Wolbachia are a bacterial endosymbiont present in the majority of insect species (Weinert et al., 2015; Werren et al., 2008). While they reside in the germline of both sexes, they are vertically transmitted exclusively through the female germline to all the offspring (Kaur et al., 2021). Consequently, Wolbachia have evolved a number of strategies that provide a selective advantage to infected females. This includes male killing, conversion of males to fertile females, induction of parthenogenesis, and most commonly cytoplasmic incompatibility (CI) (Serbus et al., 2008). CI is a form of Wolbachia-induced paternal effect embryonic lethality. Matings between infected males and uninfected females result in dramatic reductions in egg hatch rates (Hoffmann et al., 1986). However, matings between infected males and infected females, known as the ‘rescue cross’, results in normal egg hatch rates. Additionally, infected females mated with uninfected males results in normal hatch rates. Thus, in a Wolbachia-infected population, infected females have an enormous selective advantage over uninfected females as infected females produce normal hatch rates independent of the infection status of the male (Turelli and Hoffmann, 1991). This phenomenon, as well as Wolbachia-induced male sterility, is currently being employed throughout the world as a strategy for combating pest insects and insect-borne human diseases (Jiggins, 2017; Moretti et al., 2018; Zheng et al., 2019).

Since the discovery of CI and rescue (Ghelelovitach, 1952; Yen and Barr, 1971), there have been a number of insights into their molecular and cellular bases (Shropshire et al., 2020). Cytological studies demonstrate failures in condensation, alignment, and segregation of the paternal chromosomes during the first zygotic division in embryos derived from the CI cross (Breeuwer and Werren, 1990; Callaini et al., 1997; Lassy and Karr, 1996; Reed and Werren, 1995; Ryan and Saul, 1968; Tram et al., 2006; Tram and Sullivan, 2002). Subsequent studies demonstrated defects in the protamine-to-histone transition: deposition of the maternally supplied histone H3.3 is significantly delayed (Landmann et al., 2009). In addition, the male, but not the female, pronucleus of CI-derived embryos exhibits delays in DNA replication, nuclear envelope breakdown, and Cdk1 activation (Tram and Sullivan, 2002). As a result, passage of the male pronucleus through mitosis is delayed relative to the female pronucleus. Molecular insight into the mechanism of CI has come from recent studies demonstrating that a pair of Wolbachia genes originating from integrated viral DNA, the CI factors or Cifs, is likely responsible for CI (Beckmann et al., 2017; LePage et al., 2017). One of these genes, cidB, encodes a deubiquitylating enzyme and the other, cinB, a nuclease (Chen et al., 2020). When the gene pair is expressed in the male germline, paternal chromosome, and embryo abnormalities strikingly similar to Wolbachia-mediated CI are observed (Beckmann et al., 2017; LePage et al., 2017).

In addition to the well-characterized first division mitotic defects, studies in a number of species demonstrate that CI produces additional developmental defects and lethal phases later in embryogenesis (Bonneau et al., 2018; Callaini et al., 1997; Callaini et al., 1996; Duron and Weill, 2006; Jost, 1970; Lassy and Karr, 1996; Wright and Barr, 1981). Studies in wasps in which fertilized eggs develop into diploid females and unfertilized eggs develop into haploid males, provide insight into the different developmental outcomes in CI crosses (Tram et al., 2006). If CI disrupts but does not prevent paternal chromosome segregation, the resulting aneuploid embryos fail to develop. In contrast, if CI results in the complete failure of paternal chromosome segregation, embryos develop into haploid males bearing only the maternal chromosome complement (Ryan and Saul, 1968; Tram et al., 2006). In diplo-diploid organisms (where haploid embryos do not develop to adulthood), complete failure of paternal chromosome segregation in the first division leads to haploid embryos that develop and then subsequently fail to hatch (Bonneau et al., 2018; Callaini et al., 1997; Callaini et al., 1996; Duron and Weill, 2006; Jost, 1970). Additionally, in some organisms such as Drosophila simulans, a small fraction of CI-derived embryos do not undergo any first division defect and consequently hatch as diploids (Lassy and Karr, 1996).

In conjunction with broad developmental abnormalities, embryos developing from CI crosses also experience various cellular defects (Callaini et al., 1996). For example, studies in Drosophila have observed chromosome bridging in CI-derived embryos developing through the pre-blastoderm divisions (nuclear cycles 2–9) (Lassy and Karr, 1996; LePage et al., 2017). Additionally, irregular spindles are observed in syncytial and cellularized blastoderms (nuclear cycles 10–14) from CI crosses (Callaini et al., 1996). Other defects in developed embryos from CI crosses include displaced nuclei, clumped chromatin, and disorganized centrosomes in blastoderms and abnormally condensed nuclei in gastrulating embryos (Callaini et al., 1996). However, due to inherent difficulties in visualizing fertilization and early embryonic development in Drosophila, whether the cellular defects observed in later stage embryos are the direct result of aneuploidy from first division errors or due to a poorly understood second, distinct set of CI-induced defects remained unresolved. Therefore, the extent and timing with which CI induces cellular defects throughout development is an open question.

Here, through a combination of live and fixed analysis, we directly address whether late developmental and cellular defects observed in CI-derived embryos are an outcome of the well-characterized first division errors or caused by a temporally distinct set of CI-induced cell cycle defects. Consistent with previous reports, we find that the majority of embryos derived from the CI cross arrest in the first division. However, we specifically focus our analyses on a population of about one-third of CI-derived embryos that apparently progress normally through the first zygotic and subsequent internal syncytial divisions. While these embryos undergo normal pre-blastoderm divisions, they exhibit significantly increased chromosome segregation defects during the mid-blastula transition, cellularization, and gastrulation. Sequence analysis reveals that ~40% of the CI-derived embryos that reach the blastoderm stage (>nuclear cycle 10) are diploid, having undergone a normal first division to develop with both maternal and paternal chromosomes. Using fluorescence in situ hybridization (FISH) to specifically select these diploid CI-derived embryos, we observe increased chromosome segregation defects during gastrulation, suggesting the late embryonic defects do not strictly require significant first division defects. Crosses to infected females (the rescue cross) reduce the frequencies of the induced division errors. These results reveal that Wolbachia CI-induced defects in the sperm produce developmentally deferred chromosome segregation defects in the late blastoderm divisions. Thus, our studies suggest CI produces temporally distinct early and late chromosome segregation defects. These findings provide insight into the mechanisms and timing of CI.

In addition to the well-characterized early embryonic arrest, a number of reports reveal a large portion of CI-derived embryos undergo substantial embryonic development but then fail to hatch (Bonneau et al., 2018; Callaini et al., 1997; Callaini et al., 1996; Duron and Weill, 2006). In addition, these late-developing CI-derived embryos also exhibit significant cellular defects including chromosome segregation errors, irregular spindles, displaced nuclei, and disorganized centrosomes (Callaini et al., 1997; Callaini et al., 1996; Lassy and Karr, 1996). In diplo-diploid species, late embryonic lethality is widely believed to be due to haploid development explained by the behavior of the paternal chromosomes during the first division (Bonneau et al., 2018; Callaini et al., 1997; Duron and Weill, 2006). While weak CI results in defective paternal chromosome segregation creating aneuploid nuclei that arrest in early embryonic development, strong CI results in complete failure of sister chromosome segregation and haploid nuclei bearing only the maternal chromosome complement (Bonneau et al., 2018; Callaini et al., 1997; Duron and Weill, 2006; Tram et al., 2006). Haploid embryos develop normally to the cellular blastoderm stage but then fail prior to hatching.

However, whether the numerous cellular defects observed in late-developing CI-derived embryos are also the result of first division errors has not been resolved. Here, we addressed this question by analyzing the timing, extent, and causes of mitotic defects observed specifically in late D. simulans embryos derived from CI crosses that seemingly bypass the first division defects and develop normally through early embryogenesis. Collectively, our results suggest that late embryonic defects are the result of a second CI-induced affect temporally distinct from first division errors. As discussed below, this suggests CI may be produced through either (1) a common acute mechanism that acts at both early and late development time points, or (2) separate and distinct early and late mechanisms that do not depend upon one another.

In accord with previous studies in D. simulans (Callaini et al., 1997; Callaini et al., 1996; Lassy and Karr, 1996), we found that there is a second late embryonic lethality associated with CI-derived embryos: between one-fourth and one-third of embryos die after cellularization but before hatching (Figure 1). However, while we found embryos derived from CI crosses were relatively normal during nuclear cycles 2–9 (Figure 2), previous reports (Callaini et al., 1996; Lassy and Karr, 1996) observed pre-blastoderm embryos with substantial mitotic defects. In these experiments, the percentage of abnormal CI-derived embryos fixed after a 6 hr age was similar to that of CI-derived embryos fixed immediately after collection, suggesting pre-blastoderm defects manifest from first division defects (Lassy and Karr, 1996). Therefore, embryos initially undergoing extensive first division defects may develop slowly through pre-blastoderm divisions if they are aged. As we fixed embryos directly after collecting with no aging time, a possible explanation for the low percentage of abnormal pre-blastoderm embryos we observed is that there was not enough time for embryos experiencing substantial first division segregation errors to develop through the pre-blastoderm stages. Instead, our fixing mostly captured the embryos that correspond to the previously reported percentage (~44%) of the CI-derived embryos that develop normally through the pre-blastoderm stage in spite of a later lethal phase (Lassy and Karr, 1996).

As the CI-derived embryos subsequently progressed through the cortical divisions (cycles 10–14), they begin to experience increasingly more severe defects. These late embryonic defects include lagging anaphase chromosomes and chromosome bridging, which directly result in nuclear fallout, and further chromosome bridging during gastrulation (Figures 3 and 4, Figure 4—figure supplement 1). The normal progression of embryos through cycles 2–9 suggests paternal chromosome segregation was not partially defective in the first division. This is because paternal chromosome bridging in the first division would produce aneuploid daughter nuclei bearing chromosome fragments lacking telomeres. The lack of telomeres would result in detectable breakage–fusion–bridge cycles and amplifications of the aneuploidy in subsequent divisions (McClintock, 1941; Titen and Golic, 2008), which we did not observe. Our sequencing analysis of cellularized embryos (Figure 5) confirms that late-stage CI embryos did not experience partial chromosome loss during the first division. Thus, the mitotic defects first observed in cortical syncytial divisions are unlikely a consequence of CI-induced segmental aneuploidy following the first nuclear cycle.

Consequently, our single embryo PCR and whole genome sequencing of cellularized embryos containing a paternally derived egfp transgene revealed late-stage CI embryos were either haploid or diploid (Figure 5). The percentage of diploid embryos closely corresponded to the percentage of embryos hatched, suggesting late embryonic lethality is associated with CI-induced haploidy in D. simulans, as previously reported (Callaini et al., 1997). Importantly, this experiment also demonstrated that the detection of paternally derived chromosomes in that late-stage CI embryos could be used to distinguish between embryos that had experienced first division defects (haploid = only maternal chromosomes, no paternal chromosomes) and embryos that had not experienced any first division defects (diploid = both maternal and paternal chromosomes). As discussed below, this allowed us to uncouple CI-induced late embryo defects from first division defects.

In spite of the strong association between haploidy and lethality, first division-induced haploidy in and of itself cannot explain the defects we observed in CI-derived blastoderm and gastrulating embryos. This is because (1) haploidy is not intrinsically harmful to mitotic divisions in Drosophila. For example, in some Drosophila mutations that induce haploidy, chromosome segregation occurs normally during cortical divisions (Tang et al., 2017). Additionally, meiosis II—essentially a mitotic division of a haploid nucleus—is highly accurate by necessity. Furthermore, (2) any downstream effects of haploidy—such as changes to zygotic gene copy number or loss of zygotic heterozygosity—cannot explain defects first detected in syncytial cortical divisions (cycles 10–13), which do not require zygotic transcription (Yuan et al., 2016). In contrast, the observed defects in CI-derived late embryos are more likely due to a second temporally deferred CI-induced defect.

In support of this hypothesis, our observation of increased chromosome segregation errors in diploid CI gastrulating embryos bearing paternally derived Y chromosomes establishes that the defects observed in late-stage CI embryos are not limited to haploids (Figure 6). Instead, defects are present in diploid late-stage embryos. Significantly, as discussed above for the paternally derived egfp transgene, detection of the Y chromosome by FISH allowed us to select late-stage diploid embryos that had ‘escaped’ first division defects and instead continued development with both paternal and maternal chromosome complements. Therefore, the significant increase in mitotic errors observed in diploid CI-derived embryos relative to wild-type-derived embryos demonstrates the existence of a second, CI-induced defect, temporally and possibly mechanistically distinct from the first division defect (Figure 7A, B). Significantly, maternally supplied Wolbachia independently rescues this later second defect as well (Figure 7C).

Figure 7

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Interestingly, we also observed several non-Y-bearing gastrulating embryos from CI crosses that had extensive chromosome segregation errors beyond what we had observed for the diploid Y-bearing gastrulating embryos (Figure 6—figure supplement 2). Non-Y-bearing embryos may either be diploid (XX) or haploid (XØ). If these embryos were haploid, this observation would suggest that CI could affect both the maternal chromosomes and paternal chromosomes.

One intriguing aspect of the second CI-induced defect is that the embryos progress normally through the early mitotic cycles and then begin to exhibit mitotic defects in the blastoderm stage. The explanation is likely a consequence of the dramatic structural and regulator cell cycle modifications that occur when the dividing nuclei arrive at the cortex (Farrell and O’Farrell, 2014). These include heterochromatin formation, initiation of late origins of replication, slowing of DNA replication, activation of zygotic transcription, and metaphase furrow formation (Farrell and O’Farrell, 2014; Li et al., 2014; Riggs et al., 2003; Seller et al., 2019; Seller and O’Farrell, 2018). The phenotype of numerous maternal-effect mutations that either rely on or disrupt these processes is strikingly similar to the defects observed in CI embryos: normal early pre-cortical divisions followed by extensive mitotic errors and nuclear fallout during the cortical blastoderm divisions (Sullivan and Theurkauf, 1995). For example, because of the slowing of DNA replication during the cortical divisions, activation of the S-phase checkpoint is specifically required during this stage. Maternal-effect mutants that disrupt this checkpoint progress normally through the early divisions but exhibit anaphase bridging and nuclear fallout during the late cortical blastoderm divisions as a result of entering metaphase with incompletely replicated chromosomes (Fogarty et al., 1997; Fogarty et al., 1994). Given the similarity of this phenotype, both in timing and chromosome dynamics, CI-induced late division defects may be due to improper or abnormally slow chromosome replication. Additionally, defects in other events specific to the cortical blastoderm cycles, may also contribute directly or indirectly to CI-induced defects. For example, studies of hybrid incompatibilities between D. simulans and D. melanogaster show that heterochromatin establishment may be particularly sensitive, and its disruption can result in defects strikingly similar to the late CI defects (Ferree and Barbash, 2009). Other important processes, such as those involved in DNA integrity, protein turnover, and cell cycle timing, may be also involved (Momtaz et al., 2020).

In considering the mechanism by which paternal Wolbachia may induce these defects, the observation that CI-derived blastoderm embryos progress normally through pre-cortical divisions must be noted. One potential explanation is that the chromosomes in CI-derived embryos could be epigenetically marked by Wolbachia in the paternal germline. This mark would then persist through the pre-cortical divisions and become disruptive during blastoderm divisions. Interestingly, Wolbachia infection results in altered DNA methylation levels in certain wasps, mosquitos, leafhoppers, and Drosophila (LePage et al., 2014; Negri et al., 2009; Wu et al., 2020; Ye et al., 2013). Should Wolbachia in the male germline similarly change the low naturally occurring methylation levels in D. simulans (Deshmukh et al., 2018), the altered mark may become disruptive in blastoderm divisions, potentially by distorting heterochromatin establishment. However, DNA methylation does not appear to contribute to CI levels (LePage et al., 2014).

An alternative explanation for the specificity of the late blastoderm defects comes from studies into hybrid dysgenesis in D. melanogaster in which unregulated mobilization of transposable elements results in a spectrum of genetic and developmental defects in the germlines of dysgenic progeny (Kidwell et al., 1977). Transposition in progeny can be suppressed when maternally supplied small RNAs mediate silencing of the transposable element (Czech and Hannon, 2016). This silencing is associated with increased H3K9 methylation, increased heterochromatin levels, and altered splicing (Le Thomas et al., 2013; Sienski et al., 2012; Teixeira et al., 2017). Given small RNAs can affect chromosomes in trans (Hermant et al., 2015), CI may induce a similar small RNA pathway that could epigenetically alter both paternal and maternal chromosomes prior to the first division. As the blastoderm divisions do not require zygotic transcription (Yuan et al., 2016), it is unlikely an epigenetic alteration, if occurring, would cause defects via disrupted transcription. Instead, as discussed above, an epigenetic change may disrupt key aspects of the mid-blastula transition, which in turn could result in the observed errors.

Insight into the molecular mechanism of CI came with the discovery of the Wolbachia-encoded Cifs that play a key role in CI and rescue (Beckmann et al., 2017; LePage et al., 2017). A combination of molecular, genetic, and biochemical studies provided compelling evidence that the Wolbachia encoded genes, CidB and CidA, act as a paternally supplied toxin and maternally supplied anti-toxin, respectively (Beckmann et al., 2017; Horard et al., 2022; Wang et al., 2022). However, a toxin/anti-toxin model for CI does not easily explain the cortical blastoderm defects occurring after many rounds of normal mitotic cycles. This is because a paternally supplied toxin is expected to be diluted with every round of division, and therefore its induced defects would decrease over time. Similarly, a second set of CI/rescue elements, CinA and CinB, is also proposed to act in a toxin/anti-toxin manner (Chen et al., 2019; Sun et al., 2022). An alternative possibility is that Cifs may epigenetically modify paternal and maternal chromosomes to mediate CI and rescue (Kaur et al., 2022).

How and if these proteins also contribute to CI-induced late-embryo defects remains to be determined. It is possible that early and late defects are caused by common acute mechanisms involving the known Cifs. This may be due to an induced chromosomal change that produces defects at different times given natural variation in developmental processes among a population of embryos. In contrast, chromosome segregation errors in diploid embryos that have progressed normally through the first division may suggest minimal Cif activity and an additional set of Wolbachia genes that induce late-embryo defects. Unlike the first division errors, CI-induced mitotic defects in late embryos do not appear to result from abnormal condensation, alignment, or timing of metaphase exit (Figure 4). Instead, the observed chromosome bridging is strikingly similar to embryos exposed to the DNA replication inhibitor aphidicolin (Farrell et al., 2012; Fasulo et al., 2012), suggesting CI-derived blastoderm embryos may be entering anaphase with incompletely replicated chromosomes. These differences could suggest a separate proximate cause for early and late defects, or provide additional insight into Cif-induced defects. Thus, any model of CI and rescue, be it toxin/anti-toxin, lock-key, titration, or timing, must account for the fact that some effects of Wolbachia on the sperm are not realized until hours and many cell cycles later when the embryos progress through the late blastoderm divisions and the mid-blastula transition.

Sequencing data have been deposited in the Sequence Read Archive (BioProject PRJNA848235). Figure 5—source data 1 contains the raw unedited gel used to generate Figure 5B as well as Figure 5 prepared with the raw gel. Figure 5—source data 2 contains numerical source data to generate Figure 5E. All reagents used are available upon request.

1. 1. Warecki B
   2. Titen S
   3. Alam MS
   4. Vega G
   5. Lemseffer N
   6. Hug K
   7. Minden JS
   8. Sullivan W

   (2022) NCBI BioProject

   ID PRJNA848235. Drosophila simulans cellularized blastoderm embryos.

   http://www.ncbi.nlm.nih.gov/bioproject/?term=PRJNA848235

1. 1. Beckmann JF
   2. Ronau JA
   3. Hochstrasser M

   (2017) A wolbachia deubiquitylating enzyme induces cytoplasmic incompatibility

   Nature Microbiology 2:17007.

   https://doi.org/10.1038/nmicrobiol.2017.7

   • PubMed
   • Google Scholar
2. 1. Bonneau M
   2. Atyame C
   3. Beji M
   4. Justy F
   5. Cohen-Gonsaud M
   6. Sicard M
   7. Weill M

   (2018) Culex pipiens crossing type diversity is governed by an amplified and polymorphic operon of wolbachia

   Nature Communications 9:319.

   https://doi.org/10.1038/s41467-017-02749-w

   • PubMed
   • Google Scholar
3. 1. Breeuwer JA
   2. Werren JH

   (1990) Microorganisms associated with chromosome destruction and reproductive isolation between two insect species

   Nature 346:558–560.

   https://doi.org/10.1038/346558a0

   • PubMed
   • Google Scholar
4. 1. Callaini G
   2. Riparbelli MG
   3. Giordano R
   4. Dallai R

   (1996) Mitotic defects associated with cytoplasmic incompatibility in Drosophila simulans

   Journal of Invertebrate Pathology 67:55–64.

   https://doi.org/10.1006/jipa.1996.0009

   • Google Scholar
5. 1. Callaini G
   2. Dallai R
   3. Riparbelli MG

   (1997) Wolbachia-induced delay of paternal chromatin condensation does not prevent maternal chromosomes from entering anaphase in incompatible crosses of Drosophila simulans

   Journal of Cell Science 110 ( Pt 2):271–280.

   https://doi.org/10.1242/jcs.110.2.271

   • PubMed
   • Google Scholar
6. 1. Casper-Lindley C
   2. Kimura S
   3. Saxton DS
   4. Essaw Y
   5. Simpson I
   6. Tan V
   7. Sullivan W

   (2011) Rapid fluorescence-based screening for wolbachia endosymbionts in Drosophila germ line and somatic tissues

   Applied and Environmental Microbiology 77:4788–4794.

   https://doi.org/10.1128/AEM.00215-11

   • PubMed
   • Google Scholar
7. 1. Chen H
   2. Ronau JA
   3. Beckmann JF
   4. Hochstrasser M

   (2019) A wolbachia nuclease and its binding partner provide A distinct mechanism for cytoplasmic incompatibility

   PNAS 116:22314–22321.

   https://doi.org/10.1073/pnas.1914571116

   • PubMed
   • Google Scholar
8. 1. Chen H
   2. Zhang M
   3. Hochstrasser M

   (2020) The biochemistry of cytoplasmic incompatibility caused by endosymbiotic bacteria

   Genes 11:E852.

   https://doi.org/10.3390/genes11080852

   • PubMed
   • Google Scholar
9. 1. Clark AG
   2. Eisen MB
   3. Smith DR
   4. Bergman CM
   5. Oliver B
   6. Markow TA
   7. Kaufman TC
   8. Kellis M
   9. Gelbart W
   10. Iyer VN
   11. Pollard DA
   12. Sackton TB
   13. Larracuente AM
   14. Singh ND
   15. Abad JP
   16. Abt DN
   17. Adryan B
   18. Aguade M
   19. Akashi H
   20. Anderson WW
   21. Aquadro CF
   22. Ardell DH
   23. Arguello R
   24. Artieri CG
   25. Barbash DA
   26. Barker D
   27. Barsanti P
   28. Batterham P
   29. Batzoglou S
   30. Begun D
   31. Bhutkar A
   32. Blanco E
   33. Bosak SA
   34. Bradley RK
   35. Brand AD
   36. Brent MR
   37. Brooks AN
   38. Brown RH
   39. Butlin RK
   40. Caggese C
   41. Calvi BR
   42. Bernardo de Carvalho A
   43. Caspi A
   44. Castrezana S
   45. Celniker SE
   46. Chang JL
   47. Chapple C
   48. Chatterji S
   49. Chinwalla A
   50. Civetta A
   51. Clifton SW
   52. Comeron JM
   53. Costello JC
   54. Coyne JA
   55. Daub J
   56. David RG
   57. Delcher AL
   58. Delehaunty K
   59. Do CB
   60. Ebling H
   61. Edwards K
   62. Eickbush T
   63. Evans JD
   64. Filipski A
   65. Findeiss S
   66. Freyhult E
   67. Fulton L
   68. Fulton R
   69. Garcia ACL
   70. Gardiner A
   71. Garfield DA
   72. Garvin BE
   73. Gibson G
   74. Gilbert D
   75. Gnerre S
   76. Godfrey J
   77. Good R
   78. Gotea V
   79. Gravely B
   80. Greenberg AJ
   81. Griffiths-Jones S
   82. Gross S
   83. Guigo R
   84. Gustafson EA
   85. Haerty W
   86. Hahn MW
   87. Halligan DL
   88. Halpern AL
   89. Halter GM
   90. Han MV
   91. Heger A
   92. Hillier L
   93. Hinrichs AS
   94. Holmes I
   95. Hoskins RA
   96. Hubisz MJ
   97. Hultmark D
   98. Huntley MA
   99. Jaffe DB
   100. Jagadeeshan S
   101. Jeck WR
   102. Johnson J
   103. Jones CD
   104. Jordan WC
   105. Karpen GH
   106. Kataoka E
   107. Keightley PD
   108. Kheradpour P
   109. Kirkness EF
   110. Koerich LB
   111. Kristiansen K
   112. Kudrna D
   113. Kulathinal RJ
   114. Kumar S
   115. Kwok R
   116. Lander E
   117. Langley CH
   118. Lapoint R
   119. Lazzaro BP
   120. Lee S-J
   121. Levesque L
   122. Li R
   123. Lin C-F
   124. Lin MF
   125. Lindblad-Toh K
   126. Llopart A
   127. Long M
   128. Low L
   129. Lozovsky E
   130. Lu J
   131. Luo M
   132. Machado CA
   133. Makalowski W
   134. Marzo M
   135. Matsuda M
   136. Matzkin L
   137. McAllister B
   138. McBride CS
   139. McKernan B
   140. McKernan K
   141. Mendez-Lago M
   142. Minx P
   143. Mollenhauer MU
   144. Montooth K
   145. Mount SM
   146. Mu X
   147. Myers E
   148. Negre B
   149. Newfeld S
   150. Nielsen R
   151. Noor MAF
   152. O’Grady P
   153. Pachter L
   154. Papaceit M
   155. Parisi MJ
   156. Parisi M
   157. Parts L
   158. Pedersen JS
   159. Pesole G
   160. Phillippy AM
   161. Ponting CP
   162. Pop M
   163. Porcelli D
   164. Powell JR
   165. Prohaska S
   166. Pruitt K
   167. Puig M
   168. Quesneville H
   169. Ram KR
   170. Rand D
   171. Rasmussen MD
   172. Reed LK
   173. Reenan R
   174. Reily A
   175. Remington KA
   176. Rieger TT
   177. Ritchie MG
   178. Robin C
   179. Rogers Y-H
   180. Rohde C
   181. Rozas J
   182. Rubenfield MJ
   183. Ruiz A
   184. Russo S
   185. Salzberg SL
   186. Sanchez-Gracia A
   187. Saranga DJ
   188. Sato H
   189. Schaeffer SW
   190. Schatz MC
   191. Schlenke T
   192. Schwartz R
   193. Segarra C
   194. Singh RS
   195. Sirot L
   196. Sirota M
   197. Sisneros NB
   198. Smith CD
   199. Smith TF
   200. Spieth J
   201. Stage DE
   202. Stark A
   203. Stephan W
   204. Strausberg RL
   205. Strempel S
   206. Sturgill D
   207. Sutton G
   208. Sutton GG
   209. Tao W
   210. Teichmann S
   211. Tobari YN
   212. Tomimura Y
   213. Tsolas JM
   214. Valente VLS
   215. Venter E
   216. Venter JC
   217. Vicario S
   218. Vieira FG
   219. Vilella AJ
   220. Villasante A
   221. Walenz B
   222. Wang J
   223. Wasserman M
   224. Watts T
   225. Wilson D
   226. Wilson RK
   227. Wing RA
   228. Wolfner MF
   229. Wong A
   230. Wong GK-S
   231. Wu C-I
   232. Wu G
   233. Yamamoto D
   234. Yang H-P
   235. Yang S-P
   236. Yorke JA
   237. Yoshida K
   238. Zdobnov E
   239. Zhang P
   240. Zhang Y
   241. Zimin AV
   242. Baldwin J
   243. Abdouelleil A
   244. Abdulkadir J
   245. Abebe A
   246. Abera B
   247. Abreu J
   248. Acer SC
   249. Aftuck L
   250. Alexander A
   251. An P
   252. Anderson E
   253. Anderson S
   254. Arachi H
   255. Azer M
   256. Bachantsang P
   257. Barry A
   258. Bayul T
   259. Berlin A
   260. Bessette D
   261. Bloom T
   262. Blye J
   263. Boguslavskiy L
   264. Bonnet C
   265. Boukhgalter B
   266. Bourzgui I
   267. Brown A
   268. Cahill P
   269. Channer S
   270. Cheshatsang Y
   271. Chuda L
   272. Citroen M
   273. Collymore A
   274. Cooke P
   275. Costello M
   276. D’Aco K
   277. Daza R
   278. De Haan G
   279. DeGray S
   280. DeMaso C
   281. Dhargay N
   282. Dooley K
   283. Dooley E
   284. Doricent M
   285. Dorje P
   286. Dorjee K
   287. Dupes A
   288. Elong R
   289. Falk J
   290. Farina A
   291. Faro S
   292. Ferguson D
   293. Fisher S
   294. Foley CD
   295. Franke A
   296. Friedrich D
   297. Gadbois L
   298. Gearin G
   299. Gearin CR
   300. Giannoukos G
   301. Goode T
   302. Graham J
   303. Grandbois E
   304. Grewal S
   305. Gyaltsen K
   306. Hafez N
   307. Hagos B
   308. Hall J
   309. Henson C
   310. Hollinger A
   311. Honan T
   312. Huard MD
   313. Hughes L
   314. Hurhula B
   315. Husby ME
   316. Kamat A
   317. Kanga B
   318. Kashin S
   319. Khazanovich D
   320. Kisner P
   321. Lance K
   322. Lara M
   323. Lee W
   324. Lennon N
   325. Letendre F
   326. LeVine R
   327. Lipovsky A
   328. Liu X
   329. Liu J
   330. Liu S
   331. Lokyitsang T
   332. Lokyitsang Y
   333. Lubonja R
   334. Lui A
   335. MacDonald P
   336. Magnisalis V
   337. Maru K
   338. Matthews C
   339. McCusker W
   340. McDonough S
   341. Mehta T
   342. Meldrim J
   343. Meneus L
   344. Mihai O
   345. Mihalev A
   346. Mihova T
   347. Mittelman R
   348. Mlenga V
   349. Montmayeur A
   350. Mulrain L
   351. Navidi A
   352. Naylor J
   353. Negash T
   354. Nguyen T
   355. Nguyen N
   356. Nicol R
   357. Norbu C
   358. Norbu N
   359. Novod N
   360. O’Neill B
   361. Osman S
   362. Markiewicz E
   363. Oyono OL
   364. Patti C
   365. Phunkhang P
   366. Pierre F
   367. Priest M
   368. Raghuraman S
   369. Rege F
   370. Reyes R
   371. Rise C
   372. Rogov P
   373. Ross K
   374. Ryan E
   375. Settipalli S
   376. Shea T
   377. Sherpa N
   378. Shi L
   379. Shih D
   380. Sparrow T
   381. Spaulding J
   382. Stalker J
   383. Stange-Thomann N
   384. Stavropoulos S
   385. Stone C
   386. Strader C
   387. Tesfaye S
   388. Thomson T
   389. Thoulutsang Y
   390. Thoulutsang D
   391. Topham K
   392. Topping I
   393. Tsamla T
   394. Vassiliev H
   395. Vo A
   396. Wangchuk T
   397. Wangdi T
   398. Weiand M
   399. Wilkinson J
   400. Wilson A
   401. Yadav S
   402. Young G
   403. Yu Q
   404. Zembek L
   405. Zhong D
   406. Zimmer A
   407. Zwirko Z
   408. Jaffe DB
   409. Alvarez P
   410. Brockman W
   411. Butler J
   412. Chin C
   413. Gnerre S
   414. Grabherr M
   415. Kleber M
   416. Mauceli E
   417. MacCallum I
   418. Drosophila 12 Genomes Consortium

   (2007) Evolution of genes and genomes on the Drosophila phylogeny

   Nature 450:203–218.

   https://doi.org/10.1038/nature06341

   • PubMed
   • Google Scholar
10. 1. Czech B
    2. Hannon GJ

    (2016) One loop to rule them all: the ping-pong cycle and pirna-guided silencing

    Trends in Biochemical Sciences 41:324–337.

    https://doi.org/10.1016/j.tibs.2015.12.008

    • PubMed
    • Google Scholar
11. 1. Debec A

    (1978) Haploid cell cultures of Drosophila melanogaster

    Nature 274:255–256.

    https://doi.org/10.1038/274255a0

    • PubMed
    • Google Scholar
12. 1. Deshmukh S
    2. Ponnaluri VC
    3. Dai N
    4. Pradhan S
    5. Deobagkar D

    (2018) Levels of DNA cytosine methylation in the Drosophila genome

    PeerJ 6:e5119.

    https://doi.org/10.7717/peerj.5119

    • PubMed
    • Google Scholar
13. 1. Duron O
    2. Weill M

    (2006) Wolbachia infection influences the development of culex pipiens embryo in incompatible crosses

    Heredity 96:493–500.

    https://doi.org/10.1038/sj.hdy.6800831

    • PubMed
    • Google Scholar
14. 1. Farrell JA
    2. Shermoen AW
    3. Yuan K
    4. O’Farrell PH

    (2012) Embryonic onset of late replication requires cdc25 down-regulation

    Genes & Development 26:714–725.

    https://doi.org/10.1101/gad.186429.111

    • PubMed
    • Google Scholar
15. 1. Farrell JA
    2. O’Farrell PH

    (2014) From egg to gastrula: how the cell cycle is remodeled during the Drosophila mid-blastula transition

    Annual Review of Genetics 48:269–294.

    https://doi.org/10.1146/annurev-genet-111212-133531

    • PubMed
    • Google Scholar
16. 1. Fasulo B
    2. Koyama C
    3. Yu KR
    4. Homola EM
    5. Hsieh TS
    6. Campbell SD
    7. Sullivan W

    (2012) Chk1 and wee1 kinases coordinate DNA replication, chromosome condensation, and anaphase entry

    Molecular Biology of the Cell 23:1047–1057.

    https://doi.org/10.1091/mbc.E11-10-0832

    • PubMed
    • Google Scholar
17. 1. Ferree PM
    2. Barbash DA

    (2009) Species-specific heterochromatin prevents mitotic chromosome segregation to cause hybrid lethality in Drosophila

    PLOS Biology 7:e1000234.

    https://doi.org/10.1371/journal.pbio.1000234

    • PubMed
    • Google Scholar
18. 1. Foe VE

    (1989)

    Mitotic domains reveal early commitment of cells in Drosophila embryos

    Development 107:1–22.

    • PubMed
    • Google Scholar
19. 1. Fogarty P
    2. Kalpin RF
    3. Sullivan W

    (1994) The Drosophila maternal-effect mutation grapes causes a metaphase arrest at nuclear cycle 13

    Development 120:2131–2142.

    https://doi.org/10.1242/dev.120.8.2131

    • PubMed
    • Google Scholar
20. 1. Fogarty P
    2. Campbell SD
    3. Abu-Shumays R
    4. Phalle BS
    5. Yu KR
    6. Uy GL
    7. Goldberg ML
    8. Sullivan W

    (1997) The Drosophila grapes gene is related to checkpoint gene chk1/rad27 and is required for late syncytial division fidelity

    Current Biology 7:418–426.

    https://doi.org/10.1016/s0960-9822(06)00189-8

    • PubMed
    • Google Scholar
21. 1. Ghelelovitach S

    (1952)

    Genetic determinism of sterility in the cross-breeding of various strains of culex autogenicus roubaud

    Comptes Rendus Hebdomadaires Des Seances de l’Academie Des Sciences 234:2386–2388.

    • PubMed
    • Google Scholar
22. 1. Hermant C
    2. Boivin A
    3. Teysset L
    4. Delmarre V
    5. Asif-Laidin A
    6. van den Beek M
    7. Antoniewski C
    8. Ronsseray S

    (2015) Paramutation in Drosophila requires both nuclear and cytoplasmic actors of the pirna pathway and induces cis-spreading of pirna production

    Genetics 201:1381–1396.

    https://doi.org/10.1534/genetics.115.180307

    • PubMed
    • Google Scholar
23. 1. Hoffmann AA
    2. Turelli M
    3. Simmons GM

    (1986) UNIDIRECTIONAL incompatibility between populations of Drosophila simulans

    Evolution; International Journal of Organic Evolution 40:692–701.

    https://doi.org/10.1111/j.1558-5646.1986.tb00531.x

    • PubMed
    • Google Scholar
24. 1. Holtzman S
    2. Miller D
    3. Eisman R
    4. Kuwayama H
    5. Niimi T
    6. Kaufman T

    (2010) Transgenic tools for members of the genus Drosophila with sequenced genomes

    Fly 4:349–362.

    https://doi.org/10.4161/fly.4.4.13304

    • PubMed
    • Google Scholar
25. 1. Horard B
    2. Terretaz K
    3. Gosselin-Grenet AS
    4. Sobry H
    5. Sicard M
    6. Landmann F
    7. Loppin B

    (2022) Paternal transmission of the wolbachia cidb toxin underlies cytoplasmic incompatibility

    Current Biology 32:1319–1331.

    https://doi.org/10.1016/j.cub.2022.01.052

    • PubMed
    • Google Scholar
26. 1. Illmensee K
    2. Mahowald AP

    (1974) Transplantation of posterior polar plasm in Drosophila: induction of germ cells at the anterior pole of the egg

    PNAS 71:1016–1020.

    https://doi.org/10.1073/pnas.71.4.1016

    • PubMed
    • Google Scholar
27. 1. Jiggins FM

    (2017) The spread of wolbachia through mosquito populations

    PLOS Biology 15:e2002780.

    https://doi.org/10.1371/journal.pbio.2002780

    • PubMed
    • Google Scholar
28. 1. Jost E

    (1970) Genetic investigations on the incompatibility in the culex pipiens complex

    TAG. Theoretical and Applied Genetics. Theoretische Und Angewandte Genetik 40:251–256.

    https://doi.org/10.1007/BF00282034

    • PubMed
    • Google Scholar
29. 1. Kaur R
    2. Shropshire JD
    3. Cross KL
    4. Leigh B
    5. Mansueto AJ
    6. Stewart V
    7. Bordenstein SR
    8. Bordenstein SR

    (2021) Living in the endosymbiotic world of wolbachia: A centennial review

    Cell Host & Microbe 29:879–893.

    https://doi.org/10.1016/j.chom.2021.03.006

    • PubMed
    • Google Scholar
30. 1. Kaur R
    2. Leigh BA
    3. Ritchie IT
    4. Bordenstein SR

    (2022) The cif proteins from wolbachia prophage WO modify sperm genome integrity to establish cytoplasmic incompatibility

    PLOS Biology 20:e3001584.

    https://doi.org/10.1371/journal.pbio.3001584

    • PubMed
    • Google Scholar
31. 1. Kidwell MG
    2. Kidwell JF
    3. Sved JA

    (1977) Hybrid dysgenesis in Drosophila melanogaster: A syndrome of aberrant traits including mutation, sterility and male recombination

    Genetics 86:813–833.

    https://doi.org/10.1093/genetics/86.4.813

    • PubMed
    • Google Scholar
32. 1. Klasson L
    2. Westberg J
    3. Sapountzis P
    4. Näslund K
    5. Lutnaes Y
    6. Darby AC
    7. Veneti Z
    8. Chen L
    9. Braig HR
    10. Garrett R
    11. Bourtzis K
    12. Andersson SGE

    (2009) The mosaic genome structure of the wolbachia wri strain infecting Drosophila simulans

    PNAS 106:5725–5730.

    https://doi.org/10.1073/pnas.0810753106

    • PubMed
    • Google Scholar
33. 1. Landmann F
    2. Orsi GA
    3. Loppin B
    4. Sullivan W

    (2009) Wolbachia-mediated cytoplasmic incompatibility is associated with impaired histone deposition in the male pronucleus

    PLOS Pathogens 5:e1000343.

    https://doi.org/10.1371/journal.ppat.1000343

    • PubMed
    • Google Scholar
34. 1. Lassy CW
    2. Karr TL

    (1996) Cytological analysis of fertilization and early embryonic development in incompatible crosses of Drosophila simulans

    Mechanisms of Development 57:47–58.

    https://doi.org/10.1016/0925-4773(96)00527-8

    • PubMed
    • Google Scholar
35. 1. LePage DP
    2. Jernigan KK
    3. Bordenstein SR

    (2014) The relative importance of DNA methylation and dnmt2-mediated epigenetic regulation on wolbachia densities and cytoplasmic incompatibility

    PeerJ 2:e678.

    https://doi.org/10.7717/peerj.678

    • PubMed
    • Google Scholar
36. 1. LePage DP
    2. Metcalf JA
    3. Bordenstein SR
    4. On J
    5. Perlmutter JI
    6. Shropshire JD
    7. Layton EM
    8. Funkhouser-Jones LJ
    9. Beckmann JF
    10. Bordenstein SR

    (2017) Prophage WO genes recapitulate and enhance wolbachia-induced cytoplasmic incompatibility

    Nature 543:243–247.

    https://doi.org/10.1038/nature21391

    • PubMed
    • Google Scholar
37. 1. Le Thomas A
    2. Rogers AK
    3. Webster A
    4. Marinov GK
    5. Liao SE
    6. Perkins EM
    7. Hur JK
    8. Aravin AA
    9. Tóth KF

    (2013) Piwi induces pirna-guided transcriptional silencing and establishment of a repressive chromatin state

    Genes & Development 27:390–399.

    https://doi.org/10.1101/gad.209841.112

    • PubMed
    • Google Scholar
38. 1. Li XY
    2. Harrison MM
    3. Villalta JE
    4. Kaplan T
    5. Eisen MB

    (2014) Establishment of regions of genomic activity during the Drosophila maternal to zygotic transition

    eLife 3:e3737.

    https://doi.org/10.7554/eLife.03737

    • PubMed
    • Google Scholar
39. 1. McClintock B

    (1941) The stability of broken ends of chromosomes in zea mays

    Genetics 26:234–282.

    https://doi.org/10.1093/genetics/26.2.234

    • PubMed
    • Google Scholar
40. Conference

    1. Md V
    2. Misra S
    3. Li H
    4. Aluru S

    (2019) Efficient Architecture-Aware Acceleration of BWA-MEM for Multicore Systems

    IEEE.

    https://doi.org/10.1109/IPDPS.2019.00041

    • Google Scholar
41. 1. Megraw TL
    2. Li K
    3. Kao LR
    4. Kaufman TC

    (1999) The centrosomin protein is required for centrosome assembly and function during cleavage in Drosophila

    Development 126:2829–2839.

    https://doi.org/10.1242/dev.126.13.2829

    • PubMed
    • Google Scholar
42. 1. Momtaz AZ
    2. Ahumada Sabagh AD
    3. Gonzalez Amortegui JG
    4. Salazar SA
    5. Finessi A
    6. Hernandez J
    7. Christensen S
    8. Serbus LR

    (2020) A role for maternal factors in suppressing cytoplasmic incompatibility

    Frontiers in Microbiology 11:576844.

    https://doi.org/10.3389/fmicb.2020.576844

    • PubMed
    • Google Scholar
43. 1. Moretti R
    2. Yen PS
    3. Houé V
    4. Lampazzi E
    5. Desiderio A
    6. Failloux AB
    7. Calvitti M

    (2018) Combining wolbachia-induced sterility and virus protection to fight aedes albopictus-borne viruses

    PLOS Neglected Tropical Diseases 12:e0006626.

    https://doi.org/10.1371/journal.pntd.0006626

    • PubMed
    • Google Scholar
44. 1. Negri I
    2. Franchini A
    3. Gonella E
    4. Daffonchio D
    5. Mazzoglio PJ
    6. Mandrioli M
    7. Alma A

    (2009) Unravelling the wolbachia evolutionary role: the reprogramming of the host genomic imprinting

    Proceedings. Biological Sciences 276:2485–2491.

    https://doi.org/10.1098/rspb.2009.0324

    • PubMed
    • Google Scholar
45. 1. O’Neill SL
    2. Karr TL

    (1990) Bidirectional incompatibility between conspecific populations of Drosophila simulans

    Nature 348:178–180.

    https://doi.org/10.1038/348178a0

    • PubMed
    • Google Scholar
46. 1. Quinlan AR
    2. Hall IM

    (2010) BEDTools: a flexible suite of utilities for comparing genomic features

    Bioinformatics 26:841–842.

    https://doi.org/10.1093/bioinformatics/btq033

    • PubMed
    • Google Scholar
47. 1. Reed KM
    2. Werren JH

    (1995) Induction of paternal genome loss by the paternal-sex-ratio chromosome and cytoplasmic incompatibility bacteria (wolbachia): a comparative study of early embryonic events

    Molecular Reproduction and Development 40:408–418.

    https://doi.org/10.1002/mrd.1080400404

    • PubMed
    • Google Scholar
48. 1. Riggs B
    2. Rothwell W
    3. Mische S
    4. Hickson GRX
    5. Matheson J
    6. Hays TS
    7. Gould GW
    8. Sullivan W

    (2003) Actin cytoskeleton remodeling during early Drosophila furrow formation requires recycling endosomal components nuclear-fallout and rab11

    The Journal of Cell Biology 163:143–154.

    https://doi.org/10.1083/jcb.200305115

    • PubMed
    • Google Scholar
49. 1. Ryan SL
    2. Saul GB

    (1968) Post-fertilization effect of incompatibility factors in mormoniella

    Molecular & General Genetics 103:29–36.

    https://doi.org/10.1007/BF00271154

    • PubMed
    • Google Scholar
50. 1. Seller CA
    2. O’Farrell PH

    (2018) Rif1 prolongs the embryonic S phase at the Drosophila mid-blastula transition

    PLOS Biology 16:e2005687.

    https://doi.org/10.1371/journal.pbio.2005687

    • PubMed
    • Google Scholar
51. 1. Seller CA
    2. Cho CY
    3. O’Farrell PH

    (2019) Rapid embryonic cell cycles defer the establishment of heterochromatin by eggless/setdb1 in Drosophila

    Genes & Development 33:403–417.

    https://doi.org/10.1101/gad.321646.118

    • PubMed
    • Google Scholar
52. 1. Serbus LR
    2. Casper-Lindley C
    3. Landmann F
    4. Sullivan W

    (2008) The genetics and cell biology of wolbachia-host interactions

    Annual Review of Genetics 42:683–707.

    https://doi.org/10.1146/annurev.genet.41.110306.130354

    • PubMed
    • Google Scholar
53. 1. Serbus LR
    2. White PM
    3. Silva JP
    4. Rabe A
    5. Teixeira L
    6. Albertson R
    7. Sullivan W

    (2015) The impact of host diet on wolbachia titer in Drosophila

    PLOS Pathogens 11:e1004777.

    https://doi.org/10.1371/journal.ppat.1004777

    • PubMed
    • Google Scholar
54. 1. Shropshire JD
    2. Leigh B
    3. Bordenstein SR

    (2020) Symbiont-mediated cytoplasmic incompatibility: what have we learned in 50 years?

    eLife 9:e61989.

    https://doi.org/10.7554/eLife.61989

    • PubMed
    • Google Scholar
55. 1. Sienski G
    2. Dönertas D
    3. Brennecke J

    (2012) Transcriptional silencing of transposons by piwi and maelstrom and its impact on chromatin state and gene expression

    Cell 151:964–980.

    https://doi.org/10.1016/j.cell.2012.10.040

    • PubMed
    • Google Scholar
56. 1. Sokac AM
    2. Biel N
    3. De Renzis S

    (2022) Membrane-actin interactions in morphogenesis: lessons learned from Drosophila cellularization

    Seminars in Cell & Developmental Biology 1:S1084-9521(22)00101-X.

    https://doi.org/10.1016/j.semcdb.2022.03.028

    • PubMed
    • Google Scholar
57. 1. Sullivan W
    2. Minden JS
    3. Alberts BM

    (1990) Daughterless-abo-like, a Drosophila maternal-effect mutation that exhibits abnormal centrosome separation during the late blastoderm divisions

    Development 110:311–323.

    https://doi.org/10.1242/dev.110.2.311

    • PubMed
    • Google Scholar
58. 1. Sullivan W
    2. Daily DR
    3. Fogarty P
    4. Yook KJ
    5. Pimpinelli S

    (1993) Delays in anaphase initiation occur in individual nuclei of the syncytial Drosophila embryo

    Molecular Biology of the Cell 4:885–896.

    https://doi.org/10.1091/mbc.4.9.885

    • PubMed
    • Google Scholar
59. 1. Sullivan W
    2. Theurkauf WE

    (1995) The cytoskeleton and morphogenesis of the early Drosophila embryo

    Current Opinion in Cell Biology 7:18–22.

    https://doi.org/10.1016/0955-0674(95)80040-9

    • PubMed
    • Google Scholar
60. Book

    1. Sullivan W
    2. Ashburner A
    3. Hawley RS

    (2000)

    Drosophila Protocols

    Cold Spring Harbor Laboratory Press.

    • Google Scholar
61. 1. Sun G
    2. Zhang M
    3. Chen H
    4. Hochstrasser M

    (2022) The cinb nuclease from wno wolbachia is sufficient for induction of cytoplasmic incompatibility in Drosophila

    MBio 1:e0317721.

    https://doi.org/10.1101/2021.10.22.465375

    • Google Scholar
62. 1. Tang X
    2. Cao J
    3. Zhang L
    4. Huang Y
    5. Zhang Q
    6. Rong YS

    (2017) Maternal haploid, a metalloprotease enriched at the largest satellite repeat and essential for genome integrity in Drosophila embryos

    Genetics 206:1829–1839.

    https://doi.org/10.1534/genetics.117.200949

    • PubMed
    • Google Scholar
63. 1. Teixeira FK
    2. Okuniewska M
    3. Malone CD
    4. Coux RX
    5. Rio DC
    6. Lehmann R

    (2017) PiRNA-mediated regulation of transposon alternative splicing in the soma and germ line

    Nature 552:268–272.

    https://doi.org/10.1038/nature25018

    • PubMed
    • Google Scholar
64. 1. Titen SWA
    2. Golic KG

    (2008) Telomere loss provokes multiple pathways to apoptosis and produces genomic instability in Drosophila melanogaster

    Genetics 180:1821–1832.

    https://doi.org/10.1534/genetics.108.093625

    • PubMed
    • Google Scholar
65. 1. Tram U
    2. Sullivan W

    (2002) Role of delayed nuclear envelope breakdown and mitosis in wolbachia-induced cytoplasmic incompatibility

    Science 296:1124–1126.

    https://doi.org/10.1126/science.1070536

    • PubMed
    • Google Scholar
66. 1. Tram U
    2. Fredrick K
    3. Werren JH
    4. Sullivan W

    (2006) Paternal chromosome segregation during the first mitotic division determines wolbachia-induced cytoplasmic incompatibility phenotype

    Journal of Cell Science 119:3655–3663.

    https://doi.org/10.1242/jcs.03095

    • PubMed
    • Google Scholar
67. 1. Turelli M
    2. Hoffmann AA

    (1991) Rapid spread of an inherited incompatibility factor in california Drosophila

    Nature 353:440–442.

    https://doi.org/10.1038/353440a0

    • PubMed
    • Google Scholar
68. 1. Wang H
    2. Xiao Y
    3. Chen X
    4. Zhang M
    5. Sun G
    6. Wang F
    7. Wang L
    8. Zhang H
    9. Zhang X
    10. Yang X
    11. Li W
    12. Wei Y
    13. Yao D
    14. Zhang B
    15. Li J
    16. Cui W
    17. Wang F
    18. Chen C
    19. Shen W
    20. Su D
    21. Bai F
    22. Huang J
    23. Ye S
    24. Zhang L
    25. Ji X
    26. Wang W
    27. Wang Z
    28. Hochstrasser M
    29. Yang H

    (2022) Crystal structures of wolbachia cida and cidb reveal determinants of bacteria-induced cytoplasmic incompatibility and rescue

    Nature Communications 13:1608.

    https://doi.org/10.1038/s41467-022-29273-w

    • PubMed
    • Google Scholar
69. 1. Weinert LA
    2. Araujo-Jnr EV
    3. Ahmed MZ
    4. Welch JJ

    (2015) The incidence of bacterial endosymbionts in terrestrial arthropods

    Proceedings. Biological Sciences 282:20150249.

    https://doi.org/10.1098/rspb.2015.0249

    • PubMed
    • Google Scholar
70. 1. Werren JH
    2. Baldo L
    3. Clark ME

    (2008) Wolbachia: master manipulators of invertebrate biology

    Nature Reviews. Microbiology 6:741–751.

    https://doi.org/10.1038/nrmicro1969

    • PubMed
    • Google Scholar
71. Software

    1. Wickham H

    (2016) Elegant graphics for data analysis

    Ggplot2.

    https://ggplot2/tidyverse.org
72. 1. Wright JD
    2. Barr AR

    (1981) Wolbachia and the normal and incompatible eggs of aedes polynesiensis (diptera: culicidae)

    Journal of Invertebrate Pathology 38:409–418.

    https://doi.org/10.1016/0022-2011(81)90109-9

    • Google Scholar
73. 1. Wu X
    2. Lindsey ARI
    3. Chatterjee P
    4. Werren JH
    5. Stouthamer R
    6. Yi SV

    (2020) Distinct epigenomic and transcriptomic modifications associated with wolbachia-mediated asexuality

    PLOS Pathogens 16:e1008397.

    https://doi.org/10.1371/journal.ppat.1008397

    • PubMed
    • Google Scholar
74. 1. Ye YH
    2. Woolfit M
    3. Huttley GA
    4. Rancès E
    5. Caragata EP
    6. Popovici J
    7. O’Neill SL
    8. McGraw EA

    (2013) Infection with a virulent strain of wolbachia disrupts genome wide-patterns of cytosine methylation in the mosquito aedes aegypti

    PLOS ONE 8:e66482.

    https://doi.org/10.1371/journal.pone.0066482

    • PubMed
    • Google Scholar
75. 1. Yen JH
    2. Barr AR

    (1971) New hypothesis of the cause of cytoplasmic incompatibility in culex pipiens L

    Nature 232:657–658.

    https://doi.org/10.1038/232657a0

    • PubMed
    • Google Scholar
76. 1. Yuan K
    2. Seller CA
    3. Shermoen AW
    4. O’Farrell PH

    (2016) Timing the Drosophila mid-blastula transition: A cell cycle-centered view

    Trends in Genetics 32:496–507.

    https://doi.org/10.1016/j.tig.2016.05.006

    • PubMed
    • Google Scholar
77. 1. Zheng X
    2. Zhang D
    3. Li Y
    4. Yang C
    5. Wu Y
    6. Liang X
    7. Liang Y
    8. Pan X
    9. Hu L
    10. Sun Q
    11. Wang X
    12. Wei Y
    13. Zhu J
    14. Qian W
    15. Yan Z
    16. Parker AG
    17. Gilles JRL
    18. Bourtzis K
    19. Bouyer J
    20. Tang M
    21. Zheng B
    22. Yu J
    23. Liu J
    24. Zhuang J
    25. Hu Z
    26. Zhang M
    27. Gong JT
    28. Hong XY
    29. Zhang Z
    30. Lin L
    31. Liu Q
    32. Hu Z
    33. Wu Z
    34. Baton LA
    35. Hoffmann AA
    36. Xi Z

    (2019) Incompatible and sterile insect techniques combined eliminate mosquitoes

    Nature 572:56–61.

    https://doi.org/10.1038/s41586-019-1407-9

    • PubMed
    • Google Scholar

Author details

1. Brandt Warecki

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

   Contribution

   Conceptualization, Data curation, Software, Formal analysis, Supervision, Validation, Investigation, Visualization, Methodology, Writing – original draft, Writing – review and editing

   Contributed equally with

   Simon William Abraham Titen

   For correspondence

   bwarecki@ucsc.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-8025-6246

2. Simon William Abraham Titen

   1. Department of Molecular, Cell, and Developmental Biology, University of California, Santa Cruz, Santa Cruz, United States
   2. Department of Biology and Chemistry, California State University Monterey Bay, Seaside, United States

   Contribution

   Conceptualization, Formal analysis, Validation, Investigation, Methodology, Writing – review and editing

   Contributed equally with

   Brandt Warecki

   Competing interests

   No competing interests declared

3. Mohammad Shahriyar Alam

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

   Contribution

   Conceptualization, Formal analysis, Investigation, Writing – original draft, Writing – review and editing

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0003-2943-8084

4. Giovanni Vega

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

   Contribution

   Formal analysis, Validation, Investigation

   Competing interests

   No competing interests declared

5. Nassim Lemseffer

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

   Contribution

   Formal analysis, Investigation, Methodology

   Competing interests

   No competing interests declared

6. Karen Hug

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

   Contribution

   Formal analysis, Investigation

   Competing interests

   No competing interests declared

7. Jonathan S Minden

   Department of Biological Sciences, Carnegie Mellon University, Pittsburgh, United States

   Contribution

   Methodology

   Competing interests

   No competing interests declared

8. William Sullivan

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

   Contribution

   Conceptualization, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Methodology, Writing – original draft, Project administration, Writing – review and editing

   For correspondence

   wtsulliv@ucsc.edu

   Competing interests

   No competing interests declared

   "This ORCID iD identifies the author of this article:" 0000-0002-1756-4174

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