB, Coulibaly Y, Kubofcik J, Diallo AA, Klion AD, et al. (2008) Molecular identification of Wolbachia from the filarial nematode Mansonella perstans. Mol Biochem Parasitol 160: 123–128.
  52. 52. Taylor MJ, Bilo K, Cross HF, Archer JP, Underwood AP (1999) 16S rDNA phylogeny and ultrastructural characterization of Wolbachia intracellular bacteria of the filarial nematodes Brugia malayi, B. pahangi, and Wuchereria bancrofti. Exp Parasitol 91: 356–61.
  53. 53. Ferri E, Barbuto M, Bain O, Galimberti A, Uni S, et al. (2009) Integrated taxonomy: traditional approach and DNA barcoding for the identification of filarioid worms and related parasites (Nematoda). Frontiers in Zool 6: 1–12.
  54. 54. Uni S, Bain O, Takaoka H, Fujita H, Suzuki Y (2002) Diversification of Cercopithifilaria species (Nematoda Filarioidea) in Japanese wild ruminants with description of two new species. Parasite 9: 293–304.
  55. 55. Bain O, Chabaud A (1986) Atlas des larves infectantes de Filaires. Trop Med Parasitol 37: 237–340.
  56. 56. Bain O, Baker M, Chabaud A G (1982) Nouvelles données sur la lignée Dipetalonema (Filarioidea, Nematoda). Ann Parasitol Hum Comp 57: 593–620.
  57. 57. Agatsuma T, Iwagami M, Uni S, Takaoka H, Katsumi A, et al. (2005) Molecular phylogenetic relationships among seven Japanese species of Cercopithifilaria. Parasitol Int 54: 195–199.
  58. 58. Uni S (1983) Filarial parasites of the black bear of Japan. Ann Parasit Hum Comp 58: 71–84.
  59. 59. Attout T, Babayan S, Hoerauf A, Taylor DW, Kozek WJ, et al. (2004) Blood-feeding in the young adult filarial worms Litomosoides sigmdontis. Parasitology 130: 421–428.
  60. 60. Manzanell R (1986) Oswaldofilaria kanbaya n. sp., a new filarioid worm (Nematoda, Filarioidea). Ann Parasit Hum Comp 61: 245–254.
  61. 61. Saint André AV, Blackwell NM, Hall LR, Hoerauf A, Brattig NW et al (2002) The role of endosymbiotic W. pipientis bacteria in the pathogenesis of river blindness. Science 295: 1892–1895.
  62. 62. Brattig NW, Bazzocchi C, Kirschning CJ, Reiling N, Büttner DW et al (2004) The major surface protein of W. pipientis endosymbionts in filarial nematodes elicits immune responses through TLR2 and TLR4. J Immunol 173: 437–445.
  63. 63. Gillette-Ferguson I, Daehnel K, Hise AG, Sun Y, Carlson E, et al. (2007) Toll-like receptor 2 regulates CXC chemokine production and neutrophil recruitment to the corner in Onchocerca volvulus/Wolbachia-induced keratitis. Infect Immun 75: 5908–5915.
  64. 64. Higazi T B, Filiano A, Katholi CR, Dadzie Y, Remme JH et al (2005) Wolbachia endosymbiont levels in severe and mild strains of Onchocerca volvulus. Mol Biochem Parasitol 141: 109–112.
  65. 65. Bain O (2002) Evolutionary relationships among filarial nematodes. 21–29. In: World Class Parasites, Kluwer Acad Publ, vol 5, The Filaria(Eds.Klei & Rajan).
  66. 66. Blaxter M (2009) Nematodes: the worm and their relatives. PLoS Biology 9: e1001050.
  67. 67. Chirgwin SR, Porthouse KH, Nowling JM, Klei TR (2002) The filarial endosymbiont W. pipientis sp. is absent from Setaria equina. J Parasitol 88: 1248–1250.
  68. 68. Bain O, Bartlett C, Petit G (1986) Une filaire de Muridés africains dans la paroi du côlon, Monanema martini n. sp. Ann Parasit Hum Comp 61: 465–472.
  69. 69. Bain O, Petit G, Chabaud AG (1986) Une nouvelle filaire, Cercopithifilaria roussilhoni n. sp., parasite de l'Athérure au Gabon, transmise par tiques: hypothèse sur l'évolution du genre. Ann Parasit Hum Comp 61: 81–93.
  70. 70. Guerrero R, Martin C, Bain O (2003) Litomosoides yutajensis n. sp., first record of this filarial genus in a mormoopiid bat. Parasite 10: 219–225.
  71. 71. Uni S, Bain O, Takaoka H (2004) Affinities between Cutifilaria (Nematoda: Filarioidea), parasite of deer, and Mansonella as seen in a new onchocercid, M. (C.) perforata n. sp., from Japan. Parasite 11: 131–140.
  72. 72. Uni S, Bain O, Agatsuma K, Harada M, Torii H, et al. (2007) Onchocerca eberhardi n. sp. (Nematoda: Filarioidea) from the sika deer in Japan; relationships between species parasitic in cervids and bovids in the Holarctic region. Parasite 14: 199–211.
  73. 73. Guerrero R, Bain O, Martin C, Barbuto M (2011) A new species of Litomosoides (Nematoda: Onchocercidae) parasite of Nectomys palmipes (Rodentia: Cricetidae: Sigmodontinae) from Venezuela: description, molecular evidence, Wolbachia pipientis screening. Folia Parasitologica 58, in press.
  74. 74. Chabaud AG, Bain O (1976) La lignée Dipetalonema. Nouvel essai de classification. Ann Parasit Hum Comp 51: 365–397.
  75. 75. Eberhard ML, Orihel TC (1984) The genus Mansonella (Syn. Tetrapetalonema). A new classification. Ann Parasitol Hum Comp 59: 483–496.
  76. 76. O'Neill SL, Gooding RH, Aksoy S (1993) Phylogenetically distant symbiotic microorganisms reside in Glossina midgut and ovary tissues. Med Vet Entomol 7: 377–383.
  77. 77. Simoes PM, Mialdea G, Reiss D, Sagot MF, Charlat S (2011) Wolbachia detection: an assessment of standard PCR protocols. Mol Ecol Res. In Press.
  78. 78. Lane DJ Stackebrandt E, Goodfellow M, editors. (1991) 16S/23S rRNA sequencing.New York: 115–175. John Wiley & Sons.
  79. 79. Werren JH, Windsor DM (2000) W. pipientis infection frequencies in insects: evidence of a global equilibrium? Proc R Soc Lond, B 267: 1277–1285.
  80. 80. Casiraghi M, Werren JH, Bazzocchi C, Biserni A, Bandi C (2003) dnaA gene sequences from W. pipientis support subdivision into supergroups and provide no evidence for recombination in the lineages infecting nematodes. Parassitologia 45: 13–18.
  81. 81. Bazzocchi C, Jamnongluk W, O'Neill S, Anderson TJC, Genchi C, et al. (2000) wsp gene sequences from the Wolbachia of filarial nematodes. Curr Microbiol 41: 96–100.
  82. 82. Chitwood BG, Chitwood MB (1950) An introduction to nematology. Section 1. Anatomy. Baltimore, USA: Monumental Printing Company.
  83. 83. Justine J-L (2002) Male and female gametes and fertilization. 73–119.
  84. 84. Hoerauf A, Nissen-Paehle K, Schmetz C, Henkle-Dürsen K, Blaxter ML et al (1999) Tetracycline therapy targets intracellular bacteria in the filarial nematode Litomosoides sigmodontis and results in filarial infertility. J Clin Invest 103: 11–17.
  85. 85. Volkman L, Fisher K, Taylor M, Hoerauf A (2003) Antibiotic therapy in murine filariasis (Litomosoides sigmodontis): comparative effects of doxycycline and rifampicin on W. pipientis and filarial viability. Trop Med Intern Health 8: 392–401.
  86. 86. Attout T, Martin C, Babayan SA, Kozek WJ, Bazzochi C, et al. (2008) Pleural cellular reaction to the filarial infection Litomosoides sigmodontis is determined by the moulting process, the worm alteration, and the host strain. Parasitol Intern 57: 201–211.
  87. 87. Larkin MA, Blackshields G, Brown NP, Chenna R, McGettigan PA, et al. (2007) Clustal W and Clustal X version 2.0. Bioinformatics 23: 2947–2948.
  88. 88. Posada D, Crandall KA (1998) Modeltest: testing the model of DNA substitution. Bioinformatics 14: 817–818.
  89. 89. Swofford DL (1998) PAUP, phylogenetic analysis using parsimony (*and other methods): version 4. Sunderland, Massachusets, USA: Sinauer Associates.
  90. 90. Ronquist F, Huelsenbeck JP (2003) MRBAYES 3: Bayesian phylogenetic inference under mixed models. Bioinformatics 19: 1572–1574.
  91. 91. Martin DP (2009) Recombination detection and analysis using RDP3. Methods Mol Biol 537: 185–205.
  92. 92. Padidam M, Sawyer S, Fauquet CM (1999) Possible emergence of new geminiviruses by frequent recombination. Virology 265: 218–225.
  93. 93. Martin DP, Posada D, Crandall KA Williamson C (2005) A modified bootscan algorithm for automated identification of recombinant sequences and recombination breakpoints. AIDS Res Hum Retroviruses 2: 98–102.
  94. 94. Maynard Smith J (1992) Analyzing the mosaic structure of genes. J Mol Evol 34: 126–129.
  95. 95. Posada D, Crandall KA (2001) Evaluation of methods for detecting recombination from DNA sequences: Computer simulations. Proc Natl Acad Sci 98: 13757–13762.
  96. 96. Gibbs MJ, Armstrong JS, Gibbs , AJ (2000) Sister-Scanning: a Monte Carlo procedure for assessing signals in recombinant sequences. Bioinformatics 16: 573–582.
  97. 97. Holmes EC, Worobey M, Rambaut A (1999) Phylogenetic evidence for recombination in dengue virus. Mol Biol Evol 16: 405–409.
  98. 98. Shimodaira H, Hasegawa M (1999) Multiple comparisons of log-likelihoods with applications to phylogenetic inferences. Mol Biol Evol 16: 1114–1116.