Wolbachia: endosymbiont of onchocercid nematodes and their vectors.

BACKGROUND: Wolbachia is an obligate intracellular maternally transmitted, gram-negative bacterium which forms a spectrum of endosymbiotic relationships from parasitism to obligatory mutualism in a wide range of arthropods and onchocercid nematodes, respectively. In arthropods Wolbachia produces reproductive manipulations such as male killing, feminization, parthenogenesis and cytoplasmic incompatibility for its propagation and provides an additional fitness benefit for the host to protect against pathogens, whilst in onchocercid nematodes, apart from the mutual metabolic dependence, this bacterium is involved in moulting, embryogenesis, growth and survival of the host.
METHODS: This review details the molecular data of Wolbachia and its effect on host biology, immunity, ecology and evolution, reproduction, endosymbiont-based treatment and control strategies exploited for filariasis. Relevant peer-reviewed scientic papers available in various authenticated scientific data bases were considered while writing the review.
CONCLUSIONS: The information presented provides an overview on Wolbachia biology and its use in the control and/or treatment of vectors, onchocercid nematodes and viral diseases of medical and veterinary importance. This offers the development of new approaches for the control of a variety of vector-borne diseases.

Background

Endosymbiosis is an intimate form of symbiotic association in which one organism dwells within the body of another, forming a spectrum of relationships from parasitism to obligatory mutualism [1]. Many obligate mutual symbiotic associations are based on metabolic complementation and strengthen or increment the biochemical versatility and pathways of one or both hosts [2, 3]. Wolbachia is a striking example of this mechanism in both onchocercid nematodes and arthropod vectors [4]. Though many endosymbionts have been observed in arthropod and nematode hosts, Wolbachia is the one which is most widely distributed and explored [4]. Like the mitochondria organelle, this obligate intracellular gram-negative bacterium is also transmitted through the host germ line to the next generation [5]. After the initial discovery in the reproductive organs of Culex pipiens mosquito by M. Hertig and S. B. Wolbach in 1924 [6], the description of this bacterium took another 12 more years [7]. In the late 1960s and early 1970s, ultrastructural studies on filarial nematodes revealed the presence of unusual intracellular bodies in the oocyte hypodermis of these worms, which were interpreted as bacteria [8-10] and later identified as Wolbachia by Sironi and colleagues [11]. Currently, this endosymbiont has been reported in around 50% of terrestrial arthropod species (i.e. insects, mites, crustaceans, spiders, scorpions, collembolans) and in several species of onchocercid nematodes [12, 13]. Similarly, this bacterium has also been identified in non-filarial plant nematodes, Radopholus similis [14] and Pratylenchus penetrans of the order Tylenchida [15]. In onchocercid nematodes, Wolbachia has a mutual association in which it is involved in embryogenesis, moulting, growth and survival [16] of the filariae, and it has been hypothesised that the worm provides essential aminoacids for bacterial growth [17]. However, in arthropods, the “parasitic side” of Wolbachia prevails, in that this bacterium manipulates the host reproduction to increase its own fitness and spread into the host population [18, 19]. Reproductive manipulations exerted by Wolbachia on its hosts have extensively been investigated and include male killing, feminization, parthenogenesis and cytoplasmic incompatibility (CI) [4]. The effects of Wolbachia presence on its hosts (i.e. host biology, physiology, immunity, ecology, evolution and reproduction) have been exploited for the development of promising endosymbiont-based strategies for the treatment of filariasis and for the control of important vector-borne diseases of medical and veterinary relevance [4]. This review details Wolbachia's evolution, molecular identification, interaction with arthropods and nematodes and the development of endosymbiont-based treatment and control strategies.

Evolutionary history of Wolbachia

Wolbachia evolved and adapted to its intracellular lifestyle in the context of an evolutionary change that included other obligatory intracellular organisms (such as the ancestors of Rickettsia, Ehrlichia, Anaplasma and Midichloria) and extended over a hundred million years, starting from ancient alphaproteobacteria [20-22]. Wolbachia has a small genome (0.8–1.7 Mbp) with large segments of mobile and repetitive DNA, which is uncommon in vertically transmitted (generally from mother to offspring) organisms [4, 23]. Despite the erosive genomic processes due to host restriction and acquisition maintenance, these repetitive host DNA sequences are supposed to play a major role in the evolution of Wolbachia [24]. Balance among vertical transmission, host switching, recombination insertion sequences and bacteriophage sequences helps in the adaptation and global distribution of Wolbachia [4, 22]. Based on their main genetic evolution in a large variety of hosts, Wolbachia have been classified in 17 supergroups, designated by the letters A to S [23, 25, 26]. Exceptions are represented by supergroup G, which was lately been withdrawn because of the high probability of being the result of a genetic recombination event [27, 28], and supergroup R from cave spiders [29], which showed a strong association with Wolbachia strains of supergroup A, based on genetic distance measures and phylogenetic analyses [30]. Overall, Wolbachia of arthropods is categorised in supergroups A, B, E, H, I, K, that of nematodes in C, D, J [31] and supergroup L only in plant-parasitic nematodes [15]. Supergroup F is an exception which is common in many arthropod species such as termites, spiders, mites [32], bugs (i.e. Cimex lectularius and Montina sp.) [33] and in human filariae (i.e. Mansonella) [34-36], filariae of black bear, (Cercopithifilaria japonica) [12] and that of geckoes (Madathamugadia hiepei) [37]. Currently, a complete genome of Wolbachia from the supergroup F is available from the wCle strain of C. lectularius [2] and wMhie strain of M. hiepei [23].

In particular, supergroups A and B are the most represented among arthropods and it is estimated that the common ancestor of both would have diverged approximately 58–67 million years ago. Though estimation of the origin of Wolbachia is a controversial topic and a suitable outgroup for phylogenetic analysis of Wolbachia is unavailable, it has been suggested that arthropod and onchocercid nematode supergroups diverged around 100 million years ago (i.e. 500 million years after their host) [38, 39]. It is important to note that these estimations were based on small samplings. The presence of Wolbachia in phylogenetically distant hosts such as nematodes and arthropods suggests that these endosymbionts experienced some type of horizontal transmission during their ancient evolution. For example, a horizontal transfer of Wolbachia could have occurred from one host phylum to the other; alternatively, one of the two phyla could have acquired Wolbachia from a third party [39]. Incongruence in phylogenies of Wolbachia and their arthropod hosts (e.g. the unnatural occurrence of identical Wolbachia strains in distantly related species) can also be explained by the horizontal transmission of these endosymbionts [38, 40, 41]. In addition, ecological events occurring in the transmission and global distribution of this bacterium in arthropods include the relationship between and amongst hosts, such as in the case of parasitism, phoresis [42], predation and cannibalism [43], blood contact after injury [44], presence in parasitoids [45] or just sharing of common food substrates [46].

Unlike arthropods, phylogenetic congruence of nematodes with Wolbachia indicates an obligatory dependent relationship with the organism, followed by host-parasite co-evolution and vertical transmission via infected females [34, 39]. Though the association between Wolbachia and nematodes has been hypothesised to have been acquired as a single event [34, 39], recent genome analyses suggest multiple events of acquisition of Wolbachia with local coevolutionary patterns in different major lineages and wider presence of transposable elements in supergroup D (i.e. in Wolbachia from the Onchocerca genus) [13, 23]. Moreover, different patterns of symbiosis among various filarial nematodes may be due to multiple acquisitions of the bacteria and/or selective pressures imposed on it [23]. The strongest coevolution pattern has been observed in Onchocerca spp. especially in Onchocerca lupi, Onchocerca gutturosa, Onchocerca lienalis, Onchocerca volvulus and Onchocerca ochengi, strongly supported by global fit analyses [13]. Furthermore, the detection of Wolbachia-like gene transcript in Onchocerca fluxosa, which in turn is the only known Onchocerca species devoid of Wolbachia, suggests the ancestral presence of this symbiont in this nematode [47, 48]. Accordingly, the absence of Wolbachia in anuran onchocercid nematodes indicates that they would have diversified before the first bacterial invasion in onchocercid lineage (i.e. around 110 million years ago) [25, 39, 49]. However, the presence of supergroup F in both insects and filariae of humans, black bear and geckoes makes understanding of the whole picture even more complicated.

Molecular detection and identification of Wolbachia

Though the genus Wolbachia has a relatively small genome (i.e. 0.8–1.7 Mbp) it encompasses large phylogenetic variations [23, 39]. Amongst target genes, 16S rRNA showed a nucleotide divergence from 0.2% to 2.6% [50] but provided limited information for inferring phylogenetic relationships [50]. Hence, the genetic characterisation using 16S rRNA is complemented by a set of housekeeping genes (e.g. FtsZ, groEL, gltA and coxA) mainly for the phylogenetic analysis [31] (Table 1). Wolbachia surface protein (wsp) gene, ten times more variable than 16S rRNA and FtsZ, is employed to identify different groups and strains of Wolbachia [51, 52] but not for large-scale phylogenetic analysis since it is affected by recombination amongst supergroups [28]. Moreover, groE is also used for strain differentiation because of the faster evolution rate of non-coding regions that separate the coding heat shock protein (HSP) genes (i.e. groES and groEL) [53]. Therefore, PCR coupled sequencing of a combination of genes should be employed to assess the group relationships in Wolbachia [54]. It has been estimated that multiple infections can be detected by techniques such as quantitative PCR with highly specific primers [55, 56], cloning and sequencing [57], and southern hybridization [58]. Similarly, loop-mediated isothermal amplification (LAMP) is used in resource-limited laboratories for the simultaneous detection of more than one strain of Wolbachia [59]. A metagenomics-based approach can be employed to provide whole-genome sequence information for all associated endosymbionts of a nematode or an arthropod vector [60-63].

Table 1

Molecular approaches for the detection of Wolbachia in vectors and onchocercid nematodes

Primer name	Gene targeted	Type of PCR	Product size	References
99F, 99R	16S rRNA	cPCR	895	[50, 164]
Wspecf, Wspecr	16S rRNA	cPCR	438	[165]
16SWolbf, 16SWolbr	16S rRNA	cPCR	1014	[34]
INTF1, INTR	16S rRNA	cPCR	130	[166]
INTF2, INTR2	16S rRNA	cPCR	136	[166]
553F_W, 1334R_W	16S rRNA	cPCR	781	[54]
WolbF, Wspecr	16S rRNA	cPCR		[167]
63f, 1387R, 76f, 1012R	16S rRNA	Nested PCR	852	[168]
WN16S-F, WN16S-R	16S rRNA	qPCR		[169]
W-Specf, W-Specr,	16S rRNA	qPCR	438	[170]
W-Specf, W16S			102
WSPintF, WSPintR	wsp	cPCR	576	[34, 114, 168]
81F, 691R	wsp	cPCR	610	[51, 171]
136 F, 691R	wsp (Group A)		556
308 F, 691R	wsp (Subgroup Mel)		405
328 F, 691R	wsp (Subgroup AlbA)		379
173F, 691R	wsp (Subgroup Mel and AlbA)		541
181F, 691R	wsp (Subgroup wPap)		506
165F, 691R	wsp (Subgroup wAus)		506
81F, 531R	wsp (Subgroup wPap and wAus)		460
81 F, 522R	wsp (Group B)		442
183F, 691R	wsp (Subgroup Pip)		501
wspF, wspR, grF, grR	wsp	qPCR		[172]
wspTMF, wspTMR	wsp	qPCR		[59]
WSP.F3, WSP.B3, WSP.FIP, WSP.BIP	wsp	LAMP assay		[173]
FIP_wMel/wPop	wsp (wMel/ wPop)	LAMP assay		[59]
BIP_wMel/wPop
F3_wMel/wPop
B3_wMel/wPop
LpF_wMel/wPop
LpB_wMel/wPop
ftsZ_F1, ftsZ_R1	FtsZ	cPCR	524	[40]
FtsZUniF, FtsZUniR	FtsZ	cPCR		[174]
ftsZfl, ftsZrl	FtsZ	cPCR	1043–1055	[38]
Wol1F, Wol1R, Wol7F, Wol7R	FtsZ	Nested PCR	147	[175]
MLST primers	16 s rRNA, gatB, FtsZ, hcpA, fbpA coxA. wsp	MLST		[40]
ftsZ 291, ftsZ 477	FtsZ	qPCR		[176]
WSP 420, WSP 583	wsp
Bm-wFtsZ-F, BmwFtsZ-R	FtsZ	qPCR		[96]
groEL-F, groEL-R	groEL	cPCR		[32]
WgroF1, WgroRev1	groEL	cPCR	873	[35]
WgltAF1, WgltARev1	Citrate synthase (gltA)	cPCR	627	[35]
FbpA_F1, FbpA_R1	FbpA	cPCR	509	[40]
hcpA_F1, hcpA_R1	HcpA	cPCR	516	[40]
coxA_F1, coxA_R1	coxA	cPCR	487	[40]
COIintF, COIintR	COI	cPCR	689	[34]
Wseq01F, Wseq02R	gatB	cPCR	471	[31]

To date, eight complete genomes (i.e. wBm of Brugia malayi, wBp of Brugia pahangi, wOo of Onchocerca ochengi, wOv of Onchocerca volvulus, wDimm, Dirofilaria immitis, wCtub of Cruorifilaria tuberocauda, wDcau of Dipetalonema caudispina, wLsig of Litomosoides sigmodontis) and three draft genomes of Wolbachia from filarial nematodes have been published (i.e. wLbra of Litomosoides brasiliensis, wWb of Wuchereria bancrofti and wMhie of Madathamugadia hiepei) [17, 23, 64–67]. Of the 36 complete genomes and 55 draft genomes of Wolbachia available, 84% belong to supergroups A and B [23]. Advanced genome analyses on Wolbachia suggest that supergroups A and B were originated by genetic isolation events rather than convergent evolution [68]. As per genome analyses, the relationship between onchocercid nematodes and Wolbachia may represent a “genetic addiction” rather than mutualism [1]. Compared to arthropods, the filarial nematode genome has smaller size (i.e. 863,427 bp for wDcau versus 1,267,782 bp for wMel from Drosophila melanogaster or 1,801,626 bp for wFol from Folsomia candida), presence of fewer transposable elements as insertion sequence elements (ISs) and group II intron-associated genes, prophage-related genes and repeat-motif proteins as ankyrin domains [23]. Data analyses on intragenomic recombinations, transposable elements, chromosome rearrangements, mutational bias and gene loss or gain on different supergroups revealed that supergroup C strains have a very low number of genomic rearrangements, paucity of insertion sequence elements and strong GC asymmetric distribution, which is considered to be due to the long-term obligate symbiotic relationship with their host [69]. Further addition of the genome of new Wolbachia strains from different filarial nematodes will help to do detailed analyses and to have a clear picture on the divergent symbiotic mechanisms and the evolutionary pattern of this bacterium.

Wolbachia in onchocercid nematode vectors

First reported in Cx. pipiens [7], Wolbachia is a widespread endosymbiont among arthropods with an estimated prevalence ranging from 20 to around 75%, according to different studies [38, 70]. Indeed, arthropods have been found to be infected with single (e.g. C. lectularius) or multiple Wolbachia variants (e.g. Drosophila simulans, Cx. pipiens) in the same species or even in the same insect individuals (i.e. superinfection) [71-74] (Table 2). Unlike onchocercid nematodes, arthropod-Wolbachia association is more parasitic, in that the bacteria obtain fitness advantage by the reproductive manipulations of the host [16, 75–77]. Apart from these, the bacterium is involved in iron homeostasis of the host and confers immunity to viral/onchocercid nematode infections, thereby reducing the vector capacity of their hosts [78-80]. This has been demonstrated in Aedes aegypti infected by Wolbachia popcorn strain (wMelPop) wherein the symbiosis conferred protection to the mosquitoes against onchocercid nematodes and Plasmodium gallinaceum parasites [79, 81]. The wide host range, tissue distribution and ability to perpetuate in insect populations render Wolbachia very attractive as a tool to reduce the vector potentiality of their host [82, 83] also thanks to its capability to determine reproductive manipulations (e.g. CI, parthenogenesis, male killing, feminization and meiotic drive) [84, 85]. By these phenotypic alterations, the symbiont gives more reproductive advantage to infected over uninfected individuals or genders [63]; in general, Wolbachia is more beneficial for the individuals of the female sex, by virtue of the matriline vertical transmission [63]. In males, Wolbachia affects genes involved in sex differentiation and development by altering the DNA methylation whereas in females it might interfere with steroid hormone signalling [86]. In addition to this, Wolbachia wMelPop infection in Ae. aegypti mosquitoes produces hypomethylation/demethylation of genomic DNA, affecting 699 genes involved in apoptosis, defence response, phagocytosis, circadian rhythm, life span and locomotion behaviour [87-89]. The above changes may be capitalized as a mean for controlling vector-borne diseases [90] through life-shortening and increased locomotor activity and metabolism of mosquito vectors [90]. Overall, the above factors may indeed alter the mosquito-pathogen relationship reducing the vectorial capacity of mosquitoes. In addition, the ability of Wolbachia to inhibit the replication of viral RNA can be used to suppress virus infection by transinfecting mosquitoes with proper Wolbachia strains [78, 91]. Therefore, though the exact mechanism of such inhibition is not known, Wolbachia can be used as a biocontrol agent to effectively control viral pathogens such as dengue, yellow fever and chikungunya viruses and other vector-borne pathogens such as filarial nematodes and the malaria parasite, P. gallinaceum [92-94].

Table 2

Main supergroups and subgroups of Wolbachia detected in vectors

Arthropod	Vector	Supergroup detected	Subgroup	Gene targeted	Reference
Aedes	Yellow fever virus, dengue (D1, D2, D3, D4) viruses, chikungunya virus, zika virus (ZIKV), Rift Valley fever virus, Wuchereria bancrofti, Brugia malayi
Ae. albopictus		A, B	AlbA, Pip	wsp	[83, 169, 177]
Ae. albotaeniatus		A	Uni, Albo	wsp	[83, 178]
Ae. aegypti		A, B	-	wsp, 16S rRNA	[167]
Ae. pseudoalbopictus			AlbA, Pip, Pseu	wsp	[83, 178]
Ae. (Stegomyia) spp.		A, B	AlbA, Pip	wsp	[83]
Ae. niveus subgroup A		A	Mel, Niv	wsp	[83, 178]
Ae. novoniveus		A	Riv, Uni, Nov	wsp	[83, 178]
Ae. craggi		B	CauB, Crag	wsp	[83, 178]
Ae. perplexus		B	CauB, Perp	wsp	[83, 178]
Ae. pseudalbopictus		A, B	AlbA, Pip	wsp	[83]
Ae. cinereus		B		wsp	[177]
Ae. cantans		B	_	wsp	[177]
Ae. cinereus		C	Di	16S rRNA	[179, 180]
Ae. detritus		C	Di	wsp	[181]
Ae. geniculatus		C	Di	wsp	[181]
Ae. punctor		B, C	Pip, Di	wsp	[181]
Ae. fluviatilis		B	Flu	16S rRNA	[182]
Ae. cooki  Ae. polynesiensis  Ae. riversi		-	-	16S rRNA	[179, 180]
Ae. bromeliae		A	-	wsp	[183]
Ae. metallicus		A	-	wsp	[183]
Armigeres	Japanese encephalitis virus, Wuchereria bancrofti, Dirofilaria immitis, Brugia pahangi			wsp
Ar. kesseli		B	Pip	wsp	[83]
Ar. subalbatus		A	Riv, AlbA	wsp	[83, 171]
Ar. flavus		A	AlbA	wsp	[83]
Anopheles	Plasmodium falciparum, P. vivax P. malariae, P. ovale, P.knowlesi, Wuchereria bancrofti, Brugia malayi
An. funestus		A, B	Anfu	coxA, fbpA and FtsZ	[147, 184]
An. gambiae		A, B	Anga-BF, Anga-Mali	16S rRNA	[143, 185, 186]
An. coluzzii		A, B	Anga-BF, Anga-Mali	16S rRNA	[143, 185, 186]
An. arabiensis		A, B	Anga	16S rRNA	[143]
An. carnevalei		A, B	–	coxA, fbpA and FtsZ	[184]
An. coustani		B	–	coxA, fbpA and FtsZ	[184]
An. hancocki		B	–	coxA, fbpA and FtsZ	[184]
An. implexus		B	–	coxA, fbpA and FtsZ	[184]
An. jebudensis		B	–	coxA, fbpA and FtsZ	[184]
An. marshallii		B	–	coxA, fbpA and FtsZ	[184]
An. moucheti		B	–	coxA, fbpA and FtsZ	[184]
An. nigeriensis		B	–	coxA, fbpA and FtsZ	[184]
An. nili		B	–	coxA, fbpA and FtsZ	[184]
An. paludis		B	–	coxA, fbpA and FtsZ	[184]
An. vinckei		A, B	–	coxA, fbpA and FtsZ	[184]
An. minimus		F, D	–	16S rRNA	[170]
An. baimaii		D, F	–	16S rRNA	[170]
An. maculatus		B, F	–	16S rRNA	[170]
An. pseudowillmori		B	–	16S rRNA	[170]
An. sawadwongporni		B	–	16S rRNA	[170]
An. minimus		D	–	16S rRNA	[170]
An. dirus		B	–	16S rRNA	[170]
Culex	West Nile virus, equine encephalitis virus, Japanese encephalitis virus, Saint Louis encephalitis virus, Rift Valley fever virus, Wuchereria bancrofti, Brugia malayi
Cx p. pipiens		B	Pip	16S rRNA	[51]
Cx. fuscocephala		B	Pip, Fus	wsp	[83, 178]
Cx. gelidus		B	Con, Gel	wsp	[83, 178]
Cx. quinquefasciatus		A, B	Pip	wsp	[83, 169, 171]
Cx. sitiens		B	Pip, Con, Sit	wsp	[83, 178]
Cx. vishnui		A, B	Riv, Con	wsp	[83]
Cx. brevipalpis		A	Mors, Riv, Uni, Bre	wsp	[83, 178]
Cx. (Eumelanomyia) spp.		A	Eum	wsp	[178]
Cx (Lophoceraomyia) spp.		A	Lop	wsp	[178]
Cx. modestus		B	Pip	wsp	[178]
Cx. torrentium		B, C	Pip, Di	wsp	[178]
Conquillettidia	West Nile virus. Eastern equine encephalomyelitis virus, John Cunningham virus
Cq. richiardii		B	Con	wsp	[177, 181]
Cq. crassipes		A, B		wsp	[83]
Hodgesia spp.	-	A	Uni	wsp	[83]
Mansonia	Rift Valley fever virus			wsp
Mn. indiana		A, B	Riv, Con	wsp	[83]
Mn. uniformis		A, B	Riv, Con, Pip	wsp	[83, 171]
Mn. africana		B	–	wsp	[183]
Tripteroides aranoides	Sylvan yellow fever virus	B	Pip	wsp	[83]
Uranotaenia patriciae	Not known (isolated Eastern equine encephalitis virus, cyprovirus, Nounane virus)	A, B	Mors, Uni, Pip	wsp	[83]
Phlebotomus	Leishmania spp., Bartonella spp., phlebovirus, Toscana virus
P. papatasi		A	Turk 54, pap	wsp	[187]
P. (Larroussius) perfiliewi		B	AZ2331	wsp	[187]
Paraphlebotomus	Leishmania major			wsp
Pa. mongolensis		A	Turk 07, Turk 54	wsp	[187, 188]
Pa. caucasicus		A	Turk 07, Turk 54	wsp	[187, 188]
Lutzomyia	Leishmania spp.
Lu. c. cayennensis		B	Lev, Lcy	wsp	[189]
Lu. dubitans		B	Lev	wsp	[189]
Lu. evansi		B	Lev	wsp	[189]
Lu. cruciata		–	–	wsp, 16S rRNA	[190]
Lu. trapidoi		A	–	wsp	[191]
Lu. vespertilionis		A	–	wsp, FtsZ	[191]
Culicoides	Bluetongue virus, African horse sickness virus, Schmallenberg virus
C. pulicaris		A, B	–	wsp	[192]
C. imicola		A, B	–	wsp	[192]
C. kibunensis		B	–	wsp	[192]
C. vexans		B	–	wsp	[192]
C. obsoletus		B	–	wsp	[192]
C. narrabeenensis		B	–	16S rRNA	[193]
Simulium	Onchocerca spp.		–
Simulium squamosum		No identity with any of the supergroups	Dam	aspC, aspS, dnaA, fbpA, ftsZ, groEL, hcpA, IDA, rpoB, rpe, TopI wsp, FtsZ	[194, 195]
Glossina	Trypanosoma spp.
G. morsitans		A	Mors	wsp	[51]
G. austeni		A	Aus	wsp	[51]
Haematobia irritans irritans	Filarial nematode, Stephanofilaria stilesi, Staphylococcus spp.	A	wIrr	Whole genome sequencing	[163]
Tabanus sp.	Parasite transportation (Dermatobia hominis), biological transmission (Loa loa), and mechanical transmission of viruses, (equine infectious anemia virus), protozoa, (Trypanosoma evansi, Besnotia besnoiti) and bacteria (Bacillus anthracis, Anaplasma marginale)	–	–	FtsZ	[38]
Muscidae	Viruses (polioviruses, coxackie viruses), numerous bacteria (Campylobacter jejuni, Helicobacter pylori, Salmonella sp., Listeria sp., Yersinia pseudotuberculosis, Shigella, Vibrio), protozoan parasites (Giardia, Entameba) and eggs of several tapeworms
Musca sorbens		A	–	wsp	[196]
Musca domestica		B	–	wsp	[196]
Calliphoridae	Myiasis-producing agent, mechanical transmission of eggs of Taenia sp., Entamoeba coli, Giardia lamblia, Mycobacterium paratuberculosis
Chrysomya megacephala		A, B	–	wsp	[196]
Hemipyrellia pulchra		A	–	wsp	[196]
Sarcophagidae	Myiasis-producing agent
Sarcophaga dux		A	–	wsp	[196]
Sarcophaga scopariiformis		B	–	wsp	[196]
Fleas	Yersinia pestis, Rickettsia spp., Bartonella spp., Dipylidium caninum, Hymenolepis diminuta,
Ctenocephalides felis		I	–	16S rRNA	[32, 35, 197]
Tunga penetrans, Pulex irritans, P. simulans, Echidnophaga gallinacea, Ctenocephalides canis Xenopsylla brasiliensis Xenopsylla cheopis Xenopsylla nubica		Not specified	–	16S rRNA	[197–202]
Bugs	Trypanosoma cruzi, Bartonella quintana Burkholderia multivorans
Cimex lectularius		F	–	16S rRNA, FtsZ	[203]
Montina sp. (Reduvid bug)		-	–	FtsZ	[38]
Ticks
Rhipicephalus microplus	Babesia bovis, Babesia bigemina	A	–	16S rRNA
Ixodes ricinus	Borrelia burgdorferi, Anaplasma phagocytophilum, tick-borne encephalitis (TBE) virus	A			[204, 205]
Gustavia microcephala (Oribatid mite)	Anoplocephala spp. and Mesocestoidea spp.	E	–	16S rRNA, FtsZ, and gltA	[206]

Wolbachia in onchocercid nematodes

Wolbachia are obligatory endosymbionts required for the reproduction, development and long-term survival of onchocercid nematodes [95] and it has been hypothesized that they get from them, in return, essential aminoacids [17]. This endosymbiont is present in gradually increasing density from L1 to the adult filarioid [16, 96] of three subfamilies of Onchocercidae (i.e. Onchocercinae, Dirofilariinae and Splendidofilariinae) and in 16 of the 26 genera examined [12, 13, 49, 97] (Table 3). Indeed, most species in Litomosoides or Onchocerca genera have been found infected with Wolbachia but only one in the genus Cercopithifilaria (i.e. Cercopithifilaria japonica) [49, 97, 98]. However, there is a huge diversity in the localisation of these bacteria in tissues of different onchocercid nematodes and even between male and female individuals within the same species [97, 99]. In general, this bacterium is present in the female reproductive system and throughout embryonic development in the uterus of females (Fig. 1) being vertically transmitted to the progeny through the egg cytoplasm [49, 97, 100]. Other target tissues are the hypodemal lateral cords of the majority of onchocercid nematodes (e.g. Onchocerca, Brugia, Dirofilaria) and the intestinal cells of Mansonella (i.e. supergroup F) [12]. Nonetheless, the presence of Wolbachia may vary based on the species of onchocercid nematodes, being absent/less dense in lateral cords of Loxodontofilaria caprini [101] and Onchocerca dewittei japonica [97, 99]. Therefore, though the distribution of this endosymbiont is mostly concentrated in lateral cords and reproductive organs of the host, it exhibits different tropism to tissues during embryogenesis [12, 102]. In supergroups C and D there are similar patterns of embryonic segregation of Wolbachia with vertically transmitted bacteria reaching the lateral cords of the embryo by asymmetric mitotic segregation up to the ovaries [102, 103]. Though many investigations have focussed on the effects exerted by Wolbachia on the life performances of onchocercids, the role of this endosymbiont is still unclear [75, 102]. Five biosynthetic pathways (e.g. heme, riboflavin, FAD, glutathione and nucleotide synthesis) are present only in Wolbachia but not in any other rickettsiales or in onchocercid nematode hosts [17] as some involved genes (e.g. heme-biosynthesis genes) are absent in the onchocercid genome [64, 104]. In addition, the whole genome sequencing of Wolbachia from Brugia malayi (wbm) gave some clues regarding the role played by this bacterium in the filarial life cycle [17]. For example, the presence of heme metabolism and/or riboflavin genes in the wbm genome suggested a role of this bacterium in iron metabolism of the onchocercid nematodes [17, 104] though the transport, degradation and regulation of heme within filarial parasites remain a mystery for the scientific community. Therefore, the inhibition of nematode moulting following an antibiotic therapy targeting Wolbachia has been linked to the lack of production of ecdisone-like hormones because of the absence of heme, involved in the biochemical paths above [17, 105]. Similarly, heme inhibitors interfered with the vitality of onchocercid nematodes [106]. Though heme and nucleotide biosynthetic pathways are more conserved in all Wolbachia genomes, pathways like vitamin B are more variable in onchocercid nematodes (e.g. the hypothesis of Wolbachia providing vitamin B7 is clear in some insects such as bedbugs or grasshoppers but not demonstrated in filarial nematodes) [23]. In addition, the absence of Wolbachia and of any other biosynthesis pathways in the genome of Loa loa [107, 108] might suggest the presence of other alternative pathways for the essential nutritional requirement for this onchocercid species, a still open question that needs to be addressed by the scientific community along with the screening of new drug targets for filarial control.

Table 3

Onchocercid nematodes, their hosts and location, vectors and the main supergroup of Wolbachia detected

Onchocercids	Host	Location	Vector	Wolbachia supergroup	References
Onchocercinae					[102]
Acanthocheilonema dracunculoides	Canids	Coelomic cavity and subcutaneous tissue	Hippobosca Heterodoxus	NA	[102]
Acanthocheilonema reconditum	Canids	Subcutaneous tissue	Ctenocephalides Pulex Heterodoxus	Absent	[102]
Acanthocheilonema viteae	Rodents	Subcutaneous tissue	Ornithodoros	Absent	[102]
Acanthocheilonema odendhali	Northern fur seal	–	–	Absent	[13]
Brugia malayi	Humans	Lymphatic system, lymph nodes, testes	Mansonia Anopheles Aedes	D	[102]
Brugia pahangi	Dogs, felids	Lymphatic system, lymph nodes, testes	Anopheles Aedes	D	[102]
Brugia timori	Humans	Lymphatic system, lymph nodes, testes	Anopheles Aedes	D	[102]
Cercopithifilaria grassii	Dogs	Subcutaneous tissue	Rhipicephalus spp.	NA	[102]
Cercopithifilaria japonica	Ursidae (Black bear)	Oesophageal and tracheal connective tissue	–	F	[12]
Cercopithifilaria crassa	Sika deer	Dermis	Ixodid ticks	Absent	[12]
Cercopithifilaria longa	Sika deer	Subcutaneous connective tissues between muscles and skin of limbs and trunk	Ixodid ticks	Absent	[12]
Cercopithifilaria minuta	Japanese serow	Skin	Ixodid ticks	Absent	[12]
Cercopithifilaria multicauda	Japanese serow	Skin	Ixodid ticks	Absent	[12]
Cercopithifilaria roussilhoni	Brush–tailed porcupine	Skin	Ixodid ticks	Absent	[12]
Cercopithifilaria shohoi	Japanese serow	Skin	Ixodid ticks	Absent	[12]
Cercopithifilaria tumidicervicata	Japanese serow	Skin	Ixodid ticks	Absent	[12]
Litomosa westi	Rodents	Abdominal and pleural cavities	Ornithonyssus spp.	D	[49]
Litomosoides sigmodontis	Rodents	Coelomic cavity	Ornithonyssus	D	[102]
Litomosoides taylori	Water nectomys	Abdominal cavity	Suspected to be mites or bat flies	D	[12]
Litomosoides braziliensis	Bats	Abdominal cavity	Suspected to be mites or bat flies	D	[13, 49, 207]
Litomosoides solarii	Bats	Abdominal cavity	Suspected to be mites or bat flies	D	[13, 207]
Litomosoides hamletti	Bats	Abdominal cavity	Suspected to be mites or bat flies	D	[49, 207]
Litomosoides galizai	Murids	Coelomic cavity	Bdellonyssus bacoti	D	[49]
Litomosa chiropterorum				Absent	[12]
Litomosa yutajensis				Absent	[12]
Litomosoides chagasfilhoi	Mongolian gerbils	Abdominal cavity	Ornithonyssus bacoti	D	[208]
Loxodontofilaria caprini	Serows	Subcutaneous tissue mainly of limbs	Simulium japonicum T. japonensis (suspected vector)	C	[12, 209]
Montanema martini	Typical striped grass mouse	Skin	Ixodid ticks	Absent	[12]
Mansonella ozzardi	Humans	Coelomic cavity	Culicoides Simulium	F	[49, 210]
Mansonella perstans	Humans and monkeys	Coelomic cavity	Culicoides	F	[102, 210]
Mansonella streptocerca	Humans and monkeys	Intradermal	Culicoides grahamii	NA	[102, 210]
Mansonella perforata	Sika deer	Dermis	Culicoides spp.	F	[12, 210]
M. (T.) atelensis amazonae	Primates, Cebidae	Subscapular region	Culicoides spp.	F	[12, 210]
Onchocerca cervicalis	Equids	Nuchal ligament	Culicoides	C	[102]
Onchocerca gutturosa/ Onchocerca lienalis	Bovids	Nuchal ligament, connective tissue, gastro splenic ligament	Simulium arakawae Simulium daisense Simulium kyushuense Culicoides	C	[12, 209]
Onchocerca lupi	Canids	–	Unknown	C	[102]
Onchocerca ochengi	Bovids	Intradermal	Simulium	C	[102]
Onchocerca volvulus	Humans	Subcutaneous tissue	Simulium	C	[102]
Onchocerca armillata	Bovids occasionally camel	Thoracic aorta	Midges (Culicoides), Blackflies (Simulium)	C	[12]
Onchocerca borneensis n. sp.	Suids	Footpads of the hind limbs		C	[12]
Onchocerca dewittei japonica	Suids	Nodular fibrous structures in the footpads of fore- and hind limbs	Simulium bidentatum	C	[12, 207, 209]
Onchocerca caprini	Bovids	Skin	Simulium sp.	C	[12]
Onchocerca suzukii	Bovids	Subcutaneous tissue of the body, mainly in the thoracic area and pelvic limbs	Simulium japonicum Prosimulium sp. (Suspected vector)	C	[12, 209]
Onchocerca cervipedis	Cervids	Subcutaneous tissues of the legs	Prosimulium impostor Simulium decorum Simulium venustum	C	[211]
Onchocerca boehmi	Equids	Arteries and veins of the limbs	Not known	C	[212]
Onchocerca skrjabini	Cervids, bovids	Subcutaneous tissues of muzzle, hocks and to a lesser extend in brisket and shoulder	Simulium arakawae, Simulium bidentatum, Simulium oitanum (Putative vector)	C	[12, 209]
Onchocerca eberhardi	Cervids	Carpal ligament	Simulium arakawae, Simulium bidentatum, Simulium oitanum (Putative vector)	C	[12, 209]
Onchocerca gibsoni	Bovines	Subcutaneous and intermuscular nodules	Culicoides spp.	C	[34]
Onchocerca fasciata	Camels	Subcutaneous tissue and nuchal ligament	Unknown	–	[213]
Onchocerca jakutensis	Red deer, humans	Tissues of the outer thigh and the caudal part of the back; eye, neck and face nodules		–	[214]
Wuchereria bancrofti	Humans	Lymphatic system, lymph nodes, testes	Culex Anopheles Aedes	D	[102]
Dipetalonema gracile	Capuchin monkey	Abdominal cavity	Culicoides spp.	J	[13, 207, 215, 216]
Dipetalonema robini	New world monkey	Peritoneal cavity	Culicoides spp.	J	[13, 207, 216]
Dipetalonema caudispina	New world monkey	Peritoneal cavity	Culicoides spp.	J	[13, 207, 216]
Dipetalonema graciformis	New world monkey	Peritoneal cavity	Culicoides spp.	J	[216, 217]
Malayfilaria sofiani	Tree shrews	Tissues surrounding the lymph nodes of the neck	Not known	D	[207]
Cruorifilaria tuberocauda	Capybara	Kidney	Not known	J	[13]
Yatesia hydrochoerus	Capybara	Skeletal muscle	Amblyomma sp?	J	[13]
Breinlia (Breinlia) jittapalapongi	Tanezumi rat	Peritoneal cavity	–	Absent	[13]
Dirofìlariinae
Dirofilaria immitis	Canids, felids	Right ventricle pulmonary artery	Aedes	C	[102]
Dirofilaria repens	Canids, felids	Subcutaneous tissue	Culex Aedes	C	[102]
Loa loa	Humans	Subcutaneous tissue	Chrysops	Absent	[102]
Foleyella candezei	Rainbow agama	Subcutaneous tissue		Absent	[12]
Pelecitus fulicaeatrae	Black-necked grebe	Ankle region	Lice	Absent	[13]
Setariinae
Setaria equina	Horses	Coelomic cavity	Aedes	Absent	[102]
Setaria tundra	Roedeer	Peritoneal cavity	Aedes sp., Anopheles sp.	Absent	[13]
Setaria digitata	cattle	Peritoneal cavity	Aedes, Culex, Anopheles, Hyrcanus, Armigeres	Absent	[12]
Splendidofilariinae
Cardiofilaria pavlovskyi	Eurasian golden oriole	Body and pericardial cavity	–	NA	[13]
Madathamugadia hiepei	Turner's thick-toed gecko	–	Phlebotomus sp.?	F	[13]
Aproctella sp. 1	Rufous-bellied Thrush Green-winged saltator	–	–	Absent	[12]
Icosiellinae
Icosiella neglecta	Marsh frog, Edible frog	Muscle, subcutaneous tissue	–	Absent	[13]
Oswaldofilariinae
Oswaldofilaria petersi	Crocodilurus	Mesentery, intestine and thigh muscles	–	Absent	[13]
Piratuba scaffi	Lizard jungle runner	Skin		Absent	[12]
Waltonellinae
Ochoterenella phyllomedusa	Toads	–	Culicine mosquitoes	Absent	[13]
Ochoterenella royi	Cane toad	–	–	Absent	[12]
Ochoterenella sp. 1	Giant leaf frog	–	–	Absent	[12]

Fig. 1

Wolbachia in an embryo of the nematode Dirofilaria immitis (transmission electron microscopy observation). W: Wolbachia bacteria; n: nucleus; scale bar: 0.6 µm (Photograph of Luciano Sacchi and Claudio Bandi,

Modified from Bergey's Manual of Systematic Bacteriology, volume 2: The Proteobacteria) [221]

Wolbachia as a modulator of host inflammation and immunity

Filariasis has an extremely complex immunopathology with adult parasites surviving in immune competent patients for many years [49]. Many filarial nematodes harbour Wolbachia at all stages of their life cycle [49] except a few species such as L. loa, Acanthocheilonema viteae, O. flexuosa and Setaria equina [106, 109]. One of the main concerns in the treatment of filarial worms is related to the host inflammatory response provoked by the death of adult or larval worms within the parasitized tissues [77]. Under the above circumstances, the use of doxycycline targeting Wolbachia causes a soft killing of onchocercid nematodes with a slow death of adult parasites (over 12–24 months), preceded by a block of embryogenesis and larval development with clearing of microfilariae from blood or skin, thus preventing the pathology [110-112]. However, Wolbachia also plays a major role in the pathogenesis of both acute and chronic filariasis, which may cause severe systemic adverse reactions to chemotherapy as well as ocular inflammation by activating pro-inflammatory and immunomodulatory mechanisms in the host in cases of O. volvulus infection [113] (Fig. 2). Hence, Wolbachia has multiple roles in filariasis (i.e. activation of proinflammatory pathogenesis, immunomodulation of the host and survival of the parasite) starting from the entry of the parasite to the establishment of the infection [102].

Fig. 2

Role of Wolbachia in onchocecid nematode infections. Wolbachia induced changes in the host immune system such as immunomodulation for the survival of nematodes and various steps in the development of the immunopathology in filarial diseases are summarized

Wolbachia stimulates a proinflammatory immune response to onchocercid nematodes by interacting with the host monocytes, macrophages, dendritic cells and neutrophils [77]. Indeed, Wolbachia may elicit a host immune response by activating the innate and adaptive responses in human and murine models of filariasis infection [102]. Generally, in bacterial infections, the host innate immune system recognises pathogen-associated molecular patterns (PAMP) associated with bacteria which are presented by pattern recognition receptors (PRP) including the Toll-like receptors (TLR) located on the surface of antigen-presenting cells [77] (Fig. 2). Though lipopolysaccharides (LPS) act as a major PAMP in most bacterial infections, the absence of genes required for LPS biosynthesis in the Wolbachia genomes suggests that the Wolbachia surface protein (WSP) is a predominant PAMP involved in the immunopathology of filarial diseases [106, 114, 115]. Recently, further evidence on the immune-modulating property of WSP showed that the Asaia endosymbiotic bacterium engineered for the expression of this protein induces classical macrophage activation, associated with the killing of Leishmania parasites [116].

In addition to WSP, other PAMPs of Wolbachia include HSP 60 and groEL, and all these molecules mainly elicit TLR 2 or TLR 6 recognition and signalling [77]. In particular, WSP produces inflammation and regulatory marker expression (i.e. T lymphocyte antigen 4) while HSP 60 acts on monocytes and produces cytokines and TNFα, which induce an adaptive immune response against onchocercid nematodes [102, 113, 117]. Hence, in O. volvulus pathology, TLR 2 activates macrophages and local stromal cells contributing to the neutrophil-mediated corneal inflammation and permanent destruction of the cornea [118, 119]. In chronically ill patients, repeated invasion of larvae and their death produce inflammatory cell influx, eventually causing permanent tissue damage by neutrophil degranulation and the production of cytotoxic products such as nitric oxide (NO), myeloperoxidase and oxygen radicals [119]. It is believed that, in bancroftian patients, Wolbachia with its TLR2 signalling triggers vascular endothelial growth factor A and angiopoietin-1 production, which produces the dilatation of scrotal lymph vessels [120] (Fig. 2). This is supported by the results obtained after the administration of doxycycline [121]. Absence of such immunological response by the extracts of the worm A. viteae, which have no Wolbachia endosymbiont, supports the above findings [113, 117].

In addition to the contribution in immunopathology, Wolbachia plays a defensive mutualistic role in filarial biology (e.g. O. ochengi) by triggering the recruitment of more neutrophils, which will help to prevent the degranulation of eosinophils needed for the eradication of filarial worms [122]. A similar reaction is also observed in human D. immitis infection wherein WSP prevent the neutrophil apoptosis [123]. Moreover, further in vitro studies on blood cells from patients with O. volvulus suggested that chronic Wolbachia stimulation may cause the downregulation of pro-inflammatory mediators by increasing the production of interleukin 10 (IL-10) and thus help in establishing chronic infection [124]. In addition, it is suggested that together with filarial antigens, Wolbachia may induce generation of NO, which aids in the peripheral tolerance through apoptosis of antigen-specific T lymphocytes [125]. This manipulation of the host immune system helps to increase the longevity of onchocercid nematodes [126, 127] (Fig. 2). Moreover, a stronger immune reaction in response to the release of L3 larvae than to the dead worm indicates the predominant role of these bacteria at the development of the early stage of the worm [77]. Wolbachia with its TLR 2-dependent signalling helps the filarial L3 larva establishment by surpassing chemokine (c–c motif) ligand 17 (CCL 17)-mediated immune response of the host [128].

Exploiting Wolbachia for treatment

Current filariasis treatment control and Mass Drug Administration (MDA) programmes are focussed on the use of microfilaricides such as albendazole combined with either ivermectin or diethylcarbamazine [129]. Though this treatment regime helped to reduce the number of human cases of filariases, the long duration of treatment (e.g. 17 years for onchocerciasis and 5 years for lymphatic filariasis), the development of resistance to ivermectin in endemic areas as well as adverse reactions to ivermectin treatment in certain epidemiological conditions (e.g. onchocerciasis-loasis co-endemic areas) hampered the global elimination of the diseases [110, 130]. Hence, chronic debilitating pathological alterations and the economic burdens in endemic countries due to long-term treatment and control programmes highlighted the need for an alternative effective short-term potential drug target for filariasis.

Based on the unique obligatory symbiotic relationship Wolbachia has established with these onchocercid nematodes (e.g. embryogenesis and moulting) and the role of these bacteria in the immunopathology of filarial diseases, a major mission of the anti-Wolbachia (A∙WOL) consortium was to exploit the Wolbachia-filarial biology for controlling human infection [131]. Studies suggested that anti-Wolbachia therapy has both macrofilaricidal (i.e. death of adult parasites and developmental retardation) and microfilaricidal embryotoxic activity. Indeed, antibacterial agents such as doxycycline were found to be effective in clearing microfilarial stages from the blood and skin of patients, therefore preventing filarial pathology and reducing the transmission [131]. Moreover, the slow death of adult parasites over a period of 12–24 months in patients treated with doxycycline is safe to use in geographical regions were onchocerciasis and loiasis occurred in sympatry [110-112]. However, the mechanism of action of doxycycline in the treatment of filariases was not well understood until transcriptomic and proteomic analysis unveiled that the responses of Wolbachia to doxycycline cause impairment of bacterial metabolism [132]. Meta-analytical modelling suggests that a 4-week doxycycline course suffices to eliminate Wolbachia with low chances of developing drug resistance when compared to other antibiotics like penicillin or fluoroquinolones [133, 134]. However, the limited use of this drug in pregnancy, lactating mothers and children motivated the scientific community to search for other anti-Wolbachia drugs [127, 131]. As a result, more than 2 million compounds have been tested in insect cell lines and A∙WOL was formed with the objective to identify new anti-Wolbachia drugs with a short course of therapy, which could be safe in contraindicated groups [135, 136]. Mass screening of all registered antibacterials revealed four drugs as superior to doxycycline with minocycline as the most effective drug of choice [137]. Recent experimental trials claim that the use of Tylosin A, a macrofilaricide, is superior to tetracycline antibiotics (e.g. doxycycline and minocycline) and will help to reduce the duration of treatment from 3–4 to 1–2 weeks [131].

Targeting Wolbachia for vector control

Vector control methods mainly focus on the physical removal of their breeding sites in the environment or on- and off-host application of insecticides targeting immature or adult stages [138]. A combination of factors such as human population growth, globalization, rapid rise in population-dense towns, expansion of the geographical range of vectors and development of insecticide resistance affected the control of vectors and associated pathogens [139, 140]. Hence, the search for an alternative vector control approach may target either reducing the vector population or modifying the vector to make it refractory to pathogen transmission [138]. For example, the abundance of Wolbachia among vectors and its high rate of maternal transmission conjoined with CI have spurred the interest of researchers in new target strategies for vector control. It was discovered that Wolbachia can protect its natural host Drosophila melanogaster from pathogenic viruses, such as Drosophila C virus [138, 141]. Since then, many Wolbachia strains have been found to block the transmission of a range of medically important viruses and parasites [138, 142]. Thus, it was proposed that the use of large numbers of Wolbachia-infected males to sterilize local uninfected females through CI (incompatible insect technique, IIT) coupled with a pathogen-blocking Wolbachia strain could be effectively gradually replace the local permissive natural vectors with refractory insects [143]. Therefore, a non-profit research consortium, namely the “World Mosquito Program” (WMP), formerly known as the “Eliminate Dengue Program”, was instituted to eliminate mosquito-borne viral diseases like dengue, Zika and chikungunya [141, 144]. In this context, it has been shown that the intensity of Wolbachia infection is directly correlated with the strength of pathogen blocking and the tissue damage caused by the cellular load of highly replicative strains of Wolbachia (e.g. wMelPop) produces pathogen blocking in Ae. aegypti [145]. Other possible mechanisms are host immune priming by the preactivation of the immune response and gene regulation by the induction of Vago1 protein, which is involved in the innate immune pathways of Culex quinquefasciatus and Ae. aegypti. This mechanism could favour vectors to reduce West Nile and dengue virus replication [146, 147]. Though vectors like Ae. aegypti are not naturally infected with the virus inhibiting Wolbachia, these non-native strains of Wolbachia were introduced into the vectors of medical and veterinary importance by transfection [148] (Fig. 3). Other methods such as transient somatic infection, infections in cell lines, ex vivo organ culture, outcrossing and introgression can also be used for the successful introduction of Wolbachia in non-native vectors [147] (Table 4). Current research on control mainly focuses on limiting the susceptibility to infection rather than using this symbiont to reduce the life span of vectors [149]. Indeed, the fitness cost of wMelPop strain prompted the researchers to adopt another strain of Wolbachia, wMel which does not reduce the fitness of mosquito hosts [149, 150]. Approximately 300,000 wMel-infected Ae. aegypti mosquitoes were released in north Queensland over a period of 10 weeks [149]. The success of this strategy in north Queensland (i.e. a high infection frequency up to 80%–90%) was replicated in 12 countries, including Brazil, Indonesia, Vietnam, and four countries of the south-western Pacific region [141]. These studies have shown that the wMel strain of Wolbachia can quickly spread to near fixation in the wild mosquito population and become stable for a long time after the initial release [138]. Apart from these, in West Africa, stable Wolbachia infections were recently detected in natural Anopheles populations and these infections appear to be negatively correlated to Plasmodium prevalence, which opens up the possibility of utilising these endosymbionts for the control of malaria transmission [143]. The introduction of Wolbachia infections in Culicoides sonorensis cell lines and the upregulation of immune genes in the same vectors suggested the utility of using Wolbachia as a bio-control agent in the transmission of C. sonorensis vectored pathogens of veterinary importance (e.g. African horse sickness virus, Schmallenberg virus, bluetongue virus, epizootic hemorrhagic disease virus) [151]. Like any other modern technologies, Wolbachia-based vector control also has some potential vulnerabilities such as (i) loss of attenuation of Wolbachia infection in the mosquito, (ii) emergence of virus strains that are resistant to Wolbachia-mediated blocking, (iii) increasing virulence and disease pathogenesis in humans, (iv) enhancement of the arbovirus infection in transfected mosquitoes and (v) development of mutations in viruses over time that render them less susceptible or resistant to Wolbachia [141]. Apart from these the current method of control requires continual release of large numbers of males to suppress the mosquito population and the migration of mosquitoes from the untreated surroundings will hinder the long-term effectiveness of this method. Since only modified males are released into the environment, adoption of an effective sex sorting system is required [138]. Nonetheless, none of these modifying technologies has yet been approved by the WHO’s Vector Control Advisory Group [138]. In addition, it is highly advisable to avoid adverse effects such as the enhancement of pathogen development in coinfections (Table 4) by analysing the molecular mechanisms of Wolbachia-pathogen interactions before doing the field trials.

Fig. 3

Exploiting Wolbachia for vector control. Left panel: mosquito population replacement approach, in which Wolbachia-infected female and male mosquitoes are released; through CI, this strategy allows the spread of Wolbachia in the natural population. The presence of Wolbachia provides a fitness advantages (determined by CI) and can reduce the arbovirus transmission. Right panel: mosquito population reduction or suppression strategy. This approach involves the release of Wolbachia-infected males into an area; when these mosquitoes mate with wild Wolbachia-negative females (or female mosquitoes harbouring an incompatible strain of Wolbachia), a strong reduction in the rate of egg hatching is observed (CI incompatible matings). Thus, repeated releases of Wolbachia-infected males result in reduction or suppression of mosquito populations. CI cytoplasmic incompatibility, WT wild-type mosquitoes, Wolb + Wolbachia-infected mosquitoes

Table 4

Progress in Wolbachia-based vector modifications for control and its possible outcomes

Vector	Pathogens transmitted	Stable transfections	Strains used	Protected against pathogens	Effects/favourable outcome of transfected mosquitoes	Reversal outcome effects in transfected mosquitoes	Reference
Aedes	Yellow fever virus Dengue (D1, D2, D3, D4) viruses Chikungunya virus Zika virus (ZIKV) Rift Valley fever virus Wuchereria bancrofti, Brugia malayi	Aedes aegypti Aedes albopictus Aedes polynesiensis	wAlbB, wMel, wMelPopCLA	Yellow fever virus Chikungunya Plasmodium gallinaceum ZIKV and ZIKV/DENV coinfection transmission blockage Brugia pahangi	wAlB-mosquitoes successfully established and reduced human dengue incidence was registred wMel-mosquitoes successfully established across 66 km2 and no local dengue transmission was registered	Enhance DENV D2 in Aedes aegypti and Plasmodium gallinaceum in Aedes fluviatilis	[144, 147, 218]
Anopheles	Plasmodium falciparum, Plasmodium vivax, Plasmodium malariae, Plasmodium ovale, Plasmodium knowlesi Wuchereria bancrofti, Brugia malayi	Anopheles gambiae Anopheles stephensi	wAlbB	Plasmodium falciparum, modestly suppress Plasmodium berghei oocyst levels, Plasmodium yoelii at some temperatures	Modest decrease in oocyst numbers and a strong reduction in salivary gland sporozoites of Plasmodium falciparum	Enhance Plasmodium yoelii at some temperatures in Anopheles stephensi	[147]
Culex	West Nile virus Equine encephalitis virus Japanese encephalitis virus Saint Louis encephalitis virus Rift Valley fever virus Wuchereria bancrofti, Brugia malayi	Culex tarsalis	wPip	West Nile virus (WNV)	Increasing Plasmodium relictum transmission stages	Enhance WNV infection in Culex tarsalis and Plasmodium relictum in Culex quinquefasciatus	[147, 219, 220]

Conclusions and future perspectives

Though Wolbachia is a relatively well-studied endosymbiont [4], there are still lacunae in the knowledge about its exact distribution, evolution, type of symbiosis and Wolbachia-mediated antiparasitic mechanisms. It has been hypothesised that onchocercid nematodes may depend on Wolbachia for their heme metabolism [17]. For example, no new biosynthesis pathways for heme metabolism are observed in L. loa, which lacks Wolbachia [17], therefore suggesting alternative pathways or the presence of other symbionts for the essential nutritional requirement of these worms. While Wolbachia is already well studied, many other endosymbionts have received less attention, such as some Spiroplasma, Cardinium, Arsenophonus and Flavobacetrium species [152] and have not yet been investigated in detail. So, the metagenomics approach could help to assess the associated endosymbionts in Wolbachia free onchocercid nematodes. In addition, the use fluorescence in situ hybridization is advisable to assess the natural infections by Wolbachia [153]. Since Wolbachia dominant proteins expressed in each life stage of onchocercid nematodes show a gradual increase from L1 to adult [16, 96], proteomic approaches (e.g. mass spectrometry, chromatography) could be useful to assess their variation in expression in each stage in the vector and the definitive host for their survival and multiplication. This may eventually lead to exploring the type of symbiosis at each stage of the parasite life cycle, also providing insights into the Wolbachia-mediated antiparasite mechanisms and potential new drug targets for onchocercid nematodes of medical and veterinary significance. Despite doxycycline being adopted to treat filarial diseases, potential difficulties (e.g. drug adherence, toxicity, resistance, financial cost, contraindications in pregnant women and children) limit its use in the public health MDA programmes [127]. Promising in vitro drug trials with new antibiotics (e.g. berberine, rapamycin, globomycin, succinyl acetone) [106, 154–156] and the effectiveness of non-antimicrobial compounds such as anti-oxidants and anti-histamines open a new window onto filarial treatment [127]. Further clinical trials using these drugs may provide an innovative strategy for anti-Wolbachia treatment, eventually reducing the duration of treatment.

Overall, the prevalence of Wolbachia differs significantly among different climatic regions and geographic locations [157]. For example, the intensity of Wolbachia infections in natural Ae. albopictus populations was low in regions with only imported dengue cases suggesting a positive correlation with the presence of Wolbachia in vectors and dengue infection [157]. Based on the above, it would be necessary to have a cluster-randomized design, involving either long- or short-term vector-release trials in limited locations or in more sites, respectively, to optimize the impact of this control strategy in each geographical or climatic setting [158]. Nonetheless, Wolbachia-based vector control strategies to control arboviral infections targeting Aedes aegypti are being compromised in many endemic countries because of the co-localisation of the secondary vector Aedes albopictus [159]. Hence, it is important to rely also on fluorescence in situ hybridisation when reporting natural Wolbachia infections and not only on PCR. Furthermore, use of multiple strains of Wolbachia for vector transfection and integration of CI-carrying phage elements into strains that are devoid of them (e.g. wAu) [159] could provide more fitness benefits for the transfected vector. Under the above circumstances, transfected triple-strain infection of Wolbachia (e.g. wMel and wPip and wAu) into a Malaysian Ae. albopictus line produced self-compatibility, moderate fitness cost and complete resistance to Zika and dengue infections [160]. Apart from these, more field trials using wAlbB strains will also help to overcome the inability of wMelPop strains to establish in wild mosquito populations or the wMel strains to survive at high temperatures in the field [161]. This could be considered a successful strategy to reduce the incidence of dengue in an endemic area of Malaysia after the release of wAlbB-infected Ae. aegypti [161]. Similarly, use of other endosymbionts along with Wolbachia will help to accelerate the control of D. immitis through the use of genetically engineered Asaia bacteria for the expression of WSP from their Wolbachia endosymbionts [80]. However, the untoward effects of Wolbachia such as irreversible biological effects and reversal outcome on disease transmission [162] should be properly addressed before clinical trials. Successful introduction of Wolbachia infections in Anopheles gambiae, Anopheles stephensi and C. sonorensis cell lines may give a breakthrough in the control of malaria, African horse sickness, Schmallenberg, bluetongue and epizootic hemorrhagic disease. The genome sequencing of the Wolbachia strain, wIrr, of Haematobia irritans irritans suggests its unique features, including the horizontal acquisition of additional transcriptionally active CI loci, which may be exploited for the biocontrol and potential insecticide resistance of horn flies [163]. Despite all the challenges, studies on Wolbachia and their use in the control and/or treatment of vectors, onchocercid nematodes and viral diseases of medical and veterinary importance offer new approaches which undoubtedly open new avenues for the control of a variety of vector-borne diseases.