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1 Wolbachia pipientis associated to tephritid fruit fly pests: from basic research to
2 applications
3 Mariana Mateos, Humberto Martinez, Silvia B. Lanzavecchia, Claudia Conte, Karina
4 Guillén, Brenda M. Morán-Aceves, Jorge Toledo, Pablo Liedo, Elias D. Asimakis,
5 Vangelis Doudoumis, Georgios A. Kyritsis, Nikos T. Papadopoulos, Antonios A.
6 Avgoustinos, Diego F. Segura, George Tsiamis, and Kostas Bourtzis
7
8 Affiliations
9 (MM) Department of Wildlife and Fisheries Sciences, Texas A&M University, College
10 Station, Texas, USA; corresponding author [email protected]
11 (PL JT BMM KG) El Colegio de la Frontera Sur, Tapachula, Chiapas 30700, Mexico.
12 (HM) Departamento de Tecnología de Alimentos, Unidad Académica Multidisciplinaria
13 Reynosa Aztlan, Universidad Autónoma de Tamaulipas, Mexico
14 (SBL DFS CC) Laboratorio de Genética de Insectos de Importancia Económica, Instituto de
15 Genética ‘Ewald A. Favret’, CICVyA, Instituto Nacional de Tecnología Agropecuaria
16 (INTA), Hurlingham, Buenos Aires, Argentina
17 (KB AAA) Insect Pest Control Laboratory, Joint FAO/IAEA Division of Nuclear Techniques in
18 Food and Agriculture, Vienna, Austria
19 (GT EDA VD) Department of Environmental and Natural Resources Management, University
20 of Patras, 2 Seferi St, 30100 Agrinio, Greece
21 (NTP, GAK) Laboratory of Entomology and Agricultural Zoology, Department of Agriculture
22 Crop Production and Rural Environment, University of Thessaly, Phytokou St., 38446 N.
23 Ionia Magnisia, Greece
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24 Abstract
25 Members of the true fruit flies (family Tephritidae) are among the most serious
26 agricultural pests worldwide, whose control and management demands large and costly
27 international efforts. The need for cost-effective and environmentally-friendly integrated
28 pest management (IPM) has led to the development and implementation of autocidal
29 control strategies. Autocidal approaches include the widely used sterile insect technique
30 (SIT) and the incompatible insect technique (IIT). IIT relies on maternally transmitted
31 bacteria (namely Wolbachia), to cause a conditional sterility in crosses between released
32 mass-reared Wolbachia-infected males and wild females, which are either uninfected or
33 infected with a different Wolbachia strain (i.e., cytoplasmic incompatibility; CI). Herein,
34 we review the current state of knowledge on Wolbachia-tephritid interactions including
35 infection prevalence in wild populations, phenotypic consequences, and their impact on
36 life history traits. Numerous pest tephritid species are reported to harbor Wolbachia
37 infections, with a subset exhibiting high prevalence. The phenotypic effects of
38 Wolbachia have been assessed in very few tephritid species, due in part to the difficulty
39 of manipulating Wolbachia infection (removal or transinfection). Based on recent
40 methodological advances (high-throughput DNA sequencing) and a breakthrough
41 concerning the mechanistic basis of CI, we suggest research avenues that could
42 accelerate generation of necessary knowledge for the potential use of Wolbachia-based
43 IIT in area-wide integrated pest management (AW-IPM) strategies for the population
44 control of tephritid pests.
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45 1 Background
46 1.1 The economic importance of tephritids as pests
47 Flies in the family Tephritidae (Diptera) include some of the world’s most important
48 agricultural pests. The family is comprised of ~4900 described species within 481
49 genera, of which seven (Anastrepha, Bactrocera, Ceratitis, Dacus, Rhagoletis,
50 Toxotrypana and Zeugodacus) contain ~70 major pest species [1-5]. In addition to
51 causing billions of dollars in direct losses to a wide variety of fruit, vegetable and flower
52 crops, pest tephritids limit the development of agriculture in many countries due to strict
53 quarantines implemented in fruit-buying countries, and to the costs associated with
54 efforts aimed at prevention, containment, suppression, and eradication.
55
56 To prevent or minimize the harmful effects of tephritid pests, growers of affected crops
57 must comply with health and safety standards required by the market, applying an area-
58 wide management approach involving chemical, biological, cultural, and autocidal
59 control practices [6, 7]. Autocidal refers to methods that use the insect to control itself,
60 by releasing insects that are sterile or induce sterility upon mating with wild insects in the
61 next or subsequent generations [8-10]. Autocidal strategies include the sterile insect
62 technique (SIT); one of the most widespread control methods used against fruit flies.
63 SIT relies on the mass-rearing production, sterilization and recurrent release of insects
64 (preferentially males) of the targeted species. Sterilization is attained by radiation, in a
65 way that does not impair male mating and insemination capabilities. Wild females that
66 mate with sterilized males lay unfertilized eggs. At the appropriate sterile:wild (S:W)
67 ratio, the reproductive potential of the target population can be reduced [11-13].
68 Historically, at least 28 countries have used the SIT at a large-scale for the suppression
69 or eradication of pest insects [14-16]. SIT has been applied successfully for several
70 non-tephritid insect pests, for example: the New World screw worm (Cochliomyia
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71 hominivorax Coquerel); several species of tsetse fly (Glossina spp.); and the codling
72 moth (Cydia pomonella L.) [reviewed in 17, 18].
73
74 Successful SIT programs as part of Area-wide Integrated Pest Management (IPM)
75 strategies have also been implemented for several tephritids: Ceratitis capitata
76 Wiedemann; Anastrepha ludens Loew; Anastrepha obliqua Macquart; Anastrepha
77 fraterculus Wiedemann; Zeugodacus cucurbitae Coquillett; Bactrocera dorsalis Hendel;
78 and Bactrocera tryoni Froggatt [6, 12, 13, 15]. SIT is currently being developed for two
79 additional tephritid species: Dacus ciliatus Loew and Bactrocera tau Walker [19, 20].
80 The advantages of the SIT over other pest control approaches (e.g. use of pesticides)
81 are that it is the most environmentally friendly and resistance is unlikely to evolve [21,
82 22].
83
84 Another autocidal strategy where mating between mass-reared and wild insects can be
85 used to suppress pest populations is the incompatible insect technique (IIT). IIT also
86 relies on the principle of reducing female fertility, but utilizes endosymbiotic bacteria
87 instead of radiation, to induce a context-dependent sterility in wild females. It is based
88 on the ability of certain maternally inherited bacteria (namely from the genus Wolbachia)
89 to induce a form of reproductive incompatibility known as cytoplasmic incompatibility (CI;
90 explained in the section below). Herein we review the current knowledge on taxonomic
91 diversity of Wolbachia-tephritid associations and their phenotypic consequences, and
92 identify gaps in knowledge and approaches in the context of potential application of IIT in
93 AW-IPM programs to control tephritid pests. We also discuss scenarios where these
94 two autocidal strategies, SIT and IIT, could be potentially combined for the population
95 suppression of tephritid pests.
96
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97 1.2 The influence of Wolbachia on host ecology
98 Insects and other arthropods are common hosts of maternally inherited bacteria
99 [reviewed in 23]. These heritable endosymbionts can have a strong influence on host
100 ecology. Such vertically transmitted bacteria are typically vastly (or fully) dependent on
101 the host for survival and transmission. Certain associations are obligate for both
102 partners, and generally involve a nutritional benefit to the host. Other heritable bacteria
103 are facultative, with such associations ranging from mutualistic to parasitic from the
104 host’s perspective. Among these, Wolbachia is the most common and widespread
105 facultative symbiont of insects and arthropods [24-27].
106
107 Wolbachia is a diverse and old genus [possibly older than 200 million years; 28] of
108 intracellular gram-negative Alphaproteobacteria (within the order Rickettsiales)
109 associated with arthropods and filarial nematodes. Wolbachia cells resemble small
110 spheres 0.2–1.5 μm, occur in all tissue types, but tend to be more prevalent in ovaries
111 and testicles of infected hosts, and are closely associated with the female germline
112 [reviewed by 29, see also 30]. Wolbachia is estimated to infect 40–66% of insect
113 species [24-27]. Within a species or population, the infection prevalence of Wolbachia
114 can be quite variable over space [e.g. 31] and time [e.g. 32, 33].
115
116 The most commonly documented effects of Wolbachia on arthropod hosts fall under the
117 category of reproductive parasitism, which involves manipulation of host reproduction to
118 enhance symbiont transmission and persistence, in general by increasing the relative
119 frequency of Wolbachia-infected vs. uninfected females. Females are typically the sex
120 that can transmit Wolbachia and other heritable bacteria, although rare exceptions exist
121 [34-36]. Wolbachia employs all four types of reproductive manipulation [reviewed by 37,
122 38, 39]. Feminization results in genetic males that develop and function as females,
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123 and occurs in the orders Hemiptera, Lepidoptera, and Isopoda. Wolbachia-induced
124 parthenogenesis occurs in haplo-diploid hosts (e.g. Acari, Hymenoptera and
125 Thysanoptera), where unfertilized eggs, which would otherwise develop into males,
126 develop into females. Male killing eliminates infected males to the advantage of
127 surviving infected female siblings, and occurs in Coleoptera, Diptera, Lepidoptera, and
128 Pseudoscorpiones. Cytoplasmic incompatibility (CI) [40] prevents infected males
129 from producing viable offspring upon mating with females lacking Wolbachia (or a
130 compatible strain of Wolbachia; see below). CI is the most commonly reported
131 Wolbachia-induced reproductive phenotype, and is found in Acari, Coleoptera, Diptera,
132 Hemiptera, Hymenoptera, Isopoda, Lepidoptera, and Orthoptera. In unidirectional CI,
133 eggs from uninfected females that are fertilized by sperm from Wolbachia-infected males
134 fail to develop (Fig. 1a). Bi-directional CI results from crosses involving two different
135 (incompatible) Wolbachia strains (Fig. 1b). In contrast, crosses between females and
136 males infected with the same or compatible Wolbachia strains are viable. Similarly,
137 Wolbachia-infected females are compatible with uninfected males [39]. Consequently,
138 above a certain threshold of Wolbachia infection frequency in a host population, infected
139 females are at a reproductive advantage over uninfected females, resulting in a positive
140 feedback loop that leads to the increase of Wolbachia frequencies [41].
141
142 CI was discovered almost half a century ago [40], but its mechanism has not been fully
143 elucidated. A useful conceptual model to understand the observed patterns of CI is
144 “mod/resc” [42, 43]. It postulates that Wolbachia has two functions: mod (for
145 modification), which acts as a toxin or imprint of the male germline; and resc (for
146 rescue), which acts as an antidote. The mod function acts on the nucleus in the male
147 germline, before Wolbachia are shed from maturing sperm [44]. When a sperm nucleus
148 affected by mod enters the egg of an uninfected female, this nucleus encounters
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149 problems such as delays in DNA replication and cell-cycle progression, leading to
150 embryo death. In contrast, if the appropriate resc (“the antidote”) function is active in the
151 egg, the defect caused by mod in the sperm is rescued, and the embryo proceeds
152 through normal development. Based on the mod/resc model, four different CI-
153 Wolbachia types (strains) are possible: (1) the mod+/resc+ type, capable of inducing CI
154 and rescuing its own modification; (2) the mod–/resc– type, incapable of inducing CI or
155 rescuing the CI effect of other mod+ strains; (3) the mod+/resc– (“suicidal”) type, capable
156 of inducing CI but not rescuing its own modification; and (4) the mod–/resc+ type,
157 incapable of inducing CI, but able to rescue the CI defect caused by a mod+ strain [45,
158 46].
159
160 Three recent breakthrough studies identified a Wolbachia-encoded operon (of viral
161 origin) that appears to be required to induce CI [47-49]. LePage et al. [47] used a
162 combination of genomic, bioinformatic and molecular approaches to identify two
163 contiguous genes (cifA and cifB) encoded in the genome of Wolbachia wMel strain (from
164 D. melanogaster). LePage et al. [47] found that homologous genes are present in CI-
165 inducing strains, but are absent, or highly diverged, in non-CI-inducing strains. In
166 addition, LePage et al. [47] found that strain wRi (from D. simulans), which is compatible
167 with strain wMel, but not vice versa (see Fig. 1c) [50, 51], shares two sets of similar CI
168 operons with wMel, but has an additional divergent CI operon, which could explain why
169 wMel is unable to rescue the wRi-induced defects. LePage et al. [47] then expressed
170 cifA and cifB as transgenes in the germ line of D. melanogaster males. When these
171 males were mated to Wolbachia-free females, the offspring had reduced hatch rates and
172 embryonic cell-division defects that resembled those observed in CI. In contrast, when
173 these males were mated to Wolbachia-infected females, hatching rate was restored. In
174 a parallel study, Beckmann et al. [48] identified what appear to be two homologous CI
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175 operons in the strain wPip (from Culex pipiens L.) named cidA-cidB and cinA-cinB. They
176 also expressed them in D. melanogaster male germline, and found that when these
177 males were mated to Wolbachia-free females, the offspring exhibited the characteristic
178 embryonic abnormalities of CI. Based on the observation that the expression of cidB or
179 cinB in yeast blocks growth, whereas expression of cidA or cinA, respectively rescue the
180 defect, Beckmann et al. [48] proposed that cidA and cinA function as the antidote to cidB
181 or cinB, respectively. A similar rescue phenomenon was not recapitulated in the fly,
182 though. More recently, Shropshire et al. [49] demonstrated that cifA in D. melanogaster
183 acts as the rescue gene, and thus propose a “Two-by-One” model whereby cifA and cifB
184 induce CI and cifA rescues CI.
185
186 In addition to its reproductive phenotypes on arthropods, Wolbachia engages in obligate
187 mutualistic interactions with filarial nematodes [39] and with members of five insect
188 orders [reviewed in 52]. For example, it serves as a nutritional mutualist of bedbugs
189 [53], and as an enabler of oogenesis in a parasitic wasp [54]. As a facultative symbiont,
190 Wolbachia can provide direct fitness benefits to its insect hosts by influencing
191 development, nutrition, iron metabolism, lifespan, and fecundity [55-60], and most
192 notably, by conferring resistance or tolerance to pathogens (particularly single-stranded
193 RNA viruses) and parasites [61-73]. The interference of Wolbachia with the replication
194 and transmission of certain viruses forms the basis of several regional mosquito-borne
195 disease control programs [74; e.g. http://www.eliminatedengue.com, 75, 76].
196
197 Certain host-Wolbachia combinations incur fitness costs to the host, beyond
198 reproductive parasitism, including reduced longevity, sperm competitive ability, and
199 fecundity, as well as higher susceptibility to natural enemies [77-84]. Similarly, certain
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200 host-Wolbachia combinations may potentially enhance pathogen-vectoring capacities
201 [85-88].
202 2 Methods to study Wolbachia
203 2.1 Methods to assess Wolbachia infection status
204 For purposes of this review, we consider a host species or population as “infected” with
205 Wolbachia, even if the infection is transient or found at low titer. Wolbachia, and most
206 cytoplasmically transmitted endosymbionts, are fastidious to culture outside host cells,
207 such that their study typically relies on culture-independent methods. A recommended
208 flow-chart of steps is depicted in Fig. 2. The most utilized approach to date for
209 identifying hosts infected with Wolbachia is through PCR screening of Wolbachia genes
210 in DNA extracts of hosts. Different PCR primers have been used to perform such
211 surveys, traditionally targeting a portion of the 16S ribosomal (r)RNA gene or of a
212 ubiquitous protein-coding gene (e.g. wsp or ftsZ). Simoes et al. [89] evaluated the
213 relative sensitivity and specificity of different primer pairs aimed at Wolbachia detection
214 and identification, revealing that no single PCR protocol is capable of specific detection
215 of all known Wolbachia strains. A related method known as “loop mediated isothermal
216 amplification” (LAMP; not shown in Fig. 2), which requires less infrastructure than PCR,
217 has been successfully employed for Wolbachia detection in several insects [90].
218
219 The two major shortcomings of utilizing solely PCR (or LAMP) to detect Wolbachia
220 presence can be classified into the occurrence of false negatives and false positives. A
221 false negative occurs when a specimen is infected by Wolbachia, yet the screening
222 approach fails to detect its presence. The efficiency of the PCR can be affected by the
223 presence of inhibitors [91, 92], by low concentration/poor quality of the target DNA
224 molecule, as well as type and concentration of the polymerase and other PCR reagents.
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225 At the very least, negative Wolbachia detection PCRs should be validated by evaluating
226 the quality of the DNA extract, through positive amplification of a host-encoded gene
227 (e.g. the mitochondrial Cytochrome Oxidase subunit I or single-copy nuclear genes).
228 Several higher sensitivity approaches have been devised, particularly for low-titer
229 infections, such as: long PCR [93]; nested PCR [94]; quantitative PCR [95]; or the design
230 of alternative and/or more specific primers, including the use of Wolbachia multi-copy
231 genes as PCR targets [96]. These methods, however, have not been widely
232 implemented, likely due to the higher effort or cost involved.
233
234 False positives occur when a specimen not harboring Wolbachia is identified as
235 Wolbachia-infected. Several instances have been reported where insect chromosomes
236 carry Wolbachia-derived fragments, presumably from a horizontal gene transfer event
237 that occurred at some point in the host lineage as the result of an active infection that
238 was subsequently lost. The size of the horizontally-transmitted fragment can range from
239 ca. 500 bp to the equivalent of an entire Wolbachia chromosome [97]. In some cases,
240 entire Wolbachia chromosomes have been transferred more than once onto the same
241 host genome [98, 99]. The range of hosts carrying Wolbachia-derived genome
242 fragments is broad and includes several dipterans (tephritids, Glossina morsitans
243 Westwood; Drosophila spp., mosquitoes), other insects, as well as nematodes [97, 98,
244 100-102]. It is therefore desirable to corroborate PCR-based inferences with
245 approaches that detect Wolbachia cells in host tissues. Such microscopy approaches
246 can be based on nucleic acid hybridization [e.g. 103] or antibody-based detection of
247 Wolbachia proteins [e.g. ftsZ; 104, and wsp; 105]. A major drawback of these methods
248 is that they require substantial investment in time and equipment compared to PCR-
249 based approaches. False positives can also occur if the primers targeted at Wolbachia
250 turn out to amplify a fragment of the genome of the host (not derived from Wolbachia) or
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251 of another symbiont of the host. Such false positives are relatively easy to rule out upon
252 sequencing and analysis of the amplified product. Finally, as with any PCR work, false
253 positives can result from contamination of the specimen, the DNA template, or the PCR
254 reagents. Thus, it is important to implement adequate sterile practices and negative
255 controls.
256
257 The above approaches require destruction of specimens for DNA isolation or for tissue
258 fixation. As a rapid and non-destructive alternative, Near-Infrared Spectroscopy (NIR)
259 has been developed for identification of specimens infected with Wolbachia, including
260 the distinction of two different Wolbachia strains [106]. This method, however, requires
261 standardization according to species, sex, age, or any other condition that may affect
262 absorbance, and is not 100% efficient. To our knowledge, this method has not been
263 employed to assess Wolbachia infection in tephritids.
264
265 2.2 Methods to taxonomically characterize Wolbachia strains
266 The main evolutionary lineages of Wolbachia are assigned to “supergroups” [107].
267 Sixteen supergroups have been recognized to date [108, 109]. Supergroups A and B
268 are widespread in arthropods and are common reproductive manipulators [39, 110].
269 Supergroups C and D are obligate mutualists of filarial nematodes, whereas supergroup
270 F is found in both arthropods and nematodes [111]. Other supergroups have more
271 restricted host distributions [112]. Wolbachia are generally compared and classified on
272 the basis of Multilocus Sequence Typing (MLST) systems [110, 113]. The most
273 commonly used MLST is based on the PCR amplification of fragments of five ubiquitous
274 genes: coxA, fbpA, ftsZ, hcpA and gatB. However, this MLST system has limitations, in
275 that not all genes are readily amplified in all Wolbachia strains, and it fails to distinguish
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276 among very closely related strains [109, 112]. Several additional genes commonly
277 amplified and reported are the 16S rRNA, groEL, gltA, and the wolbachia surface protein
278 (wsp) [107, 112, 114, 115]. The wsp gene is highly variable and shows evidence of
279 intragenic recombination [116, 117]. An MLST database (https://pubmlst.org/wolbachia/)
280 is available to compare sequences of alleles for the five MLST loci and the wsp gene.
281 Upon submission to the MLST, new alleles for the wsp and for each of the MLST loci are
282 assigned a unique number. A Wolbachia sequence type (ST) is defined on the basis of
283 allele combinations, with each allele combination assigned a unique ST number.
284
285 Hosts can be infected by one or more distinct strains of Wolbachia. Traditionally, direct
286 Sanger sequencing of PCR products that resulted in sequences with ambiguous base
287 pairs would be subjected to cloning followed by sequencing. The allele intersection
288 analysis method [AIA; 118] can then be used to assign MSLT alleles to Wolbachia
289 strains, but it requires a priori knowledge on the number of strains present. AIA
290 identifies pairs of multiply infected individuals that share Wolbachia and differ by only
291 one strain. Alternative approaches to circumvent cloning include the use of strain-
292 specific primers [e.g. for the wsp gene; 107], or of high throughput sequencing
293 approaches (e.g., Illumina HiSeq or Illumina MiSeq) to sequence MLST or other marker
294 PCR amplicons [e.g. 119, 120]. Primer bias, however, where the fragment of one
295 Wolbachia strain is preferentially amplified over the other, has been reported [118], such
296 that presence of certain Wolbachia strains might be missed.
297
298 Use of the MLST system alone has two major drawbacks. First, strains of Wolbachia
299 sharing identical MLST or wsp alleles can differ from each other at other loci [113, 121].
300 Secondly, the MLST, 16S rRNA, and wsp loci contain limited phylogenetic signal for
301 inferring relationships within Wolbachia supergroups [109]. Therefore, to assess such
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302 intra-ST variation and to infer evolutionary relationships among closely related
303 Wolbachia strains, additional (more variable) loci must be evaluated. The multiple locus
304 variable number tandem repeat analysis developed by Riegler et al. [121] allows
305 distinction of closely related Wolbachia strains based on PCR and gel electrophoresis.
306
307 Whole genome sequencing represents a powerful approach to distinguish closely related
308 Wolbachia strains, infer their evolutionary relationships, test for recombination, and
309 identify genes of interest [e.g. 47, 122]. Due to its fastidious nature [but see 123 for a
310 recent breakthrough] and occurrence of repetitive elements, genome sequencing and
311 assembly of Wolbachia (and other host-associated uncultivable bacteria) has proven
312 difficult. Recent advances, particularly those based on targeted hybrid enrichment [124]
313 prior to high-throughput sequencing [125, 126] have been successfully applied to
314 Wolbachia for short-read technologies [i.e., Illumina; 127, 128]. The combination of
315 targeted hybrid capture and long-read technologies, such as Pacific Biosciences’ Single
316 Molecule Real-Time [e.g. 129] or Oxford Nanopore Technologies’ platforms is expected
317 to greatly advance Wolbachia genomics research.
318
319 2.3 Methods to functionally characterize Wolbachia strains
320 A major challenge to investigating the effects of Wolbachia on a host is to generate
321 Wolbachia-present and Wolbachia-free treatments while controlling for host genetic
322 background. The challenge stems from the difficulty of adding or removing Wolbachia
323 to/from particular hosts. Addition of Wolbachia to a particular host background can be
324 achieved by transinfection [reviewed in 130]. Because the vertical transmission of
325 Wolbachia appears to be dependent on its close association to the host germline,
326 successful artificial transfer of Wolbachia typically relies on injection of cytoplasm from a
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327 donor egg [but see 131] or early embryo into a recipient embryo via microinjection
328 [reviewed in 130]. The success rate of the transinfection procedure is generally very low;
329 in tephritids it is 0–0.09% (calculated as the proportion of injected embryos that emerged
330 as Wolbachia-infected adult females that transmitted Wolbachia to offspring [132, 133,
331 134; Morán-Aceves et al., unpublished data]). The low success rate is generally a result
332 of low survival of injected embryos, the low proportion of Wolbachia-positive survivors,
333 some of which do not transmit Wolbachia to their offspring.
334
335 Intra-species (or between sibling species) transfer of Wolbachia to a particular host
336 nuclear background can also be achieved through introgression, whereby males of the
337 desired background are repeatedly backcrossed with Wolbachia-infected females [e.g.
338 135, 136]. Under this scheme, after eight generations of consistent backcrossing,
339 ~99.6% of the host nuclear background is expected to have been replaced (Fig. 1d).
340 The drawback of this approach is that the mitochondrial genome will not be replaced.
341 Therefore, the effects of mitochondrial type and Wolbachia infection cannot be
342 separated.
343
344 Due to less than perfect transmission, passive loss of Wolbachia in certain host
345 individuals may be used to obtain Wolbachia-free and Wolbachia-infected hosts of
346 equivalent genetic background. Wolbachia removal has also been achieved by
347 “extreme” temperature treatment [e.g. 30°C; 137]. The most common way of removing
348 Wolbachia, however, is achieved via antibiotic treatment, but several potential biases
349 must be addressed [reviewed by 138]. Antibiotic treatment is likely to alter the
350 microbiota, other than Wolbachia, associated with the host. In addition, antibiotics may
351 affect the host in a microbe-independent manner. For instance, antibiotic treatment can
352 affect host mitochondria [139], which in turn can reduce host fitness. A common practice
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353 to circumvent these problems is to wait several generations after antibiotic treatment,
354 and to promote “restoration” of the host’s pre-antibiotic microbiota, excluding Wolbachia
355 (e.g. exposing the insects to the feces of non-treated individuals). Wolbachia does not
356 appear to be efficiently transmitted via ingestion [e.g. 140], but see discussion on
357 horizontal transmission routes below. It is important to monitor Wolbachia infection
358 status of antibiotic-treated host strains, because antibiotics may not always fully remove
359 infection. Instead, they may reduce Wolbachia densities to non-detectable levels in one
360 or few generations [138]; this has been our experience in both Anastrepha and
361 Drosophila (unpublished data).
362
363 Unidirectional CI is tested by comparing the embryo hatching rates of the CI cross
364 (uninfected female X infected male) to that of one or more control crosses. For testing
365 bidirectional CI, the reciprocal crosses of hosts infected by the different Wolbachia
366 strains are assessed. A significantly lower embryo hatching rate of the CI cross(es)
367 compared to that of the control cross(es) constitutes evidence of CI. CI can be partial or
368 complete (100% embryo failure). As with any fitness assay, care must be taken to
369 prevent potential biases, including crowding and age effects; male age can influence CI
370 [33, 81, 141]. Adequate assessment of fertilization must be performed to ensure that
371 failed embryos are not confused with unfertilized eggs. This may require testing for
372 insemination of females that produce no larval progeny [e.g. 134, 142], or exclusion of
373 females that predominantly lay unfertilized eggs, such as old D. melanogaster virgin
374 females [143].
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375 3 Wolbachia in tephritids
376 3.1 Taxonomic distribution of Wolbachia-tephritid associations
377 Based mostly on PCR and sequencing approaches, ~66% of ~86 tephritid species
378 screened have at least one record of positive Wolbachia infection (excluding
379 pseudogenes) in laboratory and natural populations (Table S1; only supergroups A and
380 B have been found in tephritids). For the genus Anastrepha, all but one species (A.
381 ludens) of 17 screened to date harbor Wolbachia [142, 144-148, 149; and this study,
382 150]. Most Anastrepha species harbor Wolbachia strains assigned to supergroup A.
383 Anastrepha striata Schiner and Anastrepha serpentina Wiedemann, however, harbor
384 supergroup B in southern Mexico [147; and this study] and supergroup A in Brazil [146].
385 Up to three Wolbachia sequence types have been detected per locality within
386 morphotypes of the A. fraterculus complex [142, 150], but co-infection of a single
387 individual is generally not observed [except for one report in A. fraterculus; 151].
388
389 Of the ~49 species of Bactrocera that have been examined, ~14 are reported to harbor
390 Wolbachia (supergroup A and/or B) and three (Bactrocera peninsularis Drew & Hancock,
391 Bactrocera perkinsi Drew & Hancock and Bactrocera nigrofemoralis White & Tsuruta)
392 carry what appear to be Wolbachia-derived pseudogenes, but not active infections [101,
393 132, 152-154]. There is also the case of Bactrocera zonata Saunders, B. dorsalis, and
394 Bactrocera correcta Bezzi that have been found to carry both active infections
395 (cytoplasmic) and pseudogenized Wolbachia sequences (Asimakis et al., in review). Up
396 to five Wolbachia strains have been reported in a single individual of Bactrocera ascita
397 Hardy [152], and double/multi infections have been reported in individuals of the
398 following five Bactrocera species in Australia: Bactrocera bryoniae Tryon; Bactrocera
399 decurtans May; Bactrocera frauenfeldi Schiner; Bactrocera neohumeralis Hardy; and
400 Bactrocera strigifinis Walker [101, 153]. Within the genus Bactrocera, polyphagous
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401 species are more likely to harbor Wolbachia compared to stenophagous or
402 monophagous ones [154].
403
404 For the genus Ceratitis, two species have been screened for Wolbachia. No evidence of
405 Wolbachia was found in Ceratitis fasciventris Bezzi. Also, no evidence of infection was
406 found in several populations of C. capitata, the Mediterranean fruit fly (medfly), in the
407 early ‘90s [155]. PCR amplification and sequencing of the 16S rRNA gene in several
408 field and lab specimens of C. capitata from Brazil suggested infection with Wolbachia
409 supergroup A [156]. However, recent thorough surveys of wild populations and lab
410 colonies indicate that Wolbachia is absent in C. capitata from numerous localities in
411 different continents (Table S1). Two out of the six species of Dacus examined to date
412 are reported to harbor Wolbachia: Dacus axanus Hering [153]; and Dacus destillatoria
413 Bezzi [152]. Wolbachia has not been detected in the monotypic genus Dirioxa [101].
414
415 Wolbachia is reported in the four species of Rhagoletis examined to date: Rhagoletis
416 cerasi L.; Rhagoletis pomonella Walsh; Rhagoletis cingulata Loew; and Rhagoletis
417 completa Cresson [133, 157-164]. Both A and B supergroups are found in R. cerasi and
418 R. completa, including a putative A-B recombinant strain, and co-infections are common
419 (e.g. R. cerasi and R. pomonella). In Zeugodacus (formerly Bactrocera), both Z.
420 cucurbitae and Z. diversa are reported to harbor Wolbachia [152, 154; Asimakis et al. in
421 review].
422
423 3.2 Wolbachia prevalence in tephritids (in time/space)
424 Numerous studies report Wolbachia infection frequencies (or data from which this
425 measure can be estimated) in natural populations of tephritids. Few of these studies,
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426 however, have adequate sample sizes for such inferences (e.g. many such studies are
427 based on 10 or fewer individuals). Notwithstanding, inferred Wolbachia prevalence in
428 tephritid populations is highly variable. In Anastrepha, ~10 species harbor at least one
429 population with prevalence ~100%, whereas populations of three species reported lower
430 frequencies (e.g. 88%, 51–60%, and 8.7%) (Table S1). In Bactrocera, one population of
431 B. caudata had a 100% prevalence, whereas all other species with positive Wolbachia
432 results exhibited low prevalence.
433
434 The best studied tephritid system in terms of spatial and temporal variation in Wolbachia
435 prevalence is probably that of R. cerasi in Europe, which was surveyed over a ~15 year
436 period in 59 localities [163]. In an early (1998) survey of Riegler and Stauffer [165], all
437 European R. cerasi individuals were infected by one strain (wCer1), most central and
438 southern European populations harbored an additional strain wCer2 (i.e., were co-
439 infected), and at least an Italian population harbored wCer4 [133]. A rapid spread of
440 wCer2 (a strain associated with cytoplasmic incompatibility) has been detected. Multiple
441 infections in various combinations considering all five known Wolbachia strains have
442 been revealed recently. Samples from Poland, Italy, and Austria, are infected with all
443 five wCer strains, those from Czech Republic and Portugal lack wCer2 only, while the
444 Swiss samples lack wCer3 [162]. Recent analysis of 15 Greek, two German and one
445 Russian population demonstrate fixation for wCer1 in all R. cerasi populations and the
446 presence of complex patterns of infections with the four of the five known wCer strains (1,
447 2, 4, and 5) and the possible existence of new Wolbachia strains for the southernmost
448 European R. cerasi population [i.e., Crete; 158]. Similarly, strain wCin2 (which is
449 identical to wCer2 based on loci examined to date) is fixed in all populations of R.
450 cingulata; a species native to the USA, but introduced into Europe at the end of the 20th
451 century. Invasive populations in Europe harbor wCin1 (identical to wCer1 based on loci
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452 examined to date) at frequencies that vary over space and time (up to 61.5%), as a
453 result of horizontal transfer (multiple events) from R. cerasi [163].
454
455 3.3 Phenotypic effects of Wolbachia in tephritids
456 Despite the numerous reports of Wolbachia in tephritids, the fitness consequences of
457 such associations remain mostly unknown. The studies reporting phenotypic effects of
458 Wolbachia have relied on transinfection and on antibiotic-curing; only two species of
459 tephritids have been successfully transinfected with Wolbachia (Table 1). Evidence of
460 Wolbachia-induced CI has been detected in three species of tephritids. Early studies
461 [166, 167] identified reproductive incompatibilities in R. cerasi that were later attributed
462 to the Wolbachia strain wCer2 [100% embryonic mortality in the CI cross; 165].
463 Artificially transferred Wolbachia (strains wCer2 and wCer4) originally from R. cerasi to
464 C. capitata also resulted in strong CI (100% embryonic mortality). wCer2 in two genetic
465 backgrounds of B. oleae resulted in strong CI as well [132]. In addition, wCer2 and
466 wCer4 are bi-directionally incompatible in C. capitata [133, 134].
467
468 In addition to CI, Wolbachia-infected C. capitata females (Benakeio strain) exhibit higher
469 embryonic mortality (17–32% in crosses with Wolbachia-free males and 65–67% in
470 crosses with Wolbachia-infected males) than their Wolbachia-free counterparts crossed
471 with Wolbachia-free males (12% embryonic mortality). Therefore, it appears that wCer2
472 and wCer4 have additional fertility effects on medfly females, other than CI. It is also
473 possible that these Wolbachia strains can only partially rescue the modification that they
474 induce in sperm [133]. A similar pattern is reported in the Vienna 8 genetic sexing strain
475 (GSS) infected with wCer2 [134]. The wCer2 also causes increased embryo death in
476 non-CI crosses of B. oleae [132].
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477
478 Two studies conducted several years apart [168-170] examined the effects of a single
479 Wolbachia strain (wCer2) on fitness components of two C. capitata genotypes (i.e.,
480 Benakeio and Vienna 8 GSS laboratory lines), as well as the effects of two different
481 Wolbachia strains (wCer2 and wCer4) on a single medfly genotype (Benakeio). The
482 following general patterns emerged (exceptions noted): (a) Wolbachia causes higher
483 egg-to-larva mortality; (b) Wolbachia causes higher egg-to-adult mortality [exception:
484 Vienna 8 GSS + wCer2 in 170]; (c) Wolbachia shortens egg-to-adult development time
485 [exception: Benakeio + wCer2 in 168, 169]. In addition, Sarakatsanou et al. [170] found
486 that Wolbachia shortens both male and female adult lifespan (exception: males of
487 Vienna 8 GSS + wCer2), and reduces life time female net fecundity. However, Kyritsis
488 G.A [168], Kyritsis G.A. et al. [169] reported no effects of Wolbachia infection on adult
489 lifespan, and a reduced fecundity in the case of wCer4 infection only. Even though
490 wCer2 and wCer4 in general tended to have consistent effects on medfly, the magnitude
491 of their effects differed. Collectively, the results from these studies indicate that the
492 effect of Wolbachia infection on life history traits depends both on the C. capitata genetic
493 background and on the Wolbachia strain. Furthermore, inconsistencies between the two
494 studies might be indicative of evolution of the host and/or Wolbachia strain during that
495 period. Adult flight ability and longevity under stress conditions also appear to be
496 determined by the interaction of Wolbachia strain and medfly genotype [168, 169].
497
498 A recent study by Conte et al. [142] examined the phenotypic effects induced by two
499 Wolbachia strains native to A. fraterculus (sp1). No evidence of bidirectional
500 cytoplasmic incompatibility was detected in reciprocal crosses. Ribeiro [137] reported
501 evidence consistent with CI caused by Wolbachia in A. obliqua and in “A. fraterculus sp.
502 1”, which according to wsp sequences, are identical. Nonetheless, confounding effects
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503 of the treatment to remove Wolbachia (removed by exposure of pupae to 30 °C) or other
504 potential biases cannot be ruled out, as all intraspecific crosses involving at least one
505 cured parent resulted in much lower (< 30%) embryo hatching than the intraspecific
506 crosses involving both infected parents (66 and 81% embryo hatching). Whether
507 Wolbachia influences any other fitness aspects of tephritids, including its interactions
508 with other microbes, has not been evaluated.
509
510 Very little is known regarding the effect of Wolbachia on tephritid behavior. Recent tests
511 demonstrated that Wolbachia infection affected male sexual competitiveness. Different
512 Wolbachia strains (wCer2 and wCer4) exerted differential impact on males mating
513 competitiveness, and a single strain (wCer2) had different impact on different medfly
514 genotypes (Benakeio and Vienna 8 GSS laboratory lines) [168, 169].
515
516 3.4 Modes of horizontal transmission of Wolbachia between tephritid hosts
517 Considering the dynamics of Wolbachia associated with arthropods in general, at the
518 population level, Wolbachia appears to be predominantly maintained by vertical
519 transmission. Above the species level, however, the lack of congruence between the
520 host and symbiont phylogenetic trees implies that Wolbachia horizontal transfers and
521 extinctions are common and underlie its widespread taxonomic and geographic
522 distribution [171].
523
524 The possible routes by which Wolbachia may be horizontally acquired by a new host can
525 generally be classified as via ingestion or via a vector. In both cases, to become
526 established as a stable cytoplasmically inherited infection, Wolbachia must cross one or
527 more cell types or tissues. For example, if Wolbachia invaded the host hemolymph
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528 directly as a result of a vector (e.g. parasitoid wasp or ectoparasitic mite), it would have
529 to invade the egg during oogenesis. Similarly, if Wolbachia were acquired via ingestion
530 (e.g. as a result of scavenging), it would have to cross the gut into the hemolymph,
531 before it invaded the egg. Support for the above routes comes from studies reporting:
532 (a) that Wolbachia can remain viable in an extracellular environment and infect mosquito
533 cell lines, as well as ovaries and testes that are maintained ex vivo [172, 173]; (b) that
534 Wolbachia cells injected into Drosophila hemolymph reach the germline after crossing
535 multiple somatic tissues [131]; (c) that Wolbachia can move between parasitic wasp
536 larvae (Trichogramma) sharing the same host egg, and achieve vertical transmission
537 [174]; and (d) that parasitic wasps of the white fly, Bemicia tabaci (Gennadius) can
538 transfer Wolbachia from an infected to a naïve host, as a result of non-lethal probing (i.e.,
539 probing without oviposition), whereby the parasitoid ovipositor or mouthparts function as
540 a “dirty needle” [175].
541
542 No direct evidence of Wolbachia transmission via parasitoids exists in tephritids, but
543 sharing of Wolbachia strains between a parasitoid and several sympatric tephritids [144,
544 153] is consistent with parasitoid-mediated transmission, or transmission from tephritid
545 host to parasitoid [176]. The potential for horizontal transfer of Wolbachia among
546 tephritids via parasitoids is high, due to the multiple instances where a single parasitoid
547 utilizes several different tephritid host species [177-179].
548
549 Wolbachia may invade a new host species via introgressive hybridization between two
550 host species. This mechanism would also transfer mitochondria from the infected to the
551 uninfected species nuclear background, akin to the artificial backcrossing approach
552 described above (Fig. 1d). Ability of tephritids to hybridize in the lab has been reported
553 in numerous species (Table 2), and hybridization in nature has been documented in B.
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554 dorsalis/B. carambolae [180], members of the C. FAR complex [181], R. cingulata/R.
555 cerasi in Europe [182]. Thus, there is potential for wild tephritid populations to acquire
556 Wolbachia infections via hybridization.
557
558 4 Considerations for Wolbachia-based IIT in tephritids
559 4.1 The need for genetic sexing strains (GSS)
560 In both SIT and IIT, the purpose of releasing insects is to reduce the fertility of the wild
561 population, as a result of mating by wild females with sterile or sterility-inducing males,
562 respectively. SIT is most effective when only males are produced and released [183,
563 184], but release of a small percentage of females, which are typically sterilized with
564 lower doses than males, is not considered overly detrimental to the program. The
565 release of only males in a large-scale operation can be accomplished by either killing
566 female zygotes during development or by selectively removing them from the mass-
567 reared population prior to release [185, 186]. In most tephritids, male sex is determined
568 by the presence of the maleness factor on the Y chromosome [187]. GSS based on
569 male-linked (e.g. Y chromosome – autosome (Y;A)) translocations have been developed
570 in a few species to produce conditional female lethality (e.g. temperature sensitive
571 lethality during embryonic development) or a visual sex marker (e.g. pupal color).
572 Unfortunately, despite substantial effort, GSS are still lacking for most tephritid pests.
573 The recent development of CRISPR/Cas9-mediated mutagenesis in tephritids, however,
574 might enable a faster development of tephritid GSS [188, see reviews by 189, 190].
575
576 Whereas the use of GSS is beneficial to SIT programs, it is an absolute necessity for
577 IIT programs, unless mitigating mechanisms can be implemented. In an IIT operation,
578 the release of even a few Wolbachia-infected females could be disastrous, because
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579 these females would be fertile with both wild and released males (which would carry a
580 compatible Wolbachia strain). It is unlikely that any sexing system will achieve zero
581 release of females. Therefore, a mechanism that guarantees that any released females
582 are sterile is necessary. If female sterility is induced with substantially lower radiation
583 doses than male sterility, then radiation could be used to render females sterile without
584 the detrimental effects of the higher doses required to sterilize males, and this is one of
585 the advantages of combining the SIT with the IIT approach [17, 21, 22, 191-193].
586
587 4.2 Choice and evaluation of Wolbachia strains
588 The goal is to generate a mass-reared colony that is infected with a mod+/resc+
589 Wolbachia strain that causes strong incompatibility with the target pest population. The
590 target pest population will be Wolbachia-free (e.g. most C. capitata wild populations and
591 all A. ludens populations) or harbor one or more Wolbachia strains. Below we list
592 recommendations regarding the selection and evaluation of the Wolbachia strain to use
593 in IIT and in SIT/IIT approaches. The target population and the donor colony should be
594 thoroughly screened for Wolbachia, ideally with the higher sensitivity methods described
595 in Section 2.1, to detect low-titer and multi-strain infections. The Wolbachia strain
596 selected should be incompatible with those found in the target population. The selected
597 Wolbachia strain should be artificially transferred to one or more lab colonies,
598 representative of the genetic background of the target pest population. Most cases of
599 successful establishment of stable transinfected insect lines have relied on embryonic
600 microinjection [130]. Introgressive backcrossing might be feasible in scenarios where
601 geographically isolated populations of the same target species harbor distinct Wolbachia
602 strains (e.g. A. striata in Mexico vs. A. striata in Brazil; Table S1).
603
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604 A thorough biological characterization of the artificial host-Wolbachia association should
605 be conducted, as both host background and Wolbachia strain are important
606 determinants of CI expression and other relevant fitness parameters [17, see also 169,
607 reviewed in 194]. The main desired characteristics of the association are: strong
608 induction of CI; no rescue by Wolbachia strain(s) present in target population; few or no
609 fitness costs for parameters relevant to the program. These fitness parameters can be
610 classified into those related to a cost-effective mass production (e.g. female fecundity
611 including embryo hatching success) and those related to the success of released males
612 (e.g. mating and sperm competitive ability). Some host-Wolbachia combinations result
613 in higher female fecundity, such as Drosophila simulans after many generations [60] and
614 Drosophila mauritiana [195]. In contrast, other host-Wolbachia combinations result in
615 lower fertility [e.g. low embryo success in C. capitata and B. oleae; 132, 133, 134].
616 Wolbachia could affect male mating success by influencing assortative mating; a
617 phenomenon detected in some studies of Drosophila [e.g. 196, 197], but not others [e.g.
618 198, 199, 200]. Such influence of Wolbachia on mating preferences was questioned
619 [201] on the basis of evidence that gut microbiota influence assortative mating in
620 Drosophila [198, 201, 202], a finding that itself has been questioned recently [203]. In
621 addition, at least one case has been reported where sperm from Wolbachia-infected
622 males were less competitive [79]. Similarly, Wolbachia-infected D. simulans produce
623 fewer sperm [83]. All of the above parameters should be evaluated under relevant
624 conditions known to interact with Wolbachia, such as temperature and nutrition
625 [reviewed in 17, 204, e.g. 205], interaction with other microorganisms [e.g. 206, 207], as
626 well as male age and mating status [e.g. 208, 209].
627
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628 4.3 Other considerations
629 4.3.1 Species recalcitrant to Wolbachia?
630 Certain species or clades appear to be “resistant” to Wolbachia infection, based on their
631 lack of infection in nature and the failure to achieve stable transfections. The reasons
632 are unknown, but could involve host and/or bacterial factors. For example, none of the
633 members of the diverse repleta species group of Drosophila, comprised mostly of
634 cactophilic flies [210], has ever been found to harbor Wolbachia [211]. Similarly, due to
635 numerous failed transinfection attempts, and the lack of natural infection in wild
636 Anopheles mosquitos, this genus was regarded impervious to Wolbachia [reviewed in
637 130]. This view has been challenged by the recent successful establishment Wolbachia-
638 transfected Anopheles stephensi Liston [62], and the recent discovery of a natural stable
639 Wolbachia infection in Anopheles coluzzii Coetzee & Wilkerson [212]. The lack of
640 natural infections and transinfection failure in A. ludens may reflect a general
641 refractoriness to Wolbachia. Nonetheless, initial attempts to transinfect C. capitata also
642 failed and transfection with Wolbachia was attained later on with different Wolbachia
643 strains [133]. Hence, transinfection attempts with additional Wolbachia strains may result
644 in successful and stable infection in A. ludens as well.
645
646 4.3.2 Potential for target populations to become resistant to sterile males
647 There are two ways in which a target population may become resistant to the effects of
648 release of Wolbachia-infected males. The first is endosymbiont-based, whereby the
649 target population may acquire (e.g. via horizontal transmission) a Wolbachia strain that
650 can rescue the modification (sterility) induced by the strain present in the released
651 males. Generally, such acquisition of a Wolbachia strain during the relatively short
652 lifespan of a release program is highly unlikely. Nonetheless, knowledge on the
653 Wolbachia infection status and strain identity of interacting species, such as other fruit
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654 flies sharing the same host plant and parasitoids, might aid in the selection of Wolbachia
655 strains that are unlikely to be compatible with strains that can potentially be horizontally
656 acquired by the target population. Permanent screening of wild flies from the target
657 population could provide valuable information in order to foresee potential lack of
658 effectiveness of the method.
659
660 The second mechanism is host-based, whereby pre- or post-mating selection on wild
661 females to avoid or reduce fertilization by incompatible sperm [reviewed by 213], acts on
662 standing (or de novo) genetic variation. Evidence consistent with influence of Wolbachia
663 on premating mechanisms comes from the observation that females and males of
664 Drosophila paulistorum Dobzhansky and Pavan exhibit assortative mating according to
665 the Wolbachia strain they harbor [197]. In addition, treatment with antibiotic (which
666 removed Wolbachia) decreases mate discrimination in D. melanogaster [196]. The
667 evolution of resistance to mating with mass-reared males by wild females can be
668 potentially minimized by frequently refreshing the genetic background of the mass-
669 reared strain [214-216], which is a routine process in mass-rearing programs [217].
670 Nonetheless, if the basis for mate discrimination were solely determined by Wolbachia
671 infection state (e.g. if females could distinguish Wolbachia-infected vs. Wolbachia-
672 uninfected males solely on the basis of a Wolbachia-encoded factor), refreshing the fly
673 genetic background of mass-reared strain is unlikely to slow down the evolution of
674 resistance to released males in the target population.
675
676 Several lines of evidence are consistent with the influence of Wolbachia infection on
677 post-mating mechanisms. The existence of genetic incompatibility is predicted to favor
678 polyandry (multiple mating by females) as a female strategy to minimize the probability
679 of her eggs being fertilized by sperm from incompatible males [218]. Consistent with this
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680 prediction, uninfected D. simulans females remate sooner than Wolbachia-infected
681 females [219]. Furthermore, Wolbachia modifies the length of the spermathecal duct of
682 females of the cricket Allonemobius socius Scudder [220], which in turn may afford the
683 female greater control on the outcome of sperm competition [e.g. D. melanogaster; 221].
684 Finally, the fact that host background can influence the CI phenotype [reviewed by 17],
685 suggests that target populations may have genetic variants that are more resistant to CI,
686 which could increase in frequency as a result of the strong selection exerted by the
687 massive release of Wolbachia-infected males.
688
689 4.3.3 Potential alternative ways of implementing Wolbachia-based approaches
690 The recent identification of Wolbachia “CI genes” offers potential alternative ways of
691 harnessing reproductive incompatibility in control of pest tephritids. First, to identify
692 strains with the desired characteristics, at least ability to induce CI, a productive
693 endeavor might be to search for CI loci in the genomes of candidate strains being
694 considered for IIT, prior to artificial transfer efforts. A candidate Wolbachia strain that
695 lacks CI loci homologues, or that contains CI loci homologues that are highly similar to
696 (and thus potentially compatible with) strains present in target population, should be
697 avoided. Secondly, it may be possible in the future to genetically engineer Wolbachia
698 strains with the desired characteristics (e.g. one or more specific CI operons) for IIT
699 programs, or to replace strains used previously in a control program, as a means of
700 addressing resistance phenomena [222]. Finally, a thorough understanding of the CI
701 mechanism might enable the development of IIT based on Wolbachia transgenes, rather
702 than Wolbachia infection. This might be particularly helpful in the control of species that
703 are resistant to Wolbachia infection. Nonetheless, the release of such genetically
704 modified insects might not be feasible due to regulatory hurdles and lack of public
705 acceptance.
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706
707 It has recently been shown that some Wolbachia strains can provide protection against
708 major pathogens and parasites of insects, including RNA viruses and bacteria [66, 71,
709 223, 224]. It is very common for pathogens to appear in rearing facilities. Thus, if a
710 Wolbachia strain could simultaneously cause strong CI and protect against a one or
711 more pathogens (e.g. RNA virus), this would have multiple benefits in an operational
712 Wolbachia-based population suppression program. Furthermore, a Wolbachia strain
713 that does not induce (strong) CI, but protects against pathogens might be desirable in a
714 program that does not rely on CI (e.g. SIT) for population suppression. Wolbachia-
715 mediated pathogen protection would enable high production and quality levels, thereby
716 contributing to a cost-effective and sustainable insect pest management program.
717
718 4.3.4 Potential influence of other heritable symbionts
719 Multiple studies have revealed that although Wolbachia appears to be the dominant
720 facultative heritable symbiont of arthropods, numerous other diverse bacteria (e.g.
721 Spiroplasma, Arsenophonus, Rickettsia, and Cardinium) form such associations with
722 insects, causing a diversity of reproductive and non-reproductive phenotypes [reviewed
723 in 225, 226, 227]. Despite the long-standing recognition that “Wolbachia do not walk
724 alone” [228], many studies of Wolbachia fail to rule out the association of their study
725 organism with other facultative heritable symbionts. Even intensely studied groups in
726 terms of heritable symbionts, such as tsetse flies (genus Glossina), can yield surprises
727 of bacterial associates [e.g. the recent discovery of Spiroplasma in two species of
728 Glossina; 229]. With few exceptions [142, 147; Asimakis et al., unpublished, 230],
729 research on tephritid facultative heritable bacteria has not examined the possibility of
730 players other than Wolbachia. Therefore, we urge that such research include screens
731 for other symbionts, including viruses, protozoans, and fungi.
29 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
732
733 5 Conclusions
734 Given the widespread occurrence of Wolbachia in tephritids and its known fitness
735 consequences in this group of dipterans and in other host taxa, Wolbachia is likely an
736 influential component of tephritid ecology. Further exploration of Wolbachia-tephritid
737 associations is expected to reveal a diversity of effects, as seen in more extensively
738 study systems such as Drosophila and mosquitos. The recent exciting progress in
739 understanding the basis of CI, and many other aspects of Wolbachia biology, should
740 accelerate progress in the development of Wolbachia-based IIT for tephritid species.
741 Nonetheless, IIT applications strongly depend on highly efficient and robust sexing
742 strategies that are not yet available. Thus, IIT will most likely be used in combination
743 with SIT since the use of radiation provides a failsafe mechanism for population
744 suppression programs. In addition, the combined SIT/IIT approach may result in the use
745 of reduced irradiation doses as well as enhanced pathogen protection and male mating
746 competitiveness of mass-reared sterile insects. It is important, however, to not overlook
747 the potential influence of other microbial players on the interaction between Wolbachia
748 and its tephritid hosts.
749
750 Acknowledgements
751 María de los Ángeles Palomeque-Rodas, Luz Verónica García-Fajardo, and Uriel
752 Gallardo-Ortiz (ECOSUR-Unidad Tapachula) provided technical support. Carlos
753 Cáceres (Insect Pest Control Laboratory, Joint FAO/IAEA Division of Nuclear
754 Techniques in Food and Agriculture, Vienna, Austria) and Emilio Hernández-Ortiz
755 (Programa Moscafrut, SAGARPA, SENASICA, IICA) provided materials. Funding was
756 provided by the Joint FAO/IAEA Coordinated Research Project “Use of Symbiotic
30 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
757 Bacteria to Reduce Mass-Rearing Costs and Increase Mating Success in Selected Fruit
758 Pests in Support of SIT Application”. We thank two anonymous reviewers for their
759 valuable suggestions.
760
31 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
761 References 762
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1413 220. Marshall JL. Rapid evolution of spermathecal duct length in the Allonemobius 1414 socius complex of crickets: species, population and Wolbachia effects. PLoS 1415 ONE 2007; 2:e720. 1416 221. Miller GT, Pitnick S. Sperm female coevolution in Drosophila. Science 2002; 1417 298:1230-1233. 1418 222. Sullivan W, O'Neill SL. Microbiology: Manipulation of the manipulators. Nature 1419 2017; 543:182-183. 1420 223. Ye YH, Woolfit M, Rances E, O'Neill SL, McGraw EA. Wolbachia-associated 1421 bacterial protection in the mosquito Aedes aegypti. PLoS Negl Trop Dis 2013; 1422 7:e2362. 1423 224. Martinez J, Longdon B, Bauer S, Chan YS, Miller WJ, Bourtzis K, Teixeira L, 1424 Jiggins FM. Symbionts commonly provide broad spectrum resistance to viruses 1425 in insects: a comparative analysis of Wolbachia strains. PLoS Pathog 2014; 1426 10:e1004369. 1427 225. Hurst GD, Frost CL. Reproductive parasitism: maternally inherited symbionts in a 1428 biparental world. Cold Spring Harbor Perspectives in Biology 2015; 7:a017699. 1429 226. McLean AHC, Parker BJ, Hrcek J, Kavanagh JC, Wellham PAD, Godfray HCJ. 1430 Consequences of symbiont co-infections for insect host phenotypes. J Anim Ecol 1431 2017. 1432 227. Zchori-Fein E, Bourtzis K (Eds.). Manipulative Tenants: Bacteria associated with 1433 Arthropods (a volume in the Frontiers in Microbiology Series). Florida, USA.: 1434 CRC Press, Taylor and Francis Group, LLC; 2011. 1435 228. Duron O, Bouchon D, Boutin S, Bellamy L, Zhou L, Engelstadter J, Hurst GD. 1436 The diversity of reproductive parasites among arthropods: Wolbachia do not walk 1437 alone. BMC Biol 2008; 6:27. 1438 229. Doudoumis V, Blow F, Saridaki A, Augustinos A, Dyer NA, Goodhead I, Solano P, 1439 Rayaisse JB, Takac P, Mekonnen S, Parker AG, Abd-Alla AMM, Darby A, 1440 Bourtzis K, Tsiamis G. Challenging the Wigglesworthia, Sodalis, Wolbachia 1441 symbiosis dogma in tsetse flies: Spiroplasma is present in both laboratory and 1442 natural populations. Sci Rep 2017; 7:4699. 1443 230. Augustinos AA, Drosopoulou E, Gariou-Papalexiou A, Asimakis ED, Cáceres C, 1444 Tsiamis G, Bourtzis K, Penelope M-T, Zacharopoulou A. Cytogenetic and 1445 symbiont analysis of five members of the B. dorsalis complex (Diptera, 1446 Tephritidae): no evidence of chromosomal or symbiont-based speciation events. 1447 Zookeys 2015:273-298. 1448 231. Ebina T, Ohto K. Morphological characters and PCR-RFLP markers in the 1449 interspecific hybrids between Bactrocera carambolae and B. papayae of the B. 1450 dorsalis species complex (Diptera: Tephritidae). Research Bulletin of the Plant 1451 Protection Service 2006. 1452 232. Morrow J, Scott L, Congdon B, Yeates D, Frommer M, Sved J. Close genetic 1453 similarity between two sympatric species of tephritid fruit fly reproductively 1454 isolated by mating time. Evolution 2000; 54:899-910. 1455 233. Shearman D, Frommer M, Morrow J, Raphael K, Gilchrist A. Interspecific 1456 hybridization as a source of novel genetic markers for the sterile insect technique 1457 in Bactrocera tryoni (Diptera: Tephritidae). J Econ Entomol 2010; 103:1071-1079. 1458 234. Pike N, Wang W, Meats A. The likely fate of hybrids of Bactrocera tryoni and 1459 Bactrocera neohumeralis. Heredity 2003; 90:365-370. 1460 235. Meats A, Pike N, An X, Raphael K, Wang W. The effects of selection for early 1461 (day) and late (dusk) mating lines of hybrids of Bactrocera tryoni and Bactrocera 1462 neohumeralis. Genetica 2003; 119:283-293.
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1463 236. Cruickshank L, Jessup AJ, Cruickshank DJ. Interspecific crosses of Bactrocera 1464 tryoni (Froggatt) and Bactrocera jarvisi (Tryon)(Diptera: Tephritidae) in the 1465 laboratory. Aust Entomol 2001; 40:278-280. 1466 237. Erbout N, De Meyer M, Lens L. Hybridization between two polyphagous fruit‐fly 1467 species (Diptera: Tephritidae) causes sex‐biased reduction in developmental 1468 stability. Biol J Linn Soc 2008; 93:579-588. 1469 238. Dos Santos P, Uramoto K, Matioli S. Experimental hybridization among 1470 Anastrepha species (Diptera: Tephritidae): production and morphological 1471 characterization of F1 hybrids. Ann Entomol Soc Am 2001; 94:717-725. 1472 239. Carvalho E, Solferini VN, Matioli SR. Alcohol dehydrogenase activities and 1473 ethanol tolerance in Anastrepha (Diptera, Tephritidae) fruit-fly species and their 1474 hybrids. Genet Mol Biol 2009; 32:177-185. 1475 240. Selivon D, Perondini AL, Morgante JS. Haldane's rule and other aspects of 1476 reproductive isolation observed in the Anastrepha fraterculus complex (Diptera: 1477 Tephritidae). Genet Mol Biol 1999; 22:507-510. 1478 241. Selivon D, Perondini A, Morgante J. A genetic–morphological characterization of 1479 two cryptic species of the Anastrepha fraterculus complex (Diptera: Tephritidae). 1480 Ann Entomol Soc Am 2005; 98:367-381. 1481 242. Segura DF, Vera MT, Rull J, Wornoayporn V, Islam A, Robinson AS. Assortative 1482 mating among Anastrepha fraterculus (Diptera: Tephritidae) hybrids as a 1483 possible route to radiation of the fraterculus cryptic species complex. Biol J Linn 1484 Soc 2011; 102:346-354. 1485 243. Roriz AKP, Japyassú HF, Joachim‐Bravo IS. Incipient speciation in the 1486 Anastrepha fraterculus cryptic species complex: reproductive compatibility 1487 between A. sp. 1 aff. fraterculus and A. sp. 3 aff. fraterculus. Entomol Exp Appl 1488 2017; 162:346-357. 1489 244. Rull J, Tadeo E, Lasa R, Rodríguez CL, Altuzar-Molina A, Aluja M. Experimental 1490 hybridization and reproductive isolation between two sympatric species of 1491 tephritid fruit flies in the Anastrepha fraterculus species group. Insect Sci 2017. 1492 245. Tadeo E, Feder JL, Egan SP, Schuler H, Aluja M, Rull J. Divergence and 1493 evolution of reproductive barriers among three allopatric populations of 1494 Rhagoletis cingulata across eastern North America and Mexico. Entomol Exp 1495 Appl 2015; 156:301-311. 1496 246. Arcella T, Hood GR, Powell TH, Sim SB, Yee WL, Schwarz D, Egan SP, 1497 Goughnour RB, Smith JJ, Feder JL. Hybridization and the spread of the apple 1498 maggot fly, Rhagoletis pomonella (Diptera: Tephritidae), in the northwestern 1499 United States. Evol Appl 2015; 8:834-846. 1500 247. Schwarz D, McPheron BA. When ecological isolation breaks down: sexual 1501 isolation is an incomplete barrier to hybridization between Rhagoletis species. 1502 Evol Ecol Res 2007; 9:829-841. 1503 248. Rull J, Aluja M, Feder JL. Evolution of intrinsic reproductive isolation among four 1504 North American populations of Rhagoletis pomonella (Diptera: Tephritidae). Biol 1505 J Linn Soc 2010; 100:213-223. 1506 249. Rull J, Tadeo E, Aluja M, Guillen L, Egan SP, Feder JL. Hybridization and 1507 sequential components of reproductive isolation between parapatric walnut- 1508 infesting sister species Rhagoletis completa and Rhagoletis zoqui. Biol J Linn 1509 Soc 2012; 107:886-898. 1510 250. Craig TP, Horner JD, Itami JK. Hybridization studies on the host races of Eurosta 1511 solidaginis: implications for sympatric speciation. Evolution 1997; 51:1552-1560.
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1512 251. Joubert DA, Walker T, Carrington LB, De Bruyne JT, Kien DH, Hoang Nle T, 1513 Chau NV, Iturbe-Ormaetxe I, Simmons CP, O'Neill SL. Establishment of a 1514 Wolbachia superinfection in Aedes aegypti mosquitoes as a potential approach 1515 for future resistance management. PLoS Pathog 2016; 12:e1005434. 1516 252. Doudoumis V, Tsiamis G, Wamwiri F, Brelsfoard C, Alam U, Aksoy E, Dalaperas 1517 S, Abd-Alla A, Ouma J, Takac P, Aksoy S, Bourtzis K. Detection and 1518 characterization of Wolbachia infections in laboratory and natural populations of 1519 different species of tsetse flies (genus Glossina). BMC Microbiol 2012; 12 Suppl 1520 1:S3. 1521 253. Hernández-Ortiz V, Canal NA, Salas JOT, Ruíz-Hurtado FM, Dzul-Cauich JF. 1522 Taxonomy and phenotypic relationships of the Anastrepha fraterculus complex in 1523 the Mesoamerican and Pacific Neotropical dominions (Diptera, Tephritidae). 1524 ZooKeys 2015; 2015:95. 1525 254. Drosopoulou E, Koeppler K, Kounatidis I, Nakou I, Papadopoulos N, Bourtzis K, 1526 Mavragani-Tsipidou P. Genetic and cytogenetic analysis of the walnut-husk fly 1527 (Diptera: Tephritidae). Ann Entomol Soc Am 2010; 103:1003-1011. 1528 255. Karimi J, Darsouei R. Presence of the endosymbiont Wolbachia among some 1529 fruit flies (Diptera: Tephritidae) from Iran: a multilocus sequence typing approach. 1530 Journal of Asia-Pacific Entomology 2014; 17:105-112. 1531 256. Yong HS, Song SL, Chua KO, Lim PE. Predominance of Wolbachia 1532 endosymbiont in the microbiota across life stages of Bactrocera latifrons (Insecta: 1533 Tephritidae). Meta Gene 2017; 14:6–17. 1534
47 bioRxiv preprint
Table 1. Successful and unsuccessful Wolbachia transfection attempts in tephritids. doi:
ID of successfully Donor species /strain Recipient species (and Wolbachia Reference certified bypeerreview)istheauthor/funder.Allrightsreserved.Noreuseallowedwithoutpermission. https://doi.org/10.1101/358333 transfected tephritid strain strain) strain
C. capitata WolMed 88.6 R. cerasi C. capitata Benakeio strain wCer2 [133]
C. capitata WolMed S10.3 R. cerasi C. capitata Benakeio strain wCer4 [133]
C. capitata VIENNA 8-E88 C. capitata WolMed 88.6 C. capitata VIENNA 8 Genetic wCer2 [134] ; this versionpostedJune29,2018.
Sexing Strain (GSS)
C. capitata VIENNA 8-E88 Bactrocera oleae wCer2 [132]
N/A (unsuccessful) A. striata A. ludens wAstriB (unpublished)
The copyrightholderforthispreprint(whichwasnot
48 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Table 2. Representative tephritid genera where hybridization between one or more
species has been reported.
Tephritid genera containing species Reference(s)
that can hybridize
Bactrocera [231-236]
Ceratitis [237]
Anastrepha [238-244]
Rhagoletis [245-249]
Eurosta [250]
49 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Figure Legends
Figure 1 a,b. Qualitative illustration of uni- and bidirectional cytoplasmic incompatibility
(CI) on the basis of the Wolbachia infection status of the parent generation. Empty male
and female symbols signify absence of Wolbachia. Blue and yellow ovals represent
distinct Wolbachia strains. Green tick marks = Successful offspring production. Red
crosses = no offspring production. c. A special case of unidirectional incompatibility in
which one Wolbachia strain (i.e., the blue one; equivalent to wRi; see text) can rescue
another strain (i.e., the yellow one; equivalent to wMel), but not vice versa. d.
Backcrossing procedure. Wolbachia infection is indicated by blue oval. Host nuclear
backgrounds are indicated by colors: white represents the initial nuclear background of
Wolbachia-infected host; red (darkest) indicates the host background of the Wolbachia-
free line contributing males every generation. Different shades of red represent the
increasing replacement of “white” nuclear background over backcrossing generations (F1
to F8) by “red” nuclear background.
50 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
Figure 2. Recommended steps to screen for Wolbachia infections in tephritids and
other arthropods. A PCR is performed with Wolbachia-specific primers on DNA isolated
from whole, or parts of (e.g. abdomens), insect. Agarose gel electrophoresis of the PCR
products is used to determine whether the amplicon is of the expected size. Amplicons
of expected size are directly sequenced (e.g. Sanger method). High sequence identity
to other Wolbachia suggests Wolbachia infection. Clean chromatograms are consistent
with a single Wolbachia strain. Otherwise, a cloning step to identify different Wolbachia
alleles is required. Other genes are then amplified and sequenced for further genetic
characterization of the strain. As an optional step, localization of Wolbachia cells within
host tissues can be achieved by Fluorescent In Situ Hybridization (FISH) with
Wolbachia-specific rRNA probe or immunolabeling with antibody specific to Wolbachia
protein. An amplicon of an unexpected size might indicate the occurrence of a
horizontally transmitted Wolbachia genome fragment to the insect chromosome, rather
than a current infection. Similarly, multiple nucleotide polymorphisms (NP) or
insertions/deletions, compared to known strains, are suggestive of Wolbachia
pseudogenes (e.g. horizontally transferred to host genome). This can be further tested
by in situ hybridization of Wolbachia-specific probe to host chromosomes, and/or by
Whole Genome Sequencing of host [images from 98, 195, 251, 252]. Photo of fly
(Anastrepha obliqua) by Fabiola Roque (ECOSUR-UT).
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Supporting Table 1 (Excel spreadsheet)
Compilation of published and unpublished reports of screenings of Wolbachia (and other
heritable bacteria) in pest Tephritidae. In our counts of species, A. fraterculus
morphotypes [253] are regarded as separate species. Additional references not cited in
main text but cited in this table [254-256].
52 bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not a certified by peer review) is the author/funder. All rights reserved.b No reuse allowed without permission. Unidirectional CI Bidirectional CI
c d W+ line W– line
F1: 50%
F2: 75%
F8: >99% bioRxiv preprint doi: https://doi.org/10.1101/358333; this version posted June 29, 2018. The copyright holder for this preprint (which was not
certified by peer review) is the author/funder. All rights reserved. No reuse allowed without permission.
ly
ee fly ee -fr
DNA extraction f -infected
with antibiotic) with
(treated Wolbachia PCR screen using Wolbachia Wolbachia specific primers
Electrophoresis can reveal amplicons of expected and unexpected size smaller or Electrophoresis a�er screening with Wolbachia- ( specific 16S rRNA primers (wspec). The amplicon larger) size of 438bp indicates the presence of intracellular Wolbachia, whereas the 296bp amplicon indicates a puta�ve Horizontal Gene Sanger sequencing Transfer (HGT) event (from Doudoumis et al. 2012)
blast Non-Wolbachia sequence False positive
Posi�ons with more than one chromatogram peak Wolbachia sequence suggest mul�ple intracellular infec�ons and require a cloning step Alignment
94-100% identity Multiple NPs, presence of deletion and/or insertions Indicate presence of intracellular/cytoplasmic Wolbachia Putative Horizontal Gene Transfer Events MLST, wsp, gltA and groEL characterization OR op�onal Confirma�on with Confirma�on of presence of Confirma�on of Wolbachia cell presence Whole Genome Sequencing Wolbachia-derived chromosome in host �ssue via FISH or immunostaining and Assembly fragment (green) in host chromosomes (red) by FISH with Wolbachia horizontally Wolbachia-specific probe transmi�ed fragment (from Brelsfoard et al. 2014)
insect chromosome Fluorescent in situ hybridization Immunostaining of (FISH) of Wolbachia (red) with Wolbachia hsp60 wMel-specific probe (from Fast et al. 2011) (from Joubert et al. 2016)