Methods and compositions using chemical or genetic control to achieve cytoplasmic incompatibility without bacterial symbionts

Abstract

This disclosure relates to use of small chemical molecules and genetically modified arthropods to control and / or reduce arthropod populations.

Description

METHODS AND COMPOSITIONS USING CHEMICAL OR GENETIC CONTROL TO ACHIEVE CYTOPLASMIC INCOMPATIBILITY WITHOUT BACTERIAL SYMBIONTS

[0001] This application claims the pnority benefit of U.S. Provisional Patent Application Senal No. 63 / 557,196, filed February 23, 2024, and U.S. Provisional Patent Application Senal No. 63 / 649,177, filed May 17, 2024, which are hereby incorporated by reference in their entirety.

[0002] This invention was made with government support under Grant Nos. AI179743 and AI143725 awarded by the National Institutes of Health. The Government has certain rights in the invention.FIELD

[0003] The present disclosure relates to methods and compositions for controlling and / or reducing arthropod populations using chemical and genetic control of arthropods by affecting histone acetylation and long noncoding RNA.SEQUENCE LISTING STATEMENT

[0004] This application contains a computer readable Sequence Listing which has been submitted electronically in XML format and is hereby incorporated by reference in its entirety. The XML file was created on February 20, 2025, is named 148411004002.xml, and is 60,737 bytes m size.BACKGROUND

[0005] Reproductive symbionts are widely distributed in diverse animal and plant hosts and often exert more influence on host biology than their symbiotic counterparts in animal guts or plant roots. In the most speciose group of animals from the phylum Arthropoda, maternally -inherited Wolbachia are an archetype of this symbiotic lifestyle. Symbiotic relationships between arthropod hosts and microorganisms have sparked global interest in the effects of microbial-encoded traits on host ecology, evolution, and biocontrol. One central goal is to pinpoint and engineer the causal pathway(s) responsible for the expression of these traits.

[0006] Endosymbionts such as the widespread genus of bacteria, Wolbachia, inhabit approximately half of the world’s arthropod species and possess the remarkable ability tomanipulate host reproduction primarily through cytoplasmic incompatibility (CI). This phenomenon, characterized by embryo mortality' in crosses between Wolbachia-infecied males and uninfected females, offers promising avenues for vector control in conjunction with Wolbachia's capacity to inhibit pathogen replication in arthropod or pest hosts. While a CI- based strategy by the World Mosquito Program has demonstrated 40-98% efficacy in mitigating vector-borne disease transmission, challenges such as the natural waning of CI effectiveness under climate change and other variables can limit their long-term impact.

[0007] New approaches are needed that, in the absence of Wolbachia, target host epigenetic factors affected by Wolbachia under natural conditions and recreate CI and / or in the presence of Wolbachia, target RNA molecules to strengthen CI. Such innovations would aim to amplify currently -ongoing efforts in arthropod vector control, and also expand their utility to any relevant arthropod vector and pest that are either naturally refractory to CI, lack Wolbachia, or Wolbachia-mduced CI strength is weak.

[0008] This disclosure is directed to overcoming these and other deficiencies in the art.SUMMARY

[0009] One aspect of the present disclosure relates to a method of population suppression of arthropods. This method involves contacting a population of male arthropods with ahistone acetyltransferase (HAT) inhibitor, where said contacting: (i) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females lacking a bacterial endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont or in comparison to uncontacted males; or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to uncontacted males; and releasing the population of male arthropods into a target population of arthropods lacking a rescuing bacterial endosymbiont under conditions effective to reduce the target population.

[0010] Another aspect of the present disclosure relates to a method of population replacement of target arthropods. This method involves contacting a population of male arthropods with a histone acetyltransferase (HAT) inhibitor, where said contacting: (i) increases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a rescuing bacterial endosymbiont in comparison to mating with arthropod females lacking a rescuing bacterial endosymbiont, or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having anon-rescuing bacterial endosymbiont in comparison to uncontacted males. The method further involves either (i) introducing the male arthropods into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, where said introducing replaces the target population lacking the rescuing endosymbiont or (ii) introducing a population of female arthropods having a rescuing bacterial endosymbiont and the contacted male arthropods into a target population of arthropods, where said introducing replaces the target population of arthropods.

[0011] A further aspect of the present disclosure relates to an arthropod control formulation comprising a histone acetyltransferase (HAT) inhibitor and a carrier comprising the HAT inhibitor, where the carrier is suitable for delivering the HAT inhibitor to a male arthropod to cause a reduction in an ability of the male arthropod to produce viable offspring with a female arthropod (i) lacking a bacterial endosymbiont or (ii) having a non-rescuing endosymbiont compared to a female arthropod having a rescuing bacterial endosymbiont.

[0012] Another aspect of the present disclosure relates to a chemically treated male arthropod, where the male arthropod has at least 10% less viable offspring when mated with a female arthropod having no endosymbiont or a non-rescuing bacterial endosymbiont in comparison to mating with a female arthropod having a rescuing bacterial endosymbiont.

[0013] Y et another aspect of the present disclosure relates to a genetically engineered arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase gene or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm development in a male arthropod causes a reduction in fertility and / or the ability of the male arthropod to produce viable offspring with arthropod females lacking a bacterial endosymbiont or having a non-rescuing endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont.

[0014] Another aspect of the present disclosure relates to a method of population suppression of arthropods. This method involves providing a genetically engineered arthropod, the arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod (i) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females lacking a bacterial endosymbiont in comparison to mating with arthropod females having a rescuingbacterial endosymbiont or in comparison to wild-type males; or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to wild-type males; and releasing the genetically engineered arthropod into a target population of arthropods under conditions effective to reduce the target population.

[0015] Another aspect of the present disclosure relates to a method of population replacement of target arthropods. This method involves providing a genetically engineered male arthropod, the male arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod (i) increases the abi 1 i ty of the male arthropod to produce viable offspring upon mating with a female arthropod having a rescuing bacterial endosymbiont in comparison to a female arthropod lacking a rescuing bacterial endosy mbiont, or (ii) decreases the ability of the male arthropod to produce viable offspring upon mating with a female arthropod having a non-rescuing bacterial endosymbiont in or in comparison to a wild-type male arthropod. This method further involves either (i) introducing the male arthropod into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, where said introducing replaces the target population lacking the rescuing endosymbiont or (ii) introducing a population of female arthropods having a rescuing bacterial endosymbiont and the male arthropods into a target population of arthropods, where said introducing replaces the target population of arthropods.

[0016] The present disclosure relates to chemically inhibiting a keystone event in animal reproductive epigenetics to synthetically replicate a paternal-effect lethality trait termed cytoplasmic incompatibility (CI) induced by the endosymbiont Wolbachia. Using a small molecule inhibitor of histone acetyltransferase, renders the arthropod sperm CI capable without Wolbachia presence. The chemical-based findings described herein were validated using transgenic knockdown of histone acetyltransferase and histone deacetylase, along with immunostaining of engineered aposymbiotic males that display reduced histone acetylation and retention of specific histones on the developing spermatids. This disclosure shifts the focus of CI from a solely organismal trait to an engineered molecular one, opening up new strategies for controlling CI, or Wolbachia-resistant arthropods as described herein.

[0017] The present disclosure also tackles the challenge of waning CI in Wolbachia infected arthropods by introducing a novel approach that, in the absence of Wolbachia, targetshost epigenetic factors affected by Wolbachia under natural conditions and recreates CL For example, in some embodiments, D. melanogaster males treated with anacardic acid, a pharmacological inhibitor, exhibit altered histone acetylation and a disrupted histone-to- protamine exchange during sperm development. Sperm of treated males develop with abnormally retained histone H3 and depleted protamines, the chromatin integrity signatures of natural CI. Consequently, molecular disentanglement of this symbiosis now enables a bypass of the symbiont to engineer a reproductive trait central to arthropod evolution and vector control. This innovation aims to amplify currently -ongoing efforts in arthropod vector control and also expands its utility in any relevant arthropod vector and pest that are either naturally refractory to CI, lack Wolbachia, or where Wolbachia-mducQd CI strength is weak.

[0018] By targeting host reproduction-epigenetic axis of molecular biology7, this innovation of chemically-engineering or programming CI in the absence of Wolbachia provides a novel approach for population suppression, replacement, and trait propagation in arthropod populations. Central to this innovation is the regulated inhibition of histone acetylation at a specific stage of spermiogenesis in arthropod males.

[0019] The present disclosure also addresses this limitation by introducing another novel approach that targets host genetic factors, specifically long non-coding RNA (“IncRNA”), to augment CI and enhance arthropod vector control efforts for population suppression or replacement. By integrating transgenes targeting IncRNA with Wolbachia infection in transgenic arthropods, the present disclosure facilitates applications including Controlled Population Suppression, Population Replacement, and Genetic Drive for Trait Propagation.

[0020] The present disclosure represents a paradigm shift in controlling commercially valuable arthropod insects and pest species. It offers comprehensive and integrated approaches that leverage both microbial and host genetic factors to recreate CI and ensure its continued efficacy in the systems devoid of Wolbachia and mitigate the risk of waning CI in systems carry ing Wolbachia. By broadening the scope of arthropod control beyond traditional methods, these innovations hold significant promise for addressing global challenges in various contexts, including disease vector control, agricultural pest management, and mitigation of invasive species in built environments.BRIEF DESCRIPTION OF THE DRAWINGS

[0021] FIGs. 1A-E illustrate that Histone H3 and H4 acetylation levels are reduced during spermatogenesis in Ci-causing w'Mel+ males. FIG. 1A is a schematic representation ofa. Drosophila melanogaster male reproductive system created with Biorender.com. In FIGs. 1B-E, testes (n = 10) from <8-hour-old males of Ci-causing wMel+ and non-CI causing wMel- males were dissected and immunostained to visualize H3 and H4 histone acetylation levels (purple) during various stages of sperm development. DAPI (grey) was used as a control stain to label spermatid DNA. FIG. IB is a series of photographs showing that in non- CI control wMel- testes, H3 acetylation (H3ac) occurred in primary spermatocytes, disappeared in post-meiotic round onion spermatids, reappeared at a low level in young elongating stage, followed by a slight increase at early canoe nuclei. H3ac is almost undetectable at late canoe nuclei, in correlation with the stage when histones are degraded from the chromatin. On the contrary, wMel+ CI testes showed remarkably reduced H3ac levels at all stages of spermiogenesis. FIG. 1C is a series of photographs showing H4 histone acetylation (H4ac) occurred at higher intensity than H3ac, and the signal intensity levels were similar in both wMel- and wMel+ males across all stages of sperm development (see FIG. 6). FIG. ID is a graph of H3ac signal intensity per stage nuclei that was quantified in ImageJ and graphed. FIG. IE is a graph of total spermatid bundles positive with DAPI and H4ac signals that were manually counted and graphed. Compared to wMel- control, wMel+ testes showed significantly reduced bundle numbers with H4ac signals at early canoe stage (see FIG.7). Horizontal lines in FIG. ID and FIG. IE represent the median. Asterisks indicate statistically significant (p < 0.05) differences as determined by pairwise Mann-Whitney tests. All p-values are reported in Table SI in Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety. Scale bars are 10 pm in FIGs. IB and 1C.

[0022] FIGs. 2A-B show a schematic illustration and a graph demonstrating that aposymbiotic wMel- males treated with a small molecule inhibitor recapitulates CL FIG. 2A shows a schematic representation of fly rearing and chemical feeding setup to generate males used for hatch rate assay created with Biorender.com. 1st and 2nd instar larvae (LI and L2) from uninfected wMel- line were fed 150 pM Anacardic acid (AA) dissolved in DMSO solvent in 10 ml standard food vials. DMSO-mixed food alone was used as a negative control group. CI hatch rate analysis was conducted using freshly hatched <4 hours old, AA- and DMSO-treated males mated to 6 days old wMel- and wMel+ females to measure CI and rescue, respectively. In parallel, wMel+ larvae were fed on standard food to generate wild type Ci-causing males as controls. In FIG. 2B, each circle data point represents percent of embryosthat hatched into larvae from a single male and female pair. Sample size is listed in parentheses. AA-treated wMel- (AA wMel-) males recapitulated rescuable CI similar to wMel- induced wild type CI. DMSO-treated wMel- (DMSO t Mel -) males showed higher embryonic hatching when crossed with both wMel- and wMel+ females indicative of no CI. Some variation in embryonic hatch rates occurred in both AA and DMSO crosses, presumably due to off-target impacts of DMSO. Horizontal bars represent the median. Asterisks indicate significant differences determined by Mann-Whitney pairwise comparison test and Kruskal- Wallis followed by Dunn’s multiple comparison tests. All p-values are reported in Table SI in Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383:1111-1117 (2024), which is hereby incorporated by reference in its entirety.

[0023] FIGs. 3A-D illustrate that Histone H3 and H4 acetylation levels are reduced in aposymbiotic CI males. Sibling males from the same setup in FIGs. 2A-B were used to dissect testes (n = 10) and immunostained to visualize H3ac and H4ac levels (purple) during various stages of sperm development. DAPI (grey) was used as a control stain to label spermatid DNA. FIGs. 3A-B are photographs demonstrating that AA-treated males showed significantly reduced H3ac and H4ac levels in all post-meiotic development stages compared to DMSO- treated controls. Both H3ac and H4ac levels did not differ in spermatocytes of AA- and DMSO-treated testes possibly because larvae had already progressed to the spermatocyte development, thus minimizing acid impact at this stage. Both signals disappear at late canoe nuclei stage in correlation with histone degradation. FIGs. 3C-D are graphs showing H3ac and H4ac signal intensity per stage nuclei that was quantified in ImageJ and graphed. Horizontal lines in FIG. 3C and FIG. 3D represent the median. Asterisks indicate statistically significant (p < 0.05) differences as determined by pairwise Mann-Whitney tests. All p-values are reported in Table SI in Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety.

[0024] FIGs. 4A-C illustrate that aposymbiotic CI spermatids abnormally retain histone H3 at the late stage of spermiogenesis. Sibling males from the same setup used in FIGs. 2A-B were used to dissect testes (n = 10) and immunostained to visualize H3 histone (yellow) localization from early to late sperm development stages. DAPI (grey) was used as a control stain to label spermatid DNA. Scale bars are 10 pm. FIG. 4A is a series of photographs showing that in non-CI control DMSO wMel- testes, H3 signal is detected from round onionto early canoe stage and disappear, as expected, during transition from early-to-late canoe followed by needle stage nuclei. On the contrary, Ci-causing AA wMel- males have abnormally retained H3 histone from late canoe to needle stage where nuclei become fully condensed and mature. FIG. 4B is a graph of total late-stage spermatid bundles with retained H3 histones that were manually counted and graphed. FIG. 4C is a graph showing that mature sperm isolated from seminal vesicles of AA wMel- males show enhanced protamine deficiency levels compared to DMSO wMel- control. Mature sperm isolated from seminal vesicles (n = 10) of <4 hour-old sibling males were stained with fluorescent chromomycin A3 (CMA3) stain to detect protamine deficiency (corresponding to high green fluorescence intensity of CMA3) in each sperm nucleus. Individual sperm head intensity was quantified in ImageJ and graphed. AA wMel- sperm show enhanced protamine deficiency levels compared to DMSO wMel- control. All p-values are reported in Table SI in Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal- effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety. Horizontal lines in FIGs. 4B-C represent the median. Asterisks indicate statistically significant (p < 0.05) differences as determined by pairwise Mann-Whitney U tests. The wild type wMel+ males showed similar H3 histone retention compared to wMel- males (FIGs. 5A-B).

[0025] FIGs. 5A-B illustrate that Histone H3 are retained in late spermiogenesis stages in wild type Ci-causing wMel+ males. Testes (n = 10) from <8-hour-old males of Ci-causing wMel- and non-CI causing wMel- males were dissected and immunostained to visualize H3 histones (yellow) from early to late sperm development stages. FIG. 5A is a series of photographs of wild type wMel+ CI males that show abnormally retained H3 histone from late canoe to needle stage where nuclei become fully condensed and mature compared to non-CI wMel- control. FIG. 5B is a graph of total late-stage spermatid bundles with retained H3 histones that were manually counted and graphed. Scale bars are 10 pm.

[0026] FIG. 6 is a graph showing that Histone H4 acetylation intensity levels are similar in wMel+ CI and wMel- non-CI males. Testes (n = 10) from <8-hour-old males of Cleansing wMel+ and non-CI causing wMel- males were dissected and immunostained to measure H4 histone acetylation (H4ac) levels during various stages of sperm development. H4ac signal intensity per stage nuclei as shown in FIGs. 1 A-E was quantified in ImageJ and graphed. Horizontal lines represent the median. Statistical significance (p < 0.05) was determined by pairwise Mann- Whitney test across stages.

[0027] FIG. 7 are photographs showing that Ci-causing wMel+ males have reduced number of spermatid bundles with H4ac. Total spermatid bundles positive with DAPI (grey) and H4ac (purple) signals as imaged in FIGs. 1B-E were manually counted and graphed. Compared to wMel- control, H4ac signals at early canoe stage were absent in wMel+ spermatid bundles positive with DAPI signals.

[0028] FIGs. 8A-B are photographs showing that in both Ci-causing AA wMel- and wild type wMel+ testes, H2B histones are degraded at late canoe stage. Sibling males from setup as in FIG. 2A were used to dissect testes (n = 10) and visualize H2B histone (red) localization from early to late sperm development stages. DAPI (grey) was used as a control stain to label spermatid DNA (FIGs. 8A-B). In both Ci-causing AA wMel- and wMel+ testes and non-CI control DMSO wMel- and wild type wMel- testes, H2B signals occurred from spermatocytes to early canoe stage and disappear, as expected, during late canoe stage.

[0029] FIGs. 9A-B are graphs showing transgenic knockdown of Hat 1 and HDAC1 enhances and suppresses wild-type CI, respectively. CI hatch rate analysis was conducted using freshly hatched <8 hours old males with RNAi-mediated knocked down expression of (FIG. 9A) Histone acetyltransferase (Hail) and (FIG. 9B) Histone deacetylase (HDACl') mated to 6 days old wMel- and wMel+ females to measure CI and rescue, respectively. To drive RNAi expression, the Vasa-Gal4 driver that is primarily active in the early stages of spermatogenesis, as germ cells differentiate into spermatocytes (Demarco et al., “Investigating Spermatogenesis n Drosophila melanogaster,” Methods 68:218-227 (2014), which is hereby incorporated by reference in its entirety) was utilized. To control for any background confounding effects of Vasa-Gal4 driver line, wild type wMel+ males were prior crossed to Gal4 females infected with Wolbachia to generate fathers with Vasa;wMel+ genotype to use as wild type CI control. In FIG. 9A, males were generated using paternal grandmothers aged 5 days old that induce incomplete / intermediate levels of wild type CI (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-induced Cytoplasmic Incompatibility in Drosophila melanogaster ” mBio 10(6): eO 1879- 19 (2019), which is hereby incorporated by reference in its entirety). In FIG. 9B, males were generated using paternal grandmothers aged 11-13 days old that typically induce very strong levels of wild type CI (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-in eA. Cytoplasmic Incompatibility in Drosophila melanogaster ” mBio 10(6): eO 1879- 19 (2019), which is hereby incorporated by reference in its entirety). Each circle data point represents the percent of embryos that hatched into larvae from a single male and female pair. Sample size is listed in parentheses. Horizontallines represent the median. Asterisks indicate statistically significant (p < 0.05) differences determined by Mann-Whitney pairwise comparison test and Kruskal-Wallis followed by Dunn’s multiple comparison tests.

[0030] FIG. 10 is a graph showing the Hatch rate assay examining median CI levels in Drosophila simulans wRi-infected males (wRi+) hatched from standard food diet, and uninfected males (wRiT-) hatched from food treated with 150 pM anacardic acid- (AA) and dimethylsulfoxide (DMSO). T indicates ‘tetracycline’ treatment to create Wolbachia- uninfected line. Crosses are listed as Male x Female.

[0031] FIGs. 11 A-B are graphs of Drosophila simulans survivorship from the larval to adult stage plotted by sex (FIG. 11 A) and total individuals hatched (FIG. 1 IB) when larvae were fed standard food treated with 150 pM anacardic acid (AA) and dimethylsulfoxide (DMSO).

[0032] FIG. 12 is a graph of a hatch rate assay examining median CI levels in infected (wRi+) and uninfected (wRi-) Drosophila simulans males hatched from food treated with 37.5 pM, 75 pM, and 112.5 pM of anacardic acid- (AA) and dimethylsulfoxide (DMSO).

[0033] FIGs. 13 A-B are graphs of Drosophila simulans survivorship from the larval to adult stage plotted by sex (FIG. 13A) and total individuals (FIG. 13B) when larvae were fed standard food treated with 37.5 pM, 75 pM, and 112.5 pM anacardic acid (AA) and dimethylsulfoxide (DMSO).

[0034] FIGs. 14A-B illustrate that CifA is an in vitro DNase and RNase, and CifB is a DNase. FIG. 14A is a schematic representation showing the Cif recombinant proteins used in the study. Domain annotations are based on structural homology-based analyses (Lindsey et al., “Evolutionary Genetics of Cytoplasmic Incompatibility Genes cifA and cifB in Prophage WO of W lbachia ” Genome Biol. Evol. 10:434-451 (2018), which is hereby incorporated by reference in its entirety). The full His-tagged CifA protein was generated with (CifApy) and without (CifA) substitutions in the QxxxY motif (SEQ ID NO: 18). Since the full length CifB protein is too large to be recombinantly expressed in E. coli, purified His-GST tagged CifB variants were generated with the amino terminus, N-terminal nuclease domain (NTND), and C-terminal nuclease domain (CTND) referred to CifBAD, along with engineered substitutions in the QxxxY motif (CifBQY). Shown are CifB NTND and CTND sequences PDDEXK (SEQ ID NO:20). FIG. 14B is a compilation of blots showing nuclease activities of the Cif proteins against single-stranded (ss)DNA, double-stranded (ds)DNA and ssRNA substrates. CifBAD cleaves both ss- and ds-DNA and not RNA. QxxxY motif substitutions ablate CifB DNaseactivity. CifA is both a DNase and RNase. The CifAQY mutant ablates ssDNase activity, whereas dsDNase and ssRNase activities remain intact. Commercial DNase and RNase enzymes were used as positive controls, and no CifA or CifB proteins were added to the reaction mixtures for negative controls. EDTA was added to a 20-fold molar excess over Mg2+ to inhibit the reactions. Samples were run in a 10% polyacrylamide / TBE gel.

[0035] FIGs. 15A-D show that CifA in situ RNase activity depletes AAGAG long non-coding RNA in association with CI. Fluorescent in situ hybridization (FISH) assay was performed on testes (n=l 5) squashes from <8 hrs old males to visualize and quantify the AAGAG repeat long noncoding (Inc) RNA abundance in primary spermatocytes (Mills et al., “RNA From a Simple-tandem Repeat is Required for Sperm Maturation and Male Fertility in Drosophila melanogasterf Elife 8:e48940 (2019), which is hereby incorporated by reference in its entirety). FIG. 15A is a compilation of photographs showing that compared to wMel- negative and transgenic non-CI control line CifAAbnlsB, AAGAG RNA (red) is less abundant in primary spermatocytes marked by DAPI staining (blue) than in the wMel+ and dual- transgenic CifAB lines, respectively. FIG. 15B shows graphs of RNA signal intensity per spermatocyte that was quantified in ImageJ as described in FIG. 21 and graphed. The Y-axis corresponds to the genotype labels shown on the left side of FIG. 15 A. Vertical bars represent the median. FIG. 15C is a graph of a hatch rate assay that confirmed AAGAG_KD(+) symbiotic males with depleted IncRNA result in stronger CI than negative control Scramble_KD(+) males. Each dot represents the percentage of embryos that hatched from a single male and female pair. Asterisks indicate statistically significant differences (p <0.05) calculated by Mann- Whitney pairwise comparison test. FIG. 15D are a photograph and graph showing representative RNA-FISH and quantification data are shown for AAGAG_KD(+) and Scramble_KD(+) sibling males to those used in the panel FIG. 15C hatch rate assay.

[0036] FIGs. 16A-C illustrate that CifAB in situ DNase activity enhances DNA damage in elongating spermatids. TUNEL staining on testes (n=15) squashes from <8 hrs old males was performed to visualize and quantify sperm bundles with DNA breaks. FIG. 16A is a compilation of representative photographic images of spermatid bundles with DNA damage from each treatment group are shown. Images were taken using Keyence All-in-One confocal microscope at lOOx magnification. Compared to wMel- control (empty arrowheads), DNA break signals marked by TUNEL (green) are highly intense in the wild type wMel+ and dual transgenic CifAB lines at the canoe stage of spermiogenesis (solid arrowheads). DAPI (blue) labels spermatid nuclei. Upon individual expression, CifB induces higher DNA damagecompared to CifA. FIG. 16B shows graphs of TUNEL signal intensity from spermatid bundles that was quantified in ImageJ and graphed. Vertical bars represent the median. FIG. 16C shows graphs of total sperm bundles and those with fragmented DNA that were manually counted. The numbers of testes investigated are shown in parentheses next to the genotype. Uncropped images of testes from CI versus non-CI males with total damaged bundles with high versus low TUNEL intensity' is shown in FIGs. 23A-B. P-value significance (p < 0.05) was calculated by Mann-Whitney pairwise comparison test. Skull icon represents CI and fly icon represents non-CI lines.

[0037] FIGs. 17A-D show that late-stage CI embryos suffer from DNA damage. FIGs. 17A-C show representative images of 2-3 hours old CI embryos (wMel+ males x wMel- females) during cellular-blastoderm formation (nuclear cycles 12-14) exhibit intense pH2Av signals (red) indicative of DNA damage compared to rescue embryos (wMel+ males x wMel+ females). pH2Av is a histone H2Av variant that phospohorylates in response to DNA damage (Harumoto et al., “Male-killing Symbiont Damages Host’s Dosage-compensated Sex Chromosome to Induce Embryonic Apoptosis,” Nat. Commun. 7: 12781 (2016) and Perlmutter et al., “The Phage Gene wmk is a Candidate for Male Killing by a Bacterial Endosymbiont,” PLoS Pathog. 15(9):el007936 (2019), each of which is hereby incorporated by reference in its entirety). CI embryonic nuclei on polar ends undergo shrinkage and distantly spaced with condensed chromatin (white arrows), likely becoming apoptotic leading to embryonic death. DAPI (grey) labels embry onic nuclei. FIG. 17B shows that in a section of CI embryos, the CI- defining chromatin bridging defect is marked by an asterisk colocalized with DNA damage signal (red). In the same embryo, the nuclei in the middle section (empty arrowhead) developed normally with no pH2Av signals. Full embryo confocal images were acquired at 63x magnification using tile scan feature and automated stitching. A’, B’, C’ are magnified images of boxed regions in FIGs. 17A-C, respectively. FIG. 17D is a graph of quantification of pH2Av signal intensity per embryo derived from CI (skull icon) and non-CI / rescue (fly icon) crosses. Values in parentheses correspond to number of embryos scored. Horizontal bars represent the median. P-values < 0.05 were significant.

[0038] FIGs. 18A-E show that CI induction and ablation directly link with AAGAG IncRNA depletion, impairment of histone-to-protamine transition, and DNA damage in embryos. (FIG. 18A) RNA-FISH on testes (n=15) from <8 hour old males of Ci-inducing CifA4B and non-CI CifA3B mutant males (as described in Fig. 2) shows CifA4B mutant depletes AAGAG IncRNA, whereas CifA3B does not. (FIG. 18B) CI embryos (nos;cifA4Bmales x wMel- females) exhibit intense pH2Av signals (red) at the polar end (white arrow) indicative of DNA damage compared to non-CI embryos (nos; cifA3Bmaies x wMel- females). Dotted periphery is drawn around the embryo shape. (FIG. 18C) CI and non-CI testes were immunostained to quantify histone-retaining spermatid bundles (purple) during late canoe stage of spermiogenesis (Kaur et al., “The Cif Proteins from Wolbachici Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). Non-CI lines (CifA3B and CifAAbnlsB) showed significantly less spermatid bundles with retained histones compared to CI groups (CifA4B and CifAB). (FIG. 18D) Mature sperm from seminal vesicles were stained with fluorescent CMA3 stain (green) to detect protamine deficiency (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). CI sperm (CifA4B and CifAB) lack protamines compared to non-CI sperm (CifA3B and vu Mel-). (FIG. 18E) pH2Av signal intensity quantification per embryo derived from CI (skull icon) and non-CI (fly icon) crosses. Vertical bars represent mean, and error bars represent standard deviation. P-values < 0.05 were significant.

[0039] FIGs. 19A-B illustrate that CifA and CifB are in vitro nucleases. FIG. 19A is a blot of dilution-based nuclease assay shows the activity of CifA DNase and RNase and CifB DNase persist at higher protein concentrations and diminish modestly with dilutions, as expected. FIG. 19B shows blots of time-course assay shows nuclease activity of Cifs diminishes modestly at shorter incubation rime points, as expected.

[0040] FIGs. 20A-B illustrate nuclease activity of phylogenetic Type 1 and Type 4 CifB associates with a QxxxY motif. FfG. 20A is a schematic of phylogenetic Type 1 (Tl) and T4 CifB proteins (Barr, J. J., “Missing a Phage: Unraveling Tripartite Symbioses within the Human Gut,” mSystems 4(3):e00105-19 (2019) and Chu et al., “Genomic Maps of Long Noncoding RNA Occupancy Reveal Principles of RNA-Chromatin Interactions,” Mol. Cell 44:667-678 (2011), each of which is hereby incorporated by reference in its entirety) and amino acid sequence showing the CTND location of QxxxY motifs that belong to the HsdR subunit of RecB nuclease family (Wen et al., “Critical Roles of Long Noncoding RNAs in Drosophila Spermatogenesis,” Genome Res. 26: 1233-1244 (2016), which is hereby incorporated by reference in its entirety). The motif and glutamine (Q) and tyrosine (Y) residues are highlighted in magenta, whereas the previously characterized aspartate (D),glutamate (E)and lysine (K) residues belonging to the PDDEXK nuclease family (Caputo et al., “A Bacterium Against the Tiger: Preliminary Evidence of Fertility Reduction After Release of Aedes albopictus Males with Manipulated Wolbachia Infection in an Italian Urban Area,” PestManag. Sci. 76: 1324-1332 (2020), which is hereby incorporated by reference in its entirety) are highlighted in grey. The QxxxY motif is present in wPip Type 4 (T4) CifB known as an in vitro nuclease (Caputo et al., “A Bacterium Against the Tiger: Preliminary Evidence of Fertility Reduction After Release of Aedes albopictus Males with Manipulated Wolbachia Infection in an Italian Urban A Q^." Pest Manag. Sci. 76: 1324-1332 (2020), which is hereby incorporated by reference in its entirety) and T1 CifB from vt’Mel Wolbachia characterized as an in vitro nuclease (this disclosure). The QxxxY motif is absent in the wPip T1 CifB that was previously characterized as a non-nuclease via artificial introduction of D, E, and K residues (T1 CifB*). Shown are CifB NTND sequence PDDEXK (SEQ ID NO:20), CTND sequences PDDEXK (SEQ ID NO:20) and PDDEXK / H (SEQ ID NO:21), partial sequence ofT4wPip (SEQ ID NOs:22-23), partial sequence of Tl*wPip (SEQ ID NOs:24-25) partial sequence of TlwMel (SEQ ID NOs:26-27). FIG. 20B is a schematic of T1 CifA protein from wMel Wolbachia shown to be an in vitro nuclease (this disclosure) also contains a QxxxY motif in a predicted a-helical region. Shown is a partial sequence of T1 CifA (wMel) (SEQ ID NOs:28-29). Predicted a-helical residues are labeled “H” and residues predicted to be part of P-sheets are labeled “E.” The numbers of excluded residues are shown in parentheses. The last residue numbers are shown at the end of each sequence.

[0041] FIG. 21 is a series of photographs showing quantification of RNA signal intensity per spermatocyte. Fluorescent in situ hybridization (FISH) assay was performed on testes squashes from <8 hrs old males to visualize and quantify the abundance of an AAGAG repeat long noncoding (Inc)RNA in primary spermatocytes (Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility in Drosophila simulans ” Genetics 126(4): 933-48 (1990), which is hereby incorporated by reference in its entirety).Representative image of primary spermatocytes from wMel- testes is shown. Images were taken using Zeiss LSM 880 confocal microscope at 63x magnification. Scale bar is 50 pm. Each individual spermatocyte per testes was quantified during the image processing. Location of five representative spermatocytes are shown with circled white periphery in individual channels. The circled regions in the DAPI and AAGAG channels were used to quantify the area and raw integrated density (RawIntDen), respectively. RawIntDens is the total signal intensity of all the pixels per channel that measured RNA transcript abundance with thatparticular area. Corrected total cell fluorescence (CTCF) was calculated as previously described (Ross & Hoffmann, “Continued Susceptibility of the wMEL Wolbachia Infection in Aedes aegypti to Heat Stress Following Field Deployment and Selection,” Insects 9(3):78 (2018), which is hereby incorporated by reference in its entirety). The average signal intensity from spermatocytes per testes was plotted on the graphs shown in FIGs. 15A-D.

[0042] FIG. 22 is a graph showing aposymbiotic males with depleted AAGAG long non-coding RNA (AAGAG_KD(-) and Scramble KD(-) males do not cause CI. Hatch rate assay was conducted to test if AAGAG IncRNA depletion could induce CI in the absence of Wolbachia (vt Mel ). To control for any background confounding effects of nos-Gal4: VP 16 driver line, Ci-causing wMel+ paternal grandmothers were prior crossed to nos- males to generate fathers with nos;wMel+ genotype. Each dot represents the percent of embryos that hatched from a single male and female pair. Vertical lines represent the median. Letter to the right indicate statistically significant (p <0.05) differences calculated by Kruskal-Wallis and Dunn’s test for multiple comparisons between all groups.

[0043] FIGs. 23A-B show photographs of CifAB expression enhances DNA damage in late elongating spermatids. TUNEL staining on testes squashes from <8 hrs old males was performed to visualize sperm bundles with DNA breaks across Ci-inducing nos;cifAB and non-CI inducing nospvMel- genotypes. Images were taken at 40x magnification. FIG. 23A shows images of representative testes where elongating spermatid bundles at different stages are shown, EC - early canoe stage, LC - late canoe stage, N - needle stage. DNA breaks marked by TUNEL staining (green) are detectable only at a low level in EC spermatids (solid white arrowhead), and at higher level in LC spermatids empty arrowhead) (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster ” Syst Biol. Reprod. Med. 56:44-61 (2010), which is hereby incorporated by reference in its entirety). DAPI stain (blue) was used to label spermatid nuclei. Antibody targeting histone H4 acetylation (AcH4) was used to discriminate EC and LC stage nuclei as it disappears in LC spermatids when protamines are deposited (Grunstein, M., “Histone Acetylation in Chromatin Structure and Transcription,” Nature 389:349-352 (1997), which is hereby incorporated by reference tn its entirety). Compared to the negative control / ?os;wMel-, DNA break signals at the LC stage of spermiogenesis (empty arrow heads) are highly intense in nos; cifAB transgenic males. DNA breaks are usually undetectable at the needle stage and in nuclei upon individualization (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceedfrom a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster f Syst. Biol. Reprod. Med. 56:44-61 (2010), which is hereby incorporated by reference in its entirety), however, there is occasional DNA damage in 2 out of 15 nos;cifAB testes examined. FIG. 23B shows a photograph with a zoomed in view of individualized and mature sperm nuclei (white arrowheads) where TUNEL signals are absent likely due to a state-specific, steric hinderance for the terminal nucleotidyl transferase enzyme to access the highly condensed sperm nucleus (Bordenstein et al.,-induced IncompatibilityPrecedes Other Hybrid Incompatibilities in Nasonia,” Nature 409:707-710 (2001), which is hereby incorporated by reference in its entirety). The experiment was performed under the same setup as shown in FIGs. 16A-C.

[0044] FIG. 24A-F are graphs of single and dual expression of cifA and cifB mutants ablates in situ nuclease activity and CI phenotype. FIGs. 24A, B, D, and E show graphs of TUNEL assays on testes squashes of <8 hrs old males was performed to quantify sperm bundles with DNA breaks across the genotype treatment groups. Single and dual expression of cifA and cifB mutants in transgenic lines fails to induce spermatid DNA fragmentation. Total sperm bundles and those with fragmented DNA were manually counted from the images acquired. The numbers of testes investigated are shown in parentheses next to the genotype. The experiment was performed in two independent biological replicates in the same setup as experiments in FIGs. 16A-C. P-value significance was calculated by Kruskal -Wallis and Dunn’s test for multiple comparisons between all groups. FIGs. 24C and F are graphs of hatch rate assays that were conducted to test if single and dual expression of cifA and cifB mutants can induce CI when transgenically expressed in aposymbiotic males (unfilled sex symbols). Each dot represents the percent of embryos that hatched from a single male and female pair. Letters to the right indicate significant differences based on p = 0.05 calculated by Kruskal- Wallis and Dunn’s test for multiple comparisons between all groups. The experiment was conducted twice in parallel to the TUNEL assay.

[0045] FIG. 25 is a compilation of photographs showing that CifA and CifB signals occur with damaged DNA spermatids at the late canoe and needle stage spermatids. TUNEL (green) and CifA and CifB antibody staining (red) was performed together on the testes squashes from <8 hrs old nos;cifAB males. DAPI stain (blue) was used to label spermatid nuclei. At canoe stage spermatids, TUNEL-based DNA damage signals co-occur with Cif signals at the tip of spermatid head. At the needle stage, CifB is undetectable due to poor antibody penetration in highly condensed nuclei (Hundertmark et al., “Nejire / dCBP-mediatedHistone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster,” PLoS One 13(9):e0203622 (2018), which is hereby incorporated by reference in its entirety), however DNA damage occurs. The experiment was performed in parallel to the same setup shown in FIG. 22.

[0046] FIG. 26 is a compilation of photographs showing DNA damage signals are undetectable in early-stage CI embryos. Representative images of 0-1 hours old embryos derived from cytoplasmic incompatibility (CI) cross (wMel+ males x wMel- females) and rescue cross (wMel+ males x wMel+ females). DAPI (in grey) was used to label embryonic nuclei. pH2Av signals (in red) indicative of DNA damage are absent in these embryos. Scale bars are 50 pm. Bottom right inlets represent magnified images of dotted boxed regions around embryonic nuclei. Skull icon represents CI cross and fly icon represents rescue cross.

[0047] FIG. 27 is a schematic summary of Host Modification Model of cytoplasmic incompatibility (CI). CifA and CifB proteins invade primary spermatocyte nuclei (Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster "' PLoS One 13(9):e0203622 (2018), which is hereby incorporated by reference in its entirety). Pre-fertilization, CifA in primary spermatocytes depletes long non-coding RNA (IncRNA). Upon elongation, CifA and CifB enhance DNA damage in late-canoe spermatids. Cif-mediated RNA and DNA modulations in early spermatogenesis lead to downstream defects in histones and protamine abundance during late spermiogenesis. Resultantly, CI sperm develops with impaired sperm chromatin integrity. Post-fertilization, when CI sperm fertilizes an aposymbiotic embryo not harboring Wolbachia, the paternal chromatin with imbalanced protamine content experience chromatin remodeling defects including delayed maternal histone deposition (Breeuwer & Werren, “Microorganisms Associated with Chromosome Destruction and Reproductive Isolation Between Two Insect Species." Nature 346(6284): 558-60 (1990), which is hereby incorporated by reference in its entirety), replication stress, DNA damage, improper condensation, and delayed entry into first mitosis leading to embry onic lethality. Using CI and non-CI causing Cif mutants, IncRNA depletion, spermatid DNA damage, and abnormal histones-to-protamine transition are linked to embryonic DNA damage associated with CI. Upon fertilizing a symbiotic embryo derived from Wolbachia-canying female, it is proposed that CifA modifies maternal chromatin integrity in a manner that enables paternal modifications to restore their synchronous entry into the first mitosis under the mistiming model (Jaenike et al., “Asymmetrical Reinforcement and Wolbachia Infection in Drosophila,"PLoS Biol. 4: 1852-1862 (2006), which is hereby incorporated by reference in its entirety). The unmodified sperm from aposymbiotic males will be compatible with embryos derived from either symbiote or aposymbiotic females because maternal chromatin determines the entry timepoint into mitosis (Shoemaker et al., “Wolbachia and the Evolution of Reproductive Isolation Between Drosophila recens and Drosophila subquinaria, ” Evolution (N Y) 53: 1157— 1164 (1999), which is hereby incorporated by reference in its entirety).

[0048] FIGs. 28A-B show that mutational changes in CifA and CifB impact nuclease activity. FIG. 28A is a schematic representation showing the Cif recombinant proteins used in the study. Since the full CifB protein is too large to be recombinantly expressed in E. coli, purified His-GST tagged CifB variants were generated with the amino terminus, N-terminal nuclease domain (NTND), and C-terminal nuclease domain (CTND) along with various engineered substitutions in conserved residues. The full His-tagged CifA protein was generated with and without substitutions in conserved residues and a truncation in the C- terminus. Shown are CifB NTND and CTND sequences PDDEXK (SEQ ID NO:20). FIG. 28B are blots of nuclease activities of the Cif proteins against DNA and RNA substrates. CifB shows both ss- and ds-DNase activity. While ss-DNase activity is ablated by mutations in both NTND (CifB2;AD) and CTND (CifB3;AD), ds-DNase activity is ablated by substitutions in NTND but not in CTND. CifA is both a DNase and RNase. Substitutions in conserved residues of the STE domain (CifA4) ablate ssDNase activity but retain dsDNase and RNase function. Substitutions residues in DUF domain (CifA3) do not alter DNase and RNase action of CifA. 1 pM of Cif protein was incubated with 500 nM Cy5-labeled ssDNA, 15 nM dsDNA and 100 nM ssRNA for 120 min. DNase and RNase enzymes were used as positive controls, and no CifA or CifB proteins were added to the reaction mixtures for negative controls. EDTA was added to a 20x molar excess over Mg2+ to inhibit the reactions. Samples were run in 10%polyacrylamide / TBE gel. Assays were replicated three times independently. The dsDNA panel belonging to CifB variants was manually cropped to bring the proteins in the same loading order. Related full uncropped gel images are submitted in the raw data file S3 in Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety.

[0049] FIG. 29 shows that CifB amino terminus alone does not induce DNase or RNase activity. FIG. 29 is a schematic representation showing the location of truncated domains of CifB recombinant proteins used in the study. CifBANACAD corresponds to CifBmutant expressing A-terminus alone with truncated nuclease and DUB domains. 1 pM of Cif protein was incubated with 500 nM Cy5-labeled ssDNA. 15 nM dsDNA and 100 nM ssRNA for 120 mm. DNase and RNase enzymes were used as positive controls. To stop the reactions, EDTA was added to a 20x molar excess over Mg2+. Cy5-labeled ssDNA samples were run in a 10% polyacrylamide / TBE gel. Non-labeled dsDNA and RNA samples were run in 10% polyacrylamide / TBE gel and stained with GelRed.DETAILED DESCRIPTION

[0050] Disclosed herein are methods and compositions useful for controlling and / or reducing populations of arthropods (for example, insects) using small chemical treatment and / or genetically modified arthropods.

[0051] Unless defined otherwise, all technical and scientific terms used herein have the same meaning as commonly understood to one of ordinary skill in the art to which this disclosure belongs. The following definitions are provided for the full understanding of terms used in this specification.

[0052] The terms “promoter” and “driver” are used interchangeably herein and refer to a nucleic acid fragment capable of controlling transcription of another nucleic acid fragment. A promoter is a non-coding genomic DNA sequence, usually upstream (5') to and operably linked to the relevant coding sequence, and its primary function is to act as a binding site for RNA polymerase to initiate transcription by the RNA polymerase. A promoter may also include distal enhancer or repressor elements, which can be located as much as several thousand base pairs from the start site of transcription. The terms “capable of controlling expression” or “initiating transcription”, refer to the primary function of a promoter. Additionally, there is “expression” of RNA, including functional RNA, or the expression of a polypeptide for operably linked encoding nucleotide sequences, as the transcribed RNA ultimately may be translated into the corresponding polypeptide. Promoters vary in their “strength” (i.e., their ability to promote transcription). The nucleotide sequence of the promoter determines the nature of the RNA polymerase binding and other related protein factors that attach to the RNA polymerase and / or promoter, and the rate of RNA synthesis. A “constitutive” promoter is a promoter that is active under most environmental and developmental conditions. A “developmental stage-specific” promoter is a promoter that is active during specific stages of development.

[0053] In some embodiments, a heterologous nucleic acid molecule is introduced into an organism. The nucleic acid molecule may be a plasmid that replicates autonomously, or it may integrate into the genome of the host organism. Host organisms containing the transformed nucleic acid molecule may be referred to as “transgenic” or “recombinant” or “transformed” organisms. A “genetically modified” organism (e.g, a genetically modified arthropod) is an organism that includes a nucleic acid that has been modified by human intervention. Examples of a nucleic acid that has been modified by human intervention include, but are not limited to, insertions, deletions, mutations, expression of nucleic acid constructs (e.g. , over-expression or expression from a non-natural promoter or control sequence or an operably linked promoter and gene nucleic acid distinct from a naturally occurring promoter and gene nucleic acid in an organism), extra-chromosomal nucleic acids, and genomically contained modified nucleic acids.

[0054] As used herein, “sequence identity” means that sequences (e.g, nucleotide sequences or protein sequences) are identical (i.e., on a nucleotide-by -nucleotide basis for nucleic acids or amino acid-by -amino acid basis for polypeptides) over a window of comparison. Methods of identifying nucleotide and polypeptide sequences with sequence identity to a sequence of interest are well known in the art. For example, sequence alignments and percent identity calculations may be determined using a variety of comparison methods designed to detect homologous sequences including, but not limited to, the Multalin program (Corpet, “Multiple Sequence Alignment with Hierarchical Clustering,” Nucleic Acids Res. 16: 10881-90 (1988), which is hereby incorporated by reference in its entirety), or the Megalign® program of the LASERGENE® bioinformatics computing suite (DNASTAR® Inc., Madison, Wis). Sequences may also be aligned using algorithms known in the art including, but not limited to, CLUSTAL V algorithm or the BLASTN or BLAST 2 sequence programs.

[0055] As used herein, “cytoplasmic incompatibility” or “CI” is a phenomenon observed in certain arthropods, particularly insects, where the reproductive success between individuals is affected by the presence of specific intracellular bacteria (e.g., Wolbachia and Cardinium endosymbionts). This incompatibility occurs when infected males mate with uninfected females or females infected with a different strain of the bacteria, leading to embryonic lethality or reduced fertility. The underlying mechanism involves the modification of the sperm by the bacteria, which can only be properly counteracted by similarly infected eggs, ensuring the bacteria's propagation through the host population. The term “cytoplasmic incompatibility factor” or “cytoplasmic incompatibility gene” refers to the genes or the factorsencoded by a prophage from bacteria which provide a function that is required and / or beneficial to produce the natural genetic drive mechanism of cytoplasmic incompatibility (CI) used by various, unrelated bacterial infections (eg., Wolbachia and Cardinium endosymbionts). Cytoplasmic incompatibility factors can include those factors that induce the CI and can also include those rescue factors that counteract the CI. In some embodiments, a single bacterial operon may encode multiple cytoplasmic incompatibility (CI) factors. In some embodiments, a single bacterial operon may encode a factor that induces the CI and can also encode a factor that can counteract the CI (for example, a rescue factor).

[0056] All aspects of the present disclosure can be earned out with any of the embodiments disclosed herein.Arthropod Endosymbionts

[0057] Reproductive microbial symbionts are widely distributed in diverse animal and plant hosts (Perlmutter & Bordenstein, “Microorganisms in the Reproductive Tissues of Arthropods,” Nature Reviews Microbiology 18:97-111 (2020), which is hereby incorporated by reference in its entirety)- In the most speciose group of animals from the phylum Arthropoda, maternally -inherited bacterial symbionts in the testes and ovaries selfishly hijack host reproduction to enhance their own transmission. The archetypes of this selfish strategy are the genus Wolbachia that cause a widespread, paternal-effect, embryonic lethality termed cytoplasmic incompatibility (CI) upon mating between Ifo / dacte-carrying symbiotic males and aposymbiotic females (Yen & Barr, “New Hypothesis of the Cause of Cytoplasmic Incompatibility in Culex Pipiens L,” Nature 232:657-658 (1971) and Shropshire et al., “Symbiont-Mediated Cytoplasmic Incompatibility : What Have We Learned in 50 Years?,” Elife 9:e61989 (2020), each of which is hereby incorporated by reference in its entirety). The lethality is rescued by females carrying the same compatible strain of Wolbachia that is transmitted to the next generation.

[0058] As disclosed herein, a “rescuing bacterial endosymbiont” is a compatible bacterial endosymbiont that can rescue lethality caused by CI or by the chemical or genetic means described herein. A “non-rescuing bacterial endosymbiont” is a non-compatible bacterial endosymbiont that cannot rescue lethality caused by CI or by the chemical or genetic means described herein.

[0059] Wolbachia symbionts are in the vanguard of global vector control programs as releases of Ci-inducing mosquitoes suppress the size of vector populations (Zheng et al., “Incompatible and Sterile Insect Techniques Combined Eliminate Mosquitoes,” Nature572:56-61 (2019) and Crawford et al., “Efficient Production of Male Wolbachia-infected Aedes aegypti Mosquitoes Enables Large-scale Suppression of Wild Populations,” Nat. Biotechnol. 38:482-492 (2020), each of which is hereby incorporated by reference in its entirety) and replace aposymbiotic vector populations with ones that cannot transmit arboviruses to humans (Hoffmann et al., “Successful Establishment of Wolbachia in Aedes Populations to Suppress Dengue Transmission,” Nature 476:454-457 (2011), O’Neill et al., “Scaled Deployment of Wolbachia to Protect the Community fromNec / c.s' Transmitted Arboviruses,” Gates Open Res. 2:36 (2018), Utarini et al., “Efficacy of IFo / doc / zzo-infected Mosquito Deployments for the Control of Dengue,” New England Journal of Medicine 384(23):2177-2186 (2021), Gesto, “Large-Scale Deployment and Establishment of Wolbachia into the Aedes aegypti Population in Rio de Janeiro, Brazil,” Fr ont. Microbiol. 12:71110 (2021), each of which is hereby incorporated by reference in its entirety).

[0060] Wolbachia can also inhibit virus transmission in an agricultural pest of rice crops (Li et al., “Stable Introduction of Plant-Virus-Inhibiting Wolbachia into Planthoppers for Rice Protection,” Current Biology’ 30(24):4837-4845 (2020), which is hereby incorporated by reference in its entirety)- However, the efficacy of Wolbachia-based releases is potentially vulnerable to changing climate conditions (Salje & Jiggins, “Risks of Releasing Imperfect Wolbachia Strains for Arbovirus Control,” Lancet Microbe 5(7):622-623 (2024), which is hereby incorporated by reference in its entirety), forecasting an interest to explore adjunct and alternative approaches for arthropod control, particularly in hosts resistant to Wolbachia or CI.

[0061] Disclosed herein is the de novo engineering of CI in the absence of the symbiont and its CI genes by chemically inhibiting a host enzy me involved in reproductive epigenetics or using genetic means to do the same. This autonomous replication of a symbiotic trait provides the means to artificially induce CI in arthropods.

[0062] The genus Wolbachia is an archetype of maternally inherited intracellular bacteria that infect the germline of millions of invertebrate species worldwide and parasitically alter arthropod sex ratios and reproductive strategies to increase the proportion of infected females (the transmitting sex) in the population. The most common of these reproductive manipulations is cytoplasmic incompatibility (CI), typically expressed as embryonic lethality in crosses between infected males and uninfected females. This lethality is completely rescued by females infected with the same or a similar Wolbachia strain.

[0063] Wolbachia pipientis is an obligate, intracellular a-proteobacteria and a member of the Rickettsiales family. These gram-negative bacteria are not culturable outside of hostcells and, as a result, knowledge on Wolbachia symbiosis has only increased in the last two decades owing to molecular techniques. Once considered an obscure bacterium in a few insect species, the most recent meta-analysis estimates that 40% of all arthropod species are infected with Wolbachia as well as 47% of the Onchocercidae family of filarial nematodes.

[0064] In some embodiments, the endosymbiont is from the genus Wolbachia. In some embodiments, the endosymbiont is from the genus Cardinium. In some embodiments, male arthropods are not infected with a bacterial endosymbiont. In some embodiments, female arthropods are not infected with a bacterial endosymbiont. In some embodiments, the male arthropods are infected with a bacterial endosymbiont. In some embodiments, the female arthropods are infected with a bacterial endosymbiont. In some embodiments, the male arthropods are infected with a Wolbachia endosymbiont. In some embodiments, the female arthropods are infected with a Wolbachia endosymbiont. In some embodiments, the male arthropods are infected with a Wolbachia pipientis endosymbiont. In some embodiments, the female arthropods are infected with a Wolbachia pipientis endosymbiont. In some embodiments, the male arthropods are infected with a Cardinium endosymbiont. In some embodiments, the female arthropods are infected with a Cardinium endosymbiont.Small Chemical Control of CI

[0065] Histone acetylation is an epigenetic histone modification to regulate gene expression and reprogramming of insect spermiogenesis. Histone acetyltransferase (“HAT”) can acetylate all four core histones H2A, H2B, H3, and H4 with stronger preferences for H3 and H4 (Dancy & Cole, “Protein Lysine Acetylation by p300 / CBP,” Chem. Rev. 115:2419— 2452 (2015), which is hereby incorporated by reference in its entirety). Histone acetyltransferases do so by transferring an acetyl group from acetyl-Coenzyme A (acetyl- CoA) to the 8-amino group of lysine side chains. As the lysine-rich histone tails are positively charged and interact with the negatively charged nucleosomal DNA, HAT-mediated acetylation weakens histone tail-DNA interactions (Bannister & Kouzarides, "Regulation of Chromatin by Histone Modifications,” Cell Res. 21 :381-395 (2011), which is hereby incorporated by reference in its entirety), causing histones to be removed.

[0066] One aspect of the present disclosure relates to a chemically treated male arthropod, where the male arthropod has at least 10% less viable offspring when mated with a female arthropod having no endosymbiont or a non-rescuing bacterial endosymbiont in comparison to mating with a female arthropod having a rescuing bacterial endosymbiont.

[0067] In some embodiments, male arthropods are contacted with a chemical that is a histone acetyltransferase (“HAT”) inhibitor. Histone acetyltransferase inhibitors function by competitively binding to the active sites of histone acetyltransferases, thereby preventing the transfer of acetyl groups to histone proteins. Histone acetyltransferase inhibitors include but are not limited to anacardic acid, garcinol, a garcinol analog, curcumin, y-butyrolactone, MB- 3, isothiazolones, quinoline derivatives such as MC1626, C646, 1-CBP112, MG149, NU9056, EP300 / CBP inhibitor-2, and TIP60 inhibitor-1. In some embodiments, the histone acetyltransferase inhibitor is anacardic acid. In some embodiments, the histone acetyltransferase inhibitor inhibits HATl.[006S] In some embodiments, contacting the male arthropod with a histone acetyltransferase inhibitor comprises contacting the male arthropod during development of the male arthropod. In some embodiments, said contacting is carried out at a larval stage of the male arthropod. In some embodiments, said contacting is carried out at first to second instar larval phase. In some embodiments, said contacting is carried out during sperm development in the male arthropod. In some embodiments, said contacting is carried out prior to spermatocyte formation.

[0069] Another aspect of the present disclosure relates to an arthropod control formulation comprising a histone acetyltransferase (HAT) inhibitor and a carrier comprising the HAT inhibitor, where the carrier is suitable for delivering the HAT inhibitor to a male arthropod to cause a reduction in an ability of the male arthropod to produce viable offspring with a female arthropod (i) lacking a bacterial endosymbiont or (ii) having a non-rescuing endosymbiont compared to a female arthropod having a rescuing bacterial endosymbiont.

[0070] In some embodiments, the carrier comprises a food. In some embodiments, the carrier comprises a spray. Carriers suitable for delivering the histone acetyltransferase (HAT) inhibitor to a male arthropod can be diverse, depending on the application method and the target arthropod species. In some embodiments, the carrier is a bait matrix, such as a sugar solution or a protein-based food, which is attractive to the target arthropod. In some embodiments, the application is intended as a spray. In some embodiments, the carrier comprises a liquid formulation that can be atomized into fine droplets. In some embodiments, the carrier comprises water, oil, or a combination of both. In some embodiment, the carrier comprises a surfactant to improve the spread and adhesion of the spray on the arthropod's body. In some embodiments, the HAT inhibitor is dissolved or suspended in the liquid carrier, allowing it to be delivered to the arthropod upon contact with the spray. In someembodiments, the carrier comprises dusts, granules, and / or microcapsules. Any nontoxic carrier that is suitable for contacting a male arthropod is encompassed herein.

[0071] In some embodiments, the amount of HAT inhibitor is 10, 20, 30, 40, 50, 60, 70, 80, 90, 100, 150, 200, 300, 400, 500, 600, 700, 800, 900, or 1000 pM or any number or range therein. In some embodiments, the HAT inhibitor is more than 1000 pM. In some embodiments, the amount of HAT inhibitor is 0.01, 0.1, 0.5, 1, 5, 10, 20, 30, 40, 50, 60, 70, 80, 90, 100, 150, 200, 300, 400, 500, 600, 700, 800, 900, or 1000 mg / ml, or any number or range therein. In some embodiments, the HAT inhibitor is more than 1000 pM or more than 1 g / ml. In some embodiments, the HAT inhibitor is present at a concentration capable of reducing fertility of a male arthropod to cause a reduction in an ability of the male arthropod to produce viable offspring with a female arthropod of the same species (i) lacking a bacterial endosymbiont or (ii) having a non-rescuing endosymbiont compared to a female arthropod of the same species having a rescuing bacterial endosymbiont.

[0072] As shown in FIGs. 3A-D, Histone H3 and H4 acetylation levels are reduced in aposymbiotic CI males treated with the HAT inhibitor, anacardic acid. In some embodiments, contacting a male arthropod with a HAT inhibitor comprises reduced histone H3 and / or H4 acetylation levels during sperm development in comparison to a non-contacted male arthropod. In some embodiments, histone H3 and / or histone H4 acetylation levels are reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2, 2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold. In some embodiments, the reduction of H3 and / or histone H4 acetylation levels is greater than 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, histone H3 acetylation levels are reduced by at least 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, histone H4 acetylation levels are reduced by at least 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, H3 and / or histone H4 acetylation levels during sperm development in the contacted male arthropod are reduced by at least 20%. In some embodiments, H3 and / or histone H4 acetylation levels during sperm development in the contacted male arthropod are reduced by at least 40%. In some embodiments, histone H3 acetylation is reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2, 2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold. In some embodiments, histone H4 acetylation is reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2,2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold.

[0073] Histone acetylation changes during sperm development can be made in comparison to males that are not contacted with the HAT inhibitor or genetic methods that affect acetylation as described infra. In some embodiments, the histone H2B acetylation levels during sperm development stages are not reduced by treatment with the HAT inhibitor. In some embodiments, the histone H3 and / or histone H4 acetylation levels are reduced during post-meiotic sperm development by treatment with the HAT inhibitor. In some embodiments, the histone H2B acetylation levels are not reduced during post-meiotic sperm development by treatment with the HAT inhibitor.

[0074] The reduction in histone H3 or histone H4 acetylation can be measured at different stages of sperm development as disclosed herein. For example, AA-treated CI males showed significantly reduced H3ac testes signal in all post-meiotic sperm development stages (fold change reduction in young elongating: 1.45; early canoe: 3.27; late canoe: 5.0) compared to DMSO testes (FIGs. 3A-B). Similarly, the characteristic pattern of H4ac was detected in all post-meiotic stages of DMSO-testes, whereas in sharp contrast, no to low H4ac signals were detected in AA-testes from round spermatids till the elongating, early canoe spermatid nuclei stages (fold change in round onion: 18.1; young elongating: 12.3; early canoe: 16.4) (FIGs. 3C-D). Both H3ac and H4ac levels did not differ in spermatocytes of AA- and DMSO-treated testes during the early stages of spermatogenesis (FIGs. 3A, 3C). Exemplar}' non-limiting methods of measuring acetylation are disclosed herein in the Examples. In some embodiments, the HAT inhibitor causes protamine depletion.

[0075] In some embodiments, the male arthropods contacted with an acetyltransferase inhibitor have reduced numbers of offspring when mated to a non-endosymbiont infected female or a female with anon-rescuing endosymbiont relative to a female with a rescuing endosymbiont. In some embodiments, when mated to a non-endosymbiont infected female, the male arthropods contacted with an acetyltransferase inhibitor have a median embryonic hatch rate of about 0%, 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, or 90% (or any number or range therein) of the median embryonic hatch rate when mating with to a female with a rescuing endosymbiont. In some embodiments, viable offspring are reduced by at least 5%, 10%, 15%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%,80%, 90%, 95% or 100% (or any number or range therein). In some embodiments, viable offspring are reduced by at least 20%.Genetic Control

[0076] Another aspect of the present disclosure relates to a genetically engineered arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyl transferase gene or a promoter operably linked to a histone deacetylase, where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm development in a male arthropod causes a reduction in ferti 1 i ty and / or the ability of the male arthropod to produce viable offspring with female arthropods lacking a bacterial endosymbiont or having a non-rescuing endosymbiont in comparison to female arthropods having a rescuing bacterial endosymbiont.

[0077] Histone acetyltransferase (HAT) and histone deacetylase (HD AC) are two enzymes that play pivotal roles in the regulation of gene expression. HAT functions primarily to acetylate the lysine residues on the histone proteins, a process that unwinds the DNA, making it more accessible for transcription and thus promoting gene expression. On the other hand, HD AC operates in a contrasting manner, removing the acet l groups from the histone proteins. This action results in the condensation of the DNA structure, thereby inhibiting gene expression.

[0078] In addition to chemical treatment to affect histone acetylation during sperm development in arthropod males, non-transgenic genetic methods such as genome editing, and transgenic methods such as reducing gene expression using RNAi, and / or transgenic overexpression of target genes can be employed. Nucleic acid molecules that reduce abundance of histone acetyltransferases include inhibitory RNAs. Non-limiting examples of inhibitory RNAs include antisense RNAs or RNAi, such as short interfering RNAs (siRNA), short hairpin RNAs (shRNA), and micro interfering RNAs (miRNA).

[0079] Antisense methods that can be used to inhibit the in vivo abundance and / or translation of genes and subsequent protein expression are well known in the art (e.g, U.S. Patent No. 7,425,544 to Dobie et al.; U.S. Patent No. 7,307,069 to Karras et al.; U.S. Patent No. 7,288,530 to Bennett et al.; U.S. Patent No. 7,1791796 to Cowsert et al., which are hereby incorporated by reference in their entirety7). Antisense nucleic acids are nucleic acid molecules (e.g, molecules containing DNA nucleotides, RNA nucleotides, or modifications (e.g, modification that increase the stability of the molecule, such as 2'-O-alkyl (e.g, methyl) substituted nucleotides) or combinations thereof) that are complementary to, or that hybridizeto, at least a portion of a specific nucleic acid molecule, such as an mRNA molecule (See e.g., Weintraub, “Antisense DNA and RNA,” Scientific Am. 262:40-46 (1990), which is hereby incorporated by reference in its entirety). The antisense nucleic acids hybridize to corresponding nucleic acids, such as mRNAs, to form a double-stranded molecule, which interferes with translation of the mRNA, as the cell will not translate a double-stranded mRNA. Antisense nucleic acids used in the methods of the present disclosure are typically at least 10-12 nucleotides in length, for example, at least 15, 20, 21, 22, 23, 24, 25, 50, 75, or 100 nucleotides in length or any nucleotide or range therein. The antisense nucleic acid can also be as long as the target nucleic acid with which it is intended to form an inhibitory duplex. Antisense nucleic acids can be introduced into cells as antisense oligonucleotides or can be produced in a cell in which a nucleic acid encoding the antisense nucleic acid has been introduced, for example, using gene engineering methods.

[0080] siRNAs are double stranded synthetic RNA molecules approximately 20-25 nucleotides in length with short 2-3 nucleotide 3' overhangs on both ends. The double stranded siRNA molecule represents the sense and anti-sense strand of a portion of the target RNA molecule such as a HAT RNA. Numerous reports have been published on critical advances in the understanding of the biochemistry and genetics of both gene silencing and RNAi (Matzke et al., “RNA-Based Silencing Strategies in Plants,” Curr. Opin. Genet. Dev. 11(2):221— 227 (2001), which is hereby incorporated by reference in its entirety).

[0081] In RNAi, the introduction of double stranded RNA (dsRNA, or iRNA, for interfering RNA) into cells leads to the destruction of the endogenous, homologous RNA, reducing expression for that specific transcript. In both post-transcriptional gene silencing and RNAi, the dsRNA is processed to short interfering molecules of 21-, 22-, or 23-nucleotide RNAs (siRNA) by a putative RNAaselll-like enzyme (Tuschl, “RNA Interference and Small Interfering RNAs,” Chembiochem 2:239-245 (2001); Zamore et al., “RNAi: Double Stranded RNA Directs the ATP -Dependent Cleavage of mRNA at 21 to 23 Nucleotide Intervals,” Cell 101:25-3, (2000), each of which is hereby incorporated by reference in its entirety). The endogenously generated siRNAs mediate and direct the specific degradation of the target RNA. In the case of RNAi, the cleavage site in the RNA molecule targeted for degradation is located near the center of the region covered by the siRNA (Elbashir et al., “RNA Interference is Mediated by 21- and 22-Nucleotide RNAs,” Gene Dev. 15(2): 188-200 (2001), which is hereby incorporated by reference in its entirety). The dsRNA targeting a HAT RNA can be generated by transcription in vivo by inserting a nucleic acid molecule encodingcomplimentary sense and antisense sequences derived from a portion of the target sequence into a suitable expression vector having the appropriate 5' and 3' regulatory nucleotide sequences operably linked for transcription, and introducing the expression vector having the nucleic acid molecule into a suitable host cell or subject. Alternatively, complementary sense and antisense RNAs of the target nucleic acid molecule are synthesized in vitro (Fire et al., “Specific Interference by Ingested dsRNA,” Nature 391:806-811 (1998); Montgomery et al, “RNA as a Target of Double-Stranded RNA-Mediated Genetic Interference in Caenorhabditis elegans ” Proc Natl Acad Sci USA 95: 15502-15507; Tabara et al., “RNAi in C. elegans: Soaking in the Genome Sequence,” Science 282:430-431 (1998), each of which is hereby incorporated by reference in its entirety). The resulting sense and antisense RNAs may be annealed in an injection buffer, and dsRNA can be administered to the subject using any method of administration known to those of skill in the art.

[0082] Short or small hairpin RNA (shRNA) molecules are similar to siRNA molecules in function but comprise RNA sequences that make a tight hairpin turn resulting in a stem loop structure. shRNA is cleaved by cellular machinery into siRNA, which silences gene expression via the cellular RNA interference pathway.

[0083] In some embodiments, the inhibitory polynucleotide is a RNAi polynucleotide comprising a sense polynucleotide strand comprising at least 21 contiguous nucleotides from a HAT1 gene, and an antisense polynucleotide strand complementary to the sense polynucleotide strand, where the sense and the antisense polynucleotide strand form a duplex. In some embodiments, the RNAi polynucleotide comprises a sense polynucleotide strand comprising at least 20, 21, 22, 23, 24, 25, 30, 40, 50, 60, 70, 80, 90, 100, 200, 300, 400 or more than 400 contiguous nucleotides from a HAT gene sequence, and an antisense polynucleotide strand that is complementary to the sense polynucleotide strand, where the sense and the antisense polynucleotide strand form a duplex.

[0084] RNAi constructs may also include a polynucleotide sequence that separates the sense and antisense strands to facilitate duplex formation. In some embodiments, the RNAi polynucleotide further comprises a polynucleotide sequence heterologous to the sense and antisense polynucleotides.

[0085] In some embodiments, an arthropod is genetically modified to comprise a heterologous polynucleotide encoding an inhibitory RNA targeting a histone acetyltransferase (HAT), e.g., the HAT1 gene. In some embodiments, the RNAi polynucleotide comprises CAGAAGAATGTTGTGGACATA (SEQ ID NO: 1). In some embodiments, the RNAipolynucleotide comprises SEQ ID NO:1 and its antisense sequence TATGTCCACAACATTCTTCTG (SEQ ID NO:2). Other nucleotide sequences targeting a HAT1 gene are contemplated (such as other sequences within the HAT1 mRNA or coding sequence) and can be developed using methods known to those of skill in the art.

[0086] SEQ ID NO: 1 and 2 can be used in an RNAi molecule to reduce expression of a HAT gene. An exemplary HAT sequence is a HAT1 mRNA sequence from Drosophila melanogaster (GenBank Accession No. NM_169106.2, which is hereby incorporated by reference in its entirety), which is set forth below as SEQ ID NO: 3:AGTATGACTC AAAATTTGTC AGAAAAATGG AAGACTGCGA AATCAAGAAA ACATCGCAAC AAGTTAGCTG CAAACTACTT AACGGTCACA AGTAAAAAGT TCCGGAAAGA TCTTCTGGTC CACCATTTGC GAAAGCGAAT CGATTGCCGC CTTTTGAAGT ACCCAACAAT GGCACAAATC GAGAAGTACC AGGACTTTGT GATTGATGCT CTCGAAGTGG TAGACTTTAA GCTGATCCGT GACAAGGCCG ACATCAACAA TGATGCCCTG ACCTTTCATC CCGCAATGGC CCACCAGATT TTCGGGGAGA CCGAGACGAT TTTCGGCTAC CAGGACTTGC ACGTCCGTGT AATGTACACG GCAGGACCTT TGCACATCTA CTTGGGCGTC GATTACGGCA AACGGGTGAA TGAGATCTCC GGCGGAGAGA TCAAGGCAGA CGACGTCGTC AGCACAATTG CTCAGAGTCT ACCGGACGGC TGCTACTTCA TCAATCTTGA TGAGTTTCTT AAGACACTGG ACAAGGCTGA CAAATTTCAG CCCTTCGGTG AGAAGATAAG CGAATACAGG CGGGTATCGG ACGACGGCAG CGAGCGCCTT TTCGAGATCT ACCAGTGCGA GTACAAGAGC TCATCATTCT TAAAGTTCTT TGCCCGGCTG CAGACGTTCA TCTTGTGGTT CGTGGATGCT GCCTCCTACA TCGACACTGA TGATCCACAG TGGTGTTACT TCTTGAGCTA CGAAAAGTAC AAGAACAACG ATGGCCAGTG GCAGTATGCC ACAGCTGGTT ACACCACCGT ATACGAGTAC TACGCCTATC CGCAAAACAA GAGGCCCAGG ATAAGCCAGA TGCTGATACT GCCGCCTTTC CAGAAACTCG GCTTGGCTAC CCAATTGGTC GAGAGCATTT ACAGGTTTTA TCAGTCACAG AAGAATGTTG TGGACATAAC AGTGGAGGAT CCATCTGAGG ATTTCCAGCG TTTGAGGAAT TTTGTGGACG CGCGATCTTG CATGAAATTG AAGTCGTTTG CACCAGGCGA GATAGTCAAG GGCTTCAACA AGGAGATGGT GCGGGAAGCT CGCGAGGCTC TGAAACTCAA TCCCCTCCAG GTGCGTAAGG TGTACGAGCT TTTGCGTTTA TTCTACACCA ATGTGAAGGA TGAAAAGGAG TACCGACCCT ATCGTCTGGA AGTTAAGAAA CGGCTTAACG CGGTCTATTA CAAGCAGTTA AAGGATTTGA AGAAAATGGA GCGCTTCAAG ATGGACACTG AAATGCTGCG TGCTCGCCTG CCAACGGTTC AACAACGAAT GGAGCAGCTG CATGAGGAAT ACCGCGTGGT CGAGCAGGAG TACCAGAGCA CCATTGCCAA GTTGAGGGCCTATGTTGATT ACAAAATACA TTATGTTGCT ATAAGCCTAC AGTAGGATTC CTTTGAATAAAT AT TAG AAA TTAAATAT

[0087] In some embodiments, the HAT1 sequence is Drosophila melanogaster HAT1 and comprises a nucleotide sequence of SEQ ID NO:3 or a nucleotide sequence having at least95% sequence identity to the nucleotide sequence of SEQ ID NO:3. In some embodiments, the HAT1 sequence comprises a nucleotide sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the nucleotide sequence of SEQ ID NO:3.

[0088] The amino acid sequence of the protein encoded by Drosophila melanogaster HAT1 mRNA (SEQ ID NO:3) is GenBank Accession No. XP 001651817.1, which is hereby incorporated by reference in its entirety, which is set forth below as SEQ ID NO:4: MAQIEKYQDF VIDALEWDF KLIRDKADIN NDALTFHPAM AHQI FGETET I FGYQDLHVR VMYTAGPLHI YLGVDYGKRV NEI SGGEIKA DDWSTIAQS LPDGCYFINL DEFLKTLDKA DKFQPFGEKI SEYRRVSDDG SERLFEIYQC EYKSSSFLKF FARLQTFILW FVDAASYIDT DDPQWCYFLS YEKYKNNDGQ WQYATAGYTT VYEYYAYPQN KRPRI SQMLI LPPFQKLGLA TQLVESIYRF YQSQKNWDI TVEDPSEDFQ RLRNFVDARS CMKLKSFAPG EIVKGFNKEM VREAREALKL NPLQVRKVYE LLRLFYTNVK DEKEYRPYRL EVKKRLNAVY YKQLKDLKKM ERFKMDTEML RARLPTVQQR MEQLHEEYRV VEQEYQSTIA KLRAS

[0089] In some embodiments, the HAT1 sequence is Drosophila melanogaster HAT1 and encodes an amino acid sequence of SEQ ID NO:4 or an amino acid sequence having at least 95% sequence identity to the amino acid sequence of SEQ ID NO:4. In some embodiments, the HAT1 sequence comprises an amino acid sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the ammo acid sequence of SEQ ID NO: 4.

[0090] An exemplary histone deacetylase 1 (HDAC1) mRNA sequence from Drosophila melanogaster is GenBank Accession No. NM_139661.4, which is hereby incorporated by reference in its entirety and as set forth below as SEQ ID NO:5.CGGTCACACT GCCCATCTGC AGAGGTTTCC TCTTCGCGAA TTTGTCTTGT GCGCTCCGAT TTTGGCTATA TAGTTGCGCC AAAGTGCCAA AAATCGGACC GGCACCAAAG TAAACCATCT TCTGAGCTCG CAAGTTCATC CGATCGACGA AGGATAGAAA AACATCGACT TTAAGGGGAT TTAAGACCCG CCAACAGCAC CGCAAAATGC AGTCTCACAG CAAAAAGCGC GTTTGTTACT ACTACGACAG CGACATTGGC AACTACTACT ATGGCCAGGG TCATCCCATG AAGCCCCATC GCATACGCAT GACCCACAAC CTGCTGCTCA ACTATGGGCT CTATCGAAAA ATGGAAATAT ACCGTCCCCA TAAAGCCACT GCCGATGAGA TGACCAAGTT CCACTCGGAC GAGTACGTCC GGTTCTTGCG ATCCATTCGG CCGGACAACA TGTCGGAGTA CAACAAGCAG ATGCAGCGTT TCAATGTCGG CGAAGATTGT CCCGTCTTCG ATGGACTATA CGAGTTTTGC CAACTCTCCG CCGGAGGATC CGTAGCTGCG GCCGTAAAAC TGAATAAGCA AGCCTCGGAG ATCTGCATCA ATTGGGGCGG TGGGCTGCAT CACGCCAAAA AATCGGAAGC CTCGGGCTTC TGCTACGTCA ACGACATTGT TCTGGGAATT CTGGAACTGC TTAAATACCA TCAGCGTGTT CTCTACATAGATATAGACGT CCATCACGGC GATGGCGTGG AGGAGGCGTT CTATACCACC GATCGTGTGATGACTGTCAG CTTCCACAAG TACGGAGAGT ATTTCCCGGG CACTGGCGAT CTGCGAGACA TTGGCGCCGG CAAGGGAAAG TACTATGCGG TGAATATACC CCTGCGTGAT GGCATGGATG ATGATGCGTA CGAGAGCATA TTTGTGCCCA TTATCAGCAA GGTGATGGAA ACATTCCAGC CGGCAGCCGT GGTGTTGCAG TGTGGCGCCG ATTCGCTGAC TGGCGATCGG TTAGGCTGCT TCAATCTCAC CGTCAAGGGT CACGGCAAGT GCGTGGAGTT CGTGAAGAAA TATAACCTGC CATTCCTGAT GGTCGGCGGT GGTGGTTATA CCATTCGTAA TGTATCCCGC TGCTGGACCT ATGAGACCTC CGTTGCACTG GCCGTGGAGA TAGCCAACGA ACTGCCCTAC AACGATTACT TCGAGTACTT TGGGCCCGAT TTTAAGCTGC ACATTAGTCC CAGCAATATG ACGAATCAGA ATACATCCGA GTACCTGGAG AAGATCAAGA ACCGTCTGTT CGAGAACCTG CGCATGCTGC CTCACGCTCC GGGCGTTCAA ATCCAAGCGA TTCCCGAGGA TGCCATCAAC GATGAGTCCG ACGACGAGGA CAAGGTCGAC AAGGATGATC GCCTGCCGCA GAGCGACAAG GACAAGCGCA TTGTGCCCGA GAACGAGTAC TCCGATTCGG AGGATGAGGG CGAAGGCGGT CGCAGGGATA ACCGTTCGTA CAAGGGTCAG CGGAAGCGGC CGCGTCTCGA CAAGGACACC AACAGCAACA AGGCATCCTC AGAGACGTCC AGCGAGATCA AGGACGAAAA GGAAAAGGGC GACGGAGCTG ATGGCGAGGA ATCGACGGCG TCTAATACCA ATAGCAACAA CAACAGTAAC AACAAGAGCG ATAATGATGC CGGAGCGACA GCTAATGCCG GATCCGGTTC GGGATCAGGT TCCGGTGCCG GGGCCAAGGG CGCCAAGGAG AACAACATTT GACGTGGCGG CCGCAATTGG TTATAGAAGC CAATTAAGCG ACCGCCGAAG TACAAGCCAC CGGATTTCAC CTAGTGTCGT CTCATTCCCG CGCCCCCCTT CACCAACACG CCCCGTAGCC ATGTCCAGGG CTTTTTCTAC ACGCCCCTAC ACAAGGTCCC ACTGCGATCG GCTCCTTTTT TTCAGTGGGA ATATTCCCAA AGGGGGATAA ATCAGTAATC AGTGGGCGGA AACACAATTT ATTTTAAGAT CGAAAGTGTA TAGCTAAGCA AAAAGTGAAT GCATACTTAC AAGCGATCCA ATGTGGCGAA TAACGCAGAA ACTAATTTAA TCAAATAAAA ATATTTATAC ATTT

[0091] In some embodiments, the HDAC1 sequence is Drosophila melanogaster HDAC1 and comprises a nucleotide sequence of SEQ ID NO:5 or a nucleotide sequence having at least 95% sequence identity to the nucleotide sequence of SEQ ID NO:5. In some embodiments, the HDAC1 sequence comprises a nucleotide sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the nucleotide sequence of SEQ ID NO:5.

[0092] The amino acid sequence of the protein encoded by Drosophila melanogaster histone deacetylase 1 (HDAC1) mRNA (SEQ ID NO:5) is GenBank Accession No.NP_647918.2, which is hereby incorporated by reference in its entirety, and is set forth below as SEQ ID NO:6:MQSHSKKRVC YYYDSDIGNY YYGQGHPMKP HRIRMTHNLL LNYGLYRKME IYRPHKATAD EMTKFHSDEY VRFLRS IRPD NMSEYNKQMQ RFNVGEDCPV FDGLYEFCQL SAGGSVAAAVKLNKQASEIC INWGGGLHHA KKSEASGFCY VNDIVLGILE LLKYHQRVLY IDIDVHHGDG VEEAFYTTDR VMTVSFHKYG EYFPGTGDLR DIGAGKGKYY AVNI PLRDGM DDDAYES I FV PI I SKVMETF QPAAWLQCG ADSLTGDRLG CFNLTVKGHG KCVEFVKKYN LPFLMVGGGG YTIRNVSRCW TYETSVALAV EIANELPYND YFEYFGPDFK LHISPSNMTN QNTSEYLEKI KNRLFENLRM LPHAPGVQIQ AI PEDAINDE SDDEDKVDKD DRLPQSDKDK RIVPENEYSD SEDEGEGGRR DNRSYKGQRK RPRLDKDTNS NKASSETSSE IKDEKEKGDG ADGEESTASN TNSNNNSNNK SDNDAGATAN AGSGSGSGSG AGAKGAKENN I

[0093] In some embodiments, the HDAC1 sequence is Drosophila melanogaster HDAC1 and encodes an amino acid sequence of SEQ ID NO:6 or an amino acid sequence having at least 95% sequence identity to the amino acid sequence of SEQ ID NO:6. In some embodiments, the HDAC1 sequence comprises an amino acid sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the amino acid sequence of SEQ ID NO: 6.

[0094] Inhibitory RNAs targeting a histone acetyltransferase 1 (HAT1) gene from other arthropods can also be used in the methods of the present disclosure. Similarly, overexpression of histone deacetylase 1 (HDAC1) in other arthropods can also be used in the methods of the present disclosure. HAT1 and HDAC1 genes from various arthropods can be identified by sequence identity using BLAST searches to the nucleotide and / or amino acid sequences disclosed herein, as non-limiting examples.

[0095] In some embodiments, the histone acetyltransferase is a HATl mRNA sequence from Drosophila simulans. An exemplary Drosophila simulans HAT1 sequence is GenBank Accession No. XM_002102248.4, which is hereby incorporated by reference in its entirety, and is set forth below as SEQ ID NO:7:TTGTATTCAC CGTCTTGCGT TGTCCAACAC TATTGGAAAG CCAATCACTT TGCTTAGCCG CGAAAATTGC GCTAGAACAC CCGCAAAAAG CTGTCTGGTT TTATAAACAA CGTGCTTTTG GATTTTTAAA GCAACAAGGT AGCTTCAAAC TACGTGACAG TCACAAGTAA AAATTTCCGG AAAGATCTTC TGGTCCACCA TTTGCGGAAG CGAACTGATT GCCGCCTTTT GAAGTACCCC ACAATGGCAC AAATCGAGAA GTACCAGGAC TTTGTGATTG ATGCTCTCGA AGTGGTAGAC TTTAAGCTGA TCCGTGACAA GGCCGACATC AACAATGATG CCCTGACCTT TCATCCCGCA ATGGCCCACC AGATTTTCGG GGAGACCGAG ACGATCTTCG GCTACCAGGA CTTGCACGTC CGTGTAATGT ACACGGCAGG ACCTTTGCAC ATCTACCTTG GCGTCGATTA CGGCAAACGGGTGAATGAGA TCTCCGGTGG AGAGATCAAG GCCGACGACG TCGTCAGCAC AATTGCTCAG AGTCTGCCGG ACGGCTGCTA CTTCATCAAT CTCGATGAGT TTCTCAAGAC ACTGGACAAG GCTGACAAAT TTCAGCCCTT CGGTGAGAAG ATAAGCGAAT ACAGGCGGGT ATCGGACGAC GGCAGCGAGC GCCTTTTCGA GATCTACCAG TGCGACTACA AGAGCTCATC ATTCTTAAAGTTCTTTGCCC GGCTGCAGAC GTTCATCTTG TGGTTCGTGG ACGCTGCCTC CTACATCGACACTGATGATC CACAGTGGTG CTACTTCTTG TGCTACGAAA AGTACAAGAA CAACGATTGCCAGTGGCAGT ACGCCACTGT TGGTTACACC ACCGTATACG AGTACTACGC CTATCCGCAA AACAAGAGGC CCAGGATAAG CCAGATGCTG ATACTGCCGC CTTTCCAGAA ACTCGGCTTG GCTACCCAAT TGGTCGAGAG CATTTACAGG TTTTATCAAG CACAGAAGAA TGTTGTGGAC ATAACAGTGG AGGATCCATC TGAGGATTTC CAGCGTTTGA GGAATTTTGT GGACGCGCGA TCTTGCATGG AATTGAAGTC GTTTGCCCGG GGCGAGATAG TCAAGGGCTT CAACAAGGAG ATGGTGCGGG AAGCCCGCGA GGCTCTGAAG CTCAATCCCA TCCAGGTGCG TAAAGTGTAC GAGCTGTTGC GTTTATTCTA CACCAATGTA AAAGATGAAA AGGAGTACCG ACCCTATCGC CTGGAAGTTA AGAAACGGCT TAATGCGGTC TATTACAAGC AGTTAAAGGA TTTGAAGAAA ATGGAGCGCT TCAAGATGGA CACTGAAATG CTGCGTGCTC GCCTGCCAAC GATTCAACAA CGCATGGAGC AGCTGCATGA GGAATACATG GTGGTCGAGC AGGAGTACCA GAGCACCATT GCCAAGTTGA GGGCCTGATA GGGTCTGTGG ATCCAAAAGG TGTTGATTAA TTTATTTCGT TTCTGTTTTC AAATATGTTG ATTACAAAAT ACATTATGTT GCTATAAGCC TACAGTAGGA TTCTTTTGAA TAAATATTAC AAATTAAA

[0096] In some embodiments, the HAT1 sequence is Drosophila simulans HAT1 and comprises a nucleotide sequence of SEQ ID NO:7 or a nucleotide sequence having at least 95% sequence identity to the nucleotide sequence of SEQ ID NO:7. In some embodiments, the HAT1 sequence comprises a nucleotide sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the nucleotide sequence of SEQ ID NO:7.

[0097] The amino acid sequence of the protein encoded by Drosophila simulansHAT1 mRNA (SEQ ID NOT) is GenBank Accession No. XP_002102284. 1, which is hereby incorporated by reference in its entirety, and is set forth below as SEQ ID NO: 8: MAQIEKYQDF VIDALEWDF KLIRDKADIN NDALTFHPAM AHQI FGETET I FGYQDLHVR VMYTAGPLHI YLGVDYGKRV NEI SGGEIKA DDWSTIAQS LPDGCYFINL DEFLKTLDKA DKFQPFGEKI SEYRRVSDDG SERLFEIYQC DYKSSSFLKF FARLQTFILW FVDAASYIDT DDPQWCYFLC YEKYKNNDCQ WQYATVGYTT VYEYYAYPQN KRPRI SQMLI LPPFQKLGLA TQLVESIYRF YQAQKNWDI TVEDPSEDFQ RLRNFVDARS CMELKSFARG EIVKGFNKEM VREAREALKL NPIQVRKVYE LLRLFYTNVK DEKEYRPYRL EVKKRLNAVY YKQLKDLKKM ERFKMDTEML RARLPTIQQR MEQLHEEYMV VEQEYQSTIA KLRA

[0098] In some embodiments, the HAT1 sequence is Drosophila simulans HAT1 and encodes an amino acid sequence of SEQ ID NO: 8 or an amino acid sequence having at least 95% sequence identity to the amino acid sequence of SEQ ID NO:8. In some embodiments, the HAT1 sequence comprises an amino acid sequence having at least 70%, 75%, 80%, 85%,90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the amino acid sequence of SEQ ID NO: 8.

[0099] In some embodiments, the histone acetyltransferase is a HATl gene sequence from Aedes aegypti. An exemplary Aedes aegypti HAT1 gene sequence is GenBank Accession No. NC_035109.1 REGION 140508611..140523812, which is hereby incorporated by reference in its entirety, and is set forth below as SEQ ID NO: 9: GCTCAAGCTG TCAGTTGCAA ATCAACACCG ATGACCTTTG TGCGCGAATT TTTGTTTGGT GTCGCAATTC TAATGAAGAT CATTTAAAGT AAATTTTTCG GGAAATAATT CATGTTTTAA TCCAATAACA CGCTGATCTG GCTGGCTTTG TCATTATTCA TACGGTAAGT TTCACATAGA GAGTTTTGTT GATTGATAGC GACGGAATAA TCAAACATAA AAGTTGACGT TGAGGTTACG GAGTCCCAAA CAAAATTTGG CGCAATTTCA TGGAGGAATT TTGGCCGCTC TATTTTCAGG AAACTTCCCT CGATTTGTGC AGTCCACCAT GACCGCTATT TCCAGTTTGC AGAATTTTGT CACCGACGCC CTCGAGTGCA CCCAGTTCCG ATTGATTCGG GATGATGCTG ACTTTGAGGA CGAATCGGTC GCGTTCCACC CGGAGATGGC TCATCAGATA TTCGGGGAAC AGGAAAGCAT CTTCGGATAC CGGGATCTGC AGATCGATGT GTGCTTCGCG GCCAGTTCGC TAGATATTTA TTTCAACATC AAATACTCGA AGAAGGTAAG TGGATAAATT TGAGGAATTT TTACAGTGAG TTGGTTAAAA AAAATACTTC ATCAATAATC ATATTTGTAT AACTGTCCCA TGTATATAGA AAATTCCATA GAACATAGGC TGTAGCGTAA CCATGATTTA GGAGACTGGG GCGCTATATG TTTCAAATTT GAAGATAATT GATTGATTTG CATGGTGCAT TCGTGCCAGC CTTGTTTACA GTTCAAAGCT TCTTCACAAT AATGTGTATA GTCAGTGGCG TCTCGTCCTA AGGTGCACTT GGTGCACTGC ACAGTCAAAG ATTTCAAACT AACTAATTAA ATATATATAT TATAATTCTC AATACTTCTA ACATTCATAA TGTGTAGTTG TTTGAAAATG TTGTTTGATT TCTTCACCTT CAATTGGTGC ATCGCCCTCA GTGAACAAAC CGTTTGATTC TCCCCCCATG GTTTTGCTCA ACCAGCAAGC GAAAACAGCT AAGCAAACAA GACAAACTGC CATGCGTCTC CATCGTATTG CAGAGAATGT TCATTTCAGC CGTGCACTCT TTCTTGTGCA CGAAAATAGC GTGTATGGAC ACAGCAGGAA TGTGCATTTG TATTAGCATT TCTAGAGCAA CAGTGGCGTT GTATAGGTAA AATCTGGTTC ACTCCAATTT GTGCACCCGA GAGAGAAATT GCCCATGACG CTCGCATTGG TTTGCAAGAA TTGAGAAAGC GCGCGCAACT GGCGTCACGA CGTCTACTGT TGGAAGAGAA AGCGTTAACG CCATACATGC TTTGTGTGTA TATATGTTGC ATGGAGCTTC CAGGTGTCGC GCGATAGTAG GCGCAATATT GATTGGAACC AAAACTAGAT GTTATAAACA TATTGGCCAA ATTGATTATG TTTTATCTTA TTAAATACAC AAAATTTATA TGGCTTTGAA TTGATTTATA AACTATGTGT GAATATAGTT TTTCTTGCTC AATAGACCGA CAACTGATAA ATTGCGGTGG GCGTAAAGTG GGTTTCAGCG TTCAACAGTC AGTCATCATT TCACTTAAAA GCTTTGATGT ATCAACAAAA GTATAAAAAG AGATGTACCA GCGCTGGACA TCGTCGAAAA GCTAAGATAT CGTGATACAA TTGACAAGCT GTAAATAATA AGAATAATTA TATAGAATAC AAACGTTAGG AATAATGGAG ATTCGGAATT TCGCGAAAGT CACGAAATCC ACGGAATTTA TCCTTCATCGCGAAACTTGG AAAATAACGC GTAAATAGTT GAATTCTGAA AGTTTATATG ATAATCTCAGAAAAGTTCAG ATTTTTGCTC ATAAGAATGG TTTATTTATT TGTAAGCCGA GTTTCATATAAAAAAATGTG CTAATCGTTT AGTTTCAAGT TAAACGTCTT TCTTCTTGCT TTATATTTTGTATTGATAAT ATGATATTAA ACTTCGCAAA ATACAAAACA TGCTTGAAAT TTACGCATATAAAGGTCATT TTTACTGTAA ATAAAATTTT TTTGTCAAAT ATTATCAAAC TATGGGGCCGCGATTAATAC GCAGAATTTT TCTTTCATTG CCTTGATATC AGCGAAATTT TGAAAATTTTTCTTGTCCTT AAACATTGTA TATTGGAATC ATCCAAAATT CTTAAGACTT ACTACTAGTGATTAAAACGA AGAAAACAAA CAAAAATGCG TCCAATGAGC AACAAATATA ATAGTTTCCAGACACTTCAA GATTGCATAC AGTATTGGAA ATTACATTTT GAGAAGAAAT TACATTTTAAAAAATCAATT GAAAAATTCG TATTCATAAT TCAAAATAAA CGGTTTGTGC GAGTCAGTGCACAGGTAATC AAATTCCTCA CGGGACGCCT CTGTGTATAG TACGGTTTCA TTCAAGCATTCCAATAACTT CTTCTTTGCT TATGGCATCT TGGAAACTGG TCATGTTGAT CTCGTTTTCCTGTCTTTCCT TCTAGTACTT TCTGCATTAA AGTATCAGGT GTCTGACAAC TATAATTGTGAAGATAATGC TTTCTTTATT TAAAAGAAAT GAGTTAAAAA TTCCCTGTCC TTTGGTTCAGAAAGGGGTAC GGGCGCGTTC TGCCAACTCG GTCGAGGCAG ATTTCTTGTG TGAACAAGTTACAACATGGA CGTTCTAAAG TGCCAGAATG AAGTGTTGTG CTTGTAGGGA GCACTTGAAGATGGGATGGA AGGTAATTAC TTCGCGTTCG TGCTATACTT CTGCTTATCT ATAAAGTTATATACAATAAC CGACCCTTAG CTTAACTATT TAAATTAAAA GTTGCAACTA CAATAATCGTTTTGAGCTTA CTAACATGTC GTCGCGTTAA TTTAATCATA ATGAGCTTTT ATCTGCATCCGTCGAACCAT TCTGAATTCC ACGAAACACT TGAAGCAAAC TTCAGGAGGC TTTTGTATGCTCAGTCGGCA AACCGTCCAG CCTACCTTGA GCATGCCCAA GTTCTTCGTT TTGCTGTCCTCAGCAACCGG TAGCCTGATC GCCGCCACCC GGGTTCCCTG CGGGCCCTTT CTAAGGTGGATAGCCTTATT GGCTACGTTG ATGTTACACT TCTCTTTGAG GGCAGCCGAG ACCTCCGCTGCATCGGTGAC CTCATCCAGA TTTTTACACT GGAGAGTCGC TTCTGCAGTA AGGGATCTTACATCAACCTC GTCGCCAGTT GCTTATAGCC TGCACCCTTT GCCTTGGCAT CCTTCTTTAAGACTAGGATC ATTTCACCAA TCCTAGTTCG CCTTATGCTC CGCACGTTTG CGCCCAATGGCGATAACTTC TCCATGATTT GCATCGCCTT CAGAACCTCG GCGTATTTCG CTCCCTCCGGTTTGATAACT AGGGCTTCAC TTAAATTTCT ATGTTTCGCT GTTTTCAGTT TAGCACTCTCCTTGGGCTCC GTTTTCGGTT TCCTCTGTTT TTTTCTTATT TCCAACTCGT ATCCACGAGTTGCTGTTGCC TTGCGCCCGT GGTTCCTGTG GATGCACTGC CTGGTTCGGG TTATCCCGTGGTTCCTTTTT CTTGGCTTTC TTGGCCTTAG GCTTGGTGTT GGCCTCTCCT TTGTTGCCATGTCCTCTTGA TCGCGGGGCT TGGCTCGTGC CAGCTTTTTC CTTAGCAACC GTGTTGTCTGTGAAGGAAAA CACTTCGGTC TGACAGGACT TAGTATCTCT TTTGCCTTCT ACCTTGCCAAGTTGTTCCTT GGCTTTCTCG TAGTCCTGCT TTGCAGCTAG GACTAATTGT CGCAATTTTCGGTAGACTTG TTTTCAAGTC TTTGCTGATA TTGCTTTTTC TCTTAACAAA CTCGATGATGACATCAAGTT GCTCAGCAGC TACCTGCATT TTAGGGAGGT ACTCCCTATT GCGATCCATGGCCTCGATTA GTCCTGGGCC ATCGGTCACC TCACATGTTG CACTGCAGCG GCCAGTGTCTGCCGTCGTCA CAGTATCTAG GCTTTTGCCC AGACTGCTTT CACTTAAATT GATGTGTTGGCGATGACAGG GTGTCTACTA CCTGGAAAAA CCTGGAATTA TCAGGGAATT TTTTCAACCTGGAAAAAACC TGGAATTCTC AGGGAATTTC GATCACAATC AGGGAAATTA TGTTGAGGCAGTAATTCATG ATAGAATATG TTCACGCCAA AGTATTTTTG CGTCATAACT CAGAGCTATCATTAATTGCC AGAAAACTCT CTTCATTCGA AAAAAAATTC GGCTGCGCCG CTCTCCAAATACTATATGAC ATGTTTAGTG AGGATCCATT CGAGTATGAA ATTTAATCAG CTTATCAAAACATGATCTAA ATTTATTCGG TGAAGAATTA ACATGTCTAG GGCTGGTAGC AGGTTTCTTTTTATATACCA TTGTCTCTAA AAAGTTTATA TTTTTGATGA AGGTGTCTGA AGTCTCTATTTTGACCAAAA TAGTCTCTAA ACTCTCTCTT TTTTCTTAAC TTGCTTGCAG ATAACTGATATGCAAAATTT CTTCTCGTAT TTCTCGAGAA CAGCTTCAGG AAAATATCCT TATGGTTCCTCCAGAGATTT TACCAGGAAT TTTTCAAGAA ATTCCTTTAG CATTTTTTTC ATAGGATGCTTATAGAAGTT CCTCCATAAT TTGCTTTAAG AAATCCTTGA ACATTTCAAC GTTACTACCTCTAGTAATTT TCTCAGGGAG TACTTTGAAC AAAGACTATC CTGGGACTTC CTCCAGATATTTTTCACTAA CTATTCAACG AGTTCTTCAG TCGAATTTCT CCCAAGAAAA CCCGACAAGAATTTCAACGC CATTTCATCT ACGGTTATTC CAAAGAATAG ATCCAAAAAC TTCTCCAGGGGTTATTCGAG TGAATATATT AAGAAAATTT CAAATTATTC CCTCAATTAG ATCTGTGTTTCATTATTAAG TTTCTTTACA TTTTTTTCTA GGAATTTCTC CAGCATTTCA TCCAAGGATTTGGAGTTCAT CCAAACGTTT TTCATTGTTT TTAAAAAAAA TCCCCATGGA ATATTGAAAAGCTCATTTAA GGTTTTACAC AATTTAATAC CACAGCATTT TTTCCAAATC CTTCGATTGTAGGAATGATT ATTCCTAAAC AATTTTCTAG TGATTTCATC CTGTGTTCCA GGAGATCCTTTGCAATATCC TGCGAGATAT TATGATGTTT CTTATTTTTT TTAATTATCT TAGTAATAATCTTGGAAATC TTCTAAAAGT TCAACAAGAG TAAATCTTCC TGAAAATCCT CTTTCATGTAGGGTGACCTA AGAAATTAAG AAACTGTAAC AAAAGATACC CGTAAATTTG CTTTAGGAATGCGTCTCGAA TTTTCTCAAC AATTTCTTAG CTTTCTGAGA AATTTCATTC 7LAATTTCTTTGGATAAAACT CCTGGAACAA TGCTTGAGAA AAAAAGTTTT GAAAACTTCC TTTATAAATCTCTAGAATTA ATAGAGGAAT TTCCGAAGAA ATCCTAGGAG TAATCTCTGA AGTAATTCCTGAAGGTATCC TCAGAGAAAT TTGTTGTCAA AATCTTATGA AAATTCTGAA AAACAATTTTGAAGAAAATC CTGAGAAATC CTTGGAGACA TCTGGAGAAG TTTCTGGTTG AATGTTACAACATGTTTTAA ACAGGAGTCT TGGAGGATGT TATAGAAAAT CTGATTGAAG AATCACCGGATAAAATTTTC GAGGAAAAAG GGCTTTTTCA AAAAATATTT GAAATGATAT GTGAATGAATTTCTGATGAA AGCCTTGAGG TATTTTTTGT GATATTTTAT GGAAAAATCC TGGTTCTATTTTGAGATATC CAAAACAATT TTTTTGGAGG AATTCCTTAG GAAAATTTTT GGAGTGAAAATCTCAAATAT AGATATCAAT CCATCCCTCT CGTAATGTCG TCGCAATGCA TCAATGAATGTTTGTCCTTT ACTGGCATAT ACCAGATGTG CTTGTATGTC GTCGTAATGT CACAAAAAAATTGAAAAAAA AAAAGTTTTA AATATAGGGG AACTTACGTA TTCTCGGCAG TCTTAGTCGATGCCGCGCTT CTTTTGTAAT TTTCTCCGAC ATCAGTAGAT AAATTCCGTG TTTGTCTGTTTACATTTATG CGTTGCTCAA TCCTCTATCG ATTCACACCG GCAGACTTGT CAAAATGCTTTTGGAAACGT TTTTACAACG CTGCGAACAT CTCTTGTCAG TGTGACTATT GTCGGCAACTTTCCCATAAG AACTGCTGGC GAATCTCTTC GGCAGCTCCA AATCAGTCGC ATTTTGAGGCGTGTTCATTT GAATTGATGA CATCTCTGGC GTTATTGTTT GTTCAGTCTC GGAAATCTCGTCAGTGGGAA GCGGACAGAA CAAAGAATCA TCTGAATGTT TTCATTGACG TGTTTTGATAGAATAATACT GTGAATTTGG CGTTTGAAAT TTGGTTGCCG ATAATAGTCG AATGGTGCCGAAGCCGTAAG TCTCCCTATT TTCAGGGAAA TGTCTGGTCT GATCAAACTA TTTTGTCCTGGTTCCTATTA GGTTTCGTTT TTTGATTACT GTTTTGTCTC TTTTCGGTCT CATTTTGGTTACTTTTTAAT CTCTTTTTTC TGGAAAAAGG TTTCTAAGTT TCATATTTTC AAGAAATTCATTTACCATGC ACAATTTTTA AAATTATTGG AAAATATTTG ATTTCAGAAA AAACAATTTTCGAAAAATAA ACTGGAATGT AAACAACTTT AAATTCAAAA TTTCGAACTA GGGAAAGTGTACCAGTAATG GCCATAGTGG TTCCCTATTT GGCCATATGT GAAATTTCGA TGACTTTCACATTTTTAATC TTTTTGAATG TTTTAACATC AAGATATATC CTATATCTTA CTGCTAAAACACACAAAATT TTTCAAAATG TGGAGACATT TAAGTTATCA CGTATGGCGA AATAGGGAACCACTATGGCC ATAACTGGTA CACCTACCCT ACATGATTAA TTTAAAACTT TTTCGAATGAAATGCAAAAA TTTATAATTT ATTTATATTA CCACAGTTTT CGAATAACTT ATGATCTCGCCCTAAAAGCC ATTGGTAACC ATGTGTATAG CTCGTTGTTT CTAAGCATTT GAATGGAGTTTCACGTTATT GACAAAGATA TGGTGAAGAA AGGATCATTC CATACCTGCC AGTAGTGTTGGTTTGGATCT TAATAATTGT TTTGCGCTGA AACAACGAGC TACACACGTG GTTACCAATGCGGCTCCTAG GGCGTTATCC CATGTTATGC GAGAGTTAAA TGTTATACGG TCGAACGTGATTACCTTGGG GCTTGACACA TACTGATGCA CTTGAATGTT CCGTTCAGTT CACAGAGCTCTTACAAACGG AATTAAGAAA TCATCCTTGG CAAATACGAT TCATTTGAAT AAAACTTAGCTTTTGTATTG TTATCATCTT CACATTGTAA TATTGGATAT TGGAATAGTT TCAGCATGCTTTACATAATG TTTAGAAGTG TATAAATAGC TATTAAATCG AAAACACTTT AAAATTTGTCATTAATTTGT TTTAGACAAA ATTAGAGTAT TTCTTGAACC TGGAATTATT TTTATAAAAC CTGGAAAAAC CTGGAAATAT CAGGGAATTT CATTTCGGTG AACGAGTAGA CACTCTGGAT GAGATTTAGT TGGCGACTAT TAATCATAGC AAACATAGAT TAGTTGATTG TTAAATACAT TCATTCCAAG TTTGTCCCTT GTCAAGATCA GAAGTTTTAT TTCAACCAAG AGTAGAATAC CCGGTCTACG GCCATTTACT ATCGAAGTTG ATGTTGGTCA CCTCCTTCCT TGGCGGGAAT GTCAGCAGGT TACTGATAGG TCCAGAAACC TCTCCTTCAC ATTGCGCTAG AAGCCTCTCC TTCACATTGC GCTAGTTGTT TGTTTTGATT GATCATCTCT TATGGGTCCC CCTAAGAGCC GCTATTCAAC TCCGCTGGAG CAGTCGCCTT TATGATCCCA TGGTTATCTA TGCAAGCAAG GAGGCCATGC TAGGGTTGAC ATAGTCCTTT ATGGGGTACT ATGTTCAGAG CCGGATCGGC GCAAGTCAGG TATTGGCATC TGAGCCTACT CCCGACCAGG CGACAGATTT GGAACGTACT CTAGGGCCTG CCACGGTTAT ACGGGACGGG GGCAGCCTGC GCCATTAACC CGCTCGCCGT TTCGGGAGGG GGAGTAAAAT TCGCCGCTGT CAGCCTCAAA GCATGTTATC CAGTACATAA ACCGTTACTT GCCCTTAGTC GTGCGCTGCT AGTATACATA AGTGGGGCCG TACTGGCGTT GGTCCCAATG TGCTCGAAGC ACTTCTATTC GTAGTTCCAT GCCTTGTCCG TTTGTATCGCGACGACCAGT ATGCCATATG ATTATGGCAT CAATCCTTTC GTCCCTGCTG CCCTGGTAGATCTTACTTCG TAACGCATCA TGTAAGATGG TCTATCTGCG TTACGGGATA AATTAAAAAA TkAAAAAAAAA AACGTTGGCT TTTTGGGGTG TAGAAGTGAA CTGCTTATAC CAAAGCTATC ACCCACTAAG CTTGACGTAG TTTGGGAAAG CGGAACCGCT GGCGAAATTC GTTATACACG GTTCCCCTAT TCCTATTGGA AAACTTCTCA TGGTAGCGCA GAATCAGCAT GTTAGTGTCC CTTGGGAAGG ATTATTGGAT ACTTCATATC AAACGTAGCG TGGTTAGCTT GCTGTAACCT ACCTCCGACA CGCACAACTC CTTCGGTGTA GAGAATAAGA TTCATTTTGT ACAATGGACT TGACTTAGCT ATTCTTTTCG GAGTTACTTC ACATTGTTGG TTTCTATCTA AAATTCTCAA TTCTTCCATG AACTCATCGA GTTGAGCCTG TTTCCATATT CAGCCGCTCA TAGTTCATCC TGTTGTAGAA GTGCTATTAG CAGACTCTCT TGTTAGGAGT AGCCTTAAAC ACTAAAATTG GAT GAT C TAG GGCCTTTCGA CGACAGTTTG CGACGAATCG AATGACGTAG GCTGTACTTT GAAGAAGACA AACGAACTTA TTTGTGGAAT ACACATTGAT GATTGGTTCG ACGTTCACAG CTTCATGATG CAGAACAGAT GAGCGCATTT CTTCGTTTGT ATCGACGGCC GGAAGCAGTC CCACAGGCCA TCGGTTTTCA GACTCCAAAA GGAATTTTGG TCCATTGAAC CACGCACTGC TAGATTTTAG TGGAGGTTCT TGAGCCCATT TTGTTACGTC AGCAATGTTT GTTTTGATAG AAATCCACTG CCAATTGGAG ATGTTCGTAT GTTCTTGAAT CTCTCCAATT CGAAATGCTA CAAATTGCTT ATAATTCCAC TGATCAGAAC GAATCCAGCT AAAGACCGTT TGGGAATCTG ACCATAGAAA ATAGTTGTGT ATTGGAAGGG AGTGATGTTG TTGAACCGTG CGTAGCAAAC GTGATCCAAA CACCATTTAA ATTCGAGGCG AGGAATCAAT TGGGAAGCCA CCAGAGAAAT CCTCACTTTA CCATCCACCA CTGTCCGGAA GTAAGCAACA CAACCGTAAG CATGCCTGCT TGCATCCATG AAGATGTGCA ACTAAAGTAA CTCGATTGCT GTTGGAAAAG CACCGCCGAT GTAACACGAG GGATTCGAAT ATCTGCTACC CGCGATAACA GACTTGCCCA GCGTTTCCAT TTTGATAGGT TAGCTTCGTC CATTTCTTTG TCCTAGCCAC ATCCCGTTCT CCACAGATCC TAAACGAGTA TTTTGCCGTG AATCGTGAAC GGTGCTAGAA GTCCAAGTGG GTCAAAAAAG CTCATGAAGC AGCTCAAAAC CAACCGTTTC GTTGGTCGGC ATGTACCATC TATAAAAGGT TTTTGTTCCG TGCGATGGTC GGTTATGAAC GAAAAGGTGC CCTAGCTTGA ATTCCAGACT TTTCTAAGGA CTCTTTCGTT CTCCGTTGAT TTGTCGACGT TGAAATGAAC ATCTTGCATT GGATTCCGTT CTCCCAGACT TTGATTAACT TCGACCCATT TTCTCACTTC AAAATTTGCC TTCAGGTGGG TGAAACGCAC CTCTTTCGCA CGATGTATTA CTTCTTCGGT GGTGTTAACA CTATCATAAT ACTCATCAAC GTGATGCTTG TCAATGATAG CAGCCGCCGC CTCTGGATAC TTCGCTGAAA ACTCCTACGC TTTAAGATTT TTAATATAAT GGGCTGAACA AGGGGAGCTG CTGGACCCAA AGGTGGCTAC ATCCATGATG TACTTGTACA CGCTCGGCTC CACGTCTCGG GGATTCTTCC CTAATATGCA TCGCTGGGCT TGCTTGTCGT CTTCACGAAT GCGTAGTTGG TGATACTGGT ACATCTCGCG TATGTAGGCC CCAAAGCCAA TACGACGCTC TCGAAAAAGA CTGAGTACAG CGGTCAGCGG CGTGAGCATA TCTAGGCCTT CTAGACTGTT AAGGGATATA CTATTCCTTC GTCATTGCTA AATTACCGTA TCGTTTTTGC TGATAATCAT TGTAACCAGT GTCAGTGTGA TGTCTAGTGT TGATGAGATT TGTCGCGTTT AGTTTCGTAA ATGTATTGCAAAAAAAAAAA GAAAACGTTT ATAGGTAGGT TAGAGTAAGA TGTCCCGAAA GATCCAACGAACGCAGCCAA CATTGCTTCA CCCTATATCA GGACAATGGC AGTGCCGCTA GTCATCATCAGAGATGCGCT AAACCACACG AATGGGCGGC CATGGGAGTG GCGAAAAGGG GTGGCCATGA TGAGACGGAT TACGTCATGA ACAAGGACGT GGGTCAAAGG CCAAAGGTTC AGGGTTAATG ATCACCAGAA CGACACTTCA TTAGCCTGGT GATCTTTGGA GAAGAAAGTC GTTGAGCTGA GAGTTAAGAG CAAGTGCTGT GAAAAATTGA GTTTAAGCTA AGAAGTTCAA TTAAAAGGAG AGGAGGAAAG TATGAAGGAT TTCTAGACTT CCAGGTAAAT TCCGTCGTTC CAAGTAAACC CCGAACCAAG CCCAATCGCG GAGTACCTAA TACTCGCCAT TTTGCACCCC CTAAGCGCTT GCACAGCCAA TATTGAAGAG CATCCTGGAA ATACCGAAAT TTCTTTCAGT ATTTTCTTGT CAGCATTAAT GTTTGCAGCA TATCAATGAT TTGATACAAA CATTAAAATG GCCAGGCCCA CTGTGCAGAC TTGGGGTTTG AAGAGAATTC AAAAATTAAC GTCGATCAGG TTATTCACGA ATGAATAACA TGATCGACGA AAAAGTTTGA AGTATTGTGA AAGAAGAAAC CGGGACAACC GTACCGACCG TTCCATTTAG GATGATAAGC AAAATCGTTG GGAGAGGCGT TGCTAAGAAA GTTAATGCGC ACTCCATTGT TGTTTTTGGT CCTTTTCCTG GGATGGGACA CATGTATTTC TCCCGTATGT CCTGGCTTCA GAGCCTAGGC TTAAGCGCCC TTACTCGCTC TCTGAAAAGA AAGAAAATCT GACCTCCCTC GCGTTTTGTA GGTTTCTGTG ATATAAAATA GATTCAACCT TTTGTTACAA TATTGAAAAT TGTATCATCC ACTTTATATC TAAAATGCTT GTAAATAAAA AGTGAAAAAC AAATTCTAAT GTTGCTAAAA TTCGAAATGT ATGTGATTGA TAATGGCTCA CCAACATTTG ATCCAAGATA AGTCTTTTTG CAATAAAAAT TTAAAAAAAA ATAAAAAACT GCTTCTTGCA TTACGTATCA TGTTATATGA TTTGTTACGC CATGAGAACC ATTACCTGTA AACGCATAAC ATCCAGCTAG AGAAGCTCTT GAAATCAAAT TTTACGCGCA GTCCAAAAAT TGTCCAAACA GCGTTGACTA TCACTATCAG ATAGCCATTC CGTCTACCAA GACTAATCTT CATCTCCCAT CCAGTCCACG ATGACCTTTC TGCTATAAAA ATATACTGGA AGAGTAGCTC TTGCATGGTT TTCGAAAAGC TCATAAACCA CTTGCATTCA CCGCATCAAA CACAAACCTG ATCACCAATG GACCAATGCA CGAGTTCACT ATATGACGTT TGAGCACTGC ACCCTATTTA TGTGGTCAAA TAACGAGCGA ATATGGCACC GCACAAATGT CAAATGATTG AACTCGTGCA TTGGTCCCTG ACATTCAAGC TATCACAAAC AATGCAAACC CAAATTGAAA CGAACTCACC GGATCCAACC TTTACAAGAA ACCTTCCGTC GTCTCAGAAT TGTTCAAAAG GGATGATGAG CGTGCCGATT ATCCTCCTGT AATGTTAGTT CCTAGTGGAG GCATTGCTGG AGGCAGCCAA CGATTGCTGG AATGTCACAT CCCGCATTGA ACAAACTTGA ATCAACACGA ACTTCTTCAG AATATCCAAA TAAGTGCACA ATCACACCAA CTGTTTTTCA TAAATAAAAT TTCAAAATTT TCACAAAACT GACGGAGCAT AAAATATTTG CTTCCGTTTT AGCCGCCGCA CACTTCGATC CTCTCCCAAA CAATTCCCAC AGGACCTCGC GGTTTTTGTT TAAAAACGCG ATTGCGATTT GGTGTATTAT CACCGTAAAC TACCTGTGTT GACCCTACGC GCCAATTAAG TAAGTCCGTG AACACCTGTA CCCAAGTACG ACAACCGAGT GCGATTGGAA ACCCAAGACT TCCAGTGAAT CCACGGATCA CTGAATACTT GTTTTTAATT AGGAATCGTG GTTTACGGCT AACCAGCCGAGTGGAAGTTT AACAACTACC GAAAAGCTAA ACATTACAAT AGACCTTACA CCCCTTGAAAAAGGCCAACA CCCATCGGCC GAAACGTCGG GAATGTAAAA AACATCGTTT TTCGACTTCCAGACTGAGAA AGCCATCCGA AAGTGGTAGG AGTAGTTGTA AAAAAGTGGG AGGCGTCGTCTGGTGTTGAG GACGGACGCA AGGATAGAGA AGGGAAAGTC AGGAGTAGAA GAAGTAGGAC GTCAAAGAAG TGAACTGCCC ATAACTGCAA AACAGTCACA TTCGACATTT TTGACAAAAT GGAGTTAATA CCATGGAGAG TCATCAAATG ATAAATACTT TCGATCAACT TACTGAAAGC TGTGAGATTG TTCTAGAAAA TTCGAAAAAA ATACCAAGTT GTTTTGTCAC ATTGATAATT ATAACCGCAT AACAGTCACA TTGAGATTAC AAATGGGTCT CGTAATGTGA TAGCAATGAA ATTTCTATCA GAATTATTTT CTGACAATTC ACCTAGTACG CGATATGAGT TGTACAAAAT ATCAGCCTCA AATAAGCACT TTTGAGTTCC TGGTAATTTA TAGAAATTTT AGTTTTCTCC CATACTGCCA TAAAATGCAC ACTTGGTATT CCATTTACTC AATGCCTACT TTTGTCGAAT GTTACAAATA TGCAGTTATG GGCAGTGAAG ATACAAGTGT ACACGAGACC ACGAAATAGA GGGAGTTTTA CTGGAGTGTT TTCTTTCGTG GCGTAGCGCC GACCCTGAGG CTTGGAGACG CGGGTCTCAT CTAGACTGAG AGTAACACGG ACTGCAAGTT ATTTACATCA TCAATTAATT GTTTGAACAA CAGTATGAAG GGCTTATCCA ATCATACAAA GCTTCAAAAT ACAGTTATTG TATGTTTCAA TTCTGATAAG TGTACGAATA TGAATTTGGG TCAATTTTAG ACTAAAATTC AATACATTTC TATTGTTCCA AAGATTAAAG TAAATGGAAA TATCTTCATT ATTGATAGTG TTTGAAATTA GATATGTACA TCATTTGATA GTTTTGACCA ACTTGATATA GAAACAGTGC CCCGTTCAAA GTATGAAATT AAATCACTGT AAGTGTAGCT CTGATATGAATTTTTTTTTTTTGCAGGTTG ATGAGGTATG TTCCGAAGGA GTTAAGCCCG ACGACGTTGA AAAGGCTCTG GCCGAAATAG TAGAAGATGG CTGCTATTAC ACCAACCTGG AAGAGTACAA GAAAGTGGTT AAAGCTAAAA CCGAATCGTT CAAACCATTC GGCGAGAAGG TCGATGAGTT CCAAATAAGT TCCGGAACTG CGGACGGCGG AACTCGAACC TTTGAAGTGT ACGTGAGTGA CGTGAATGAC AAGGAATTCC TAAAGTTCCA CTCCCGTCTT GAGTCGTTCT CGTTCTGGTT CATCGATGCT TTCAGCCGGG TGGAACACGA CCCTTTGTGG TTGTTCTTCA TGGTCTACGA GAAATATTCG AACAACAATA ACGAAACTCG TTATGCTACG GCTGGGTATT TTACCGTTTA TCAGTATTTC TCGTATCCGC AATTCATTAG GCCACGCGTC AGCCAAATTC TGGTGCTGCC TCCATTCCAA AAGCTGGGCA TCGCTTCTCG GTTGATCGAG GAGACAGTGT TCAAATACTT TGTCCCTAAA GAGAACGTTG CCGATATCAC TTACGAACAG CCGACCGATA TCGTCCAGCA TATTCGATCT GTGTCCGATG CCAAGCGCTG TATGACGCTT CCATCGTTTG CCAAGGAGCA TCTGCTGGCT GGCTTTTCGA AGAGTATGCT CAGAGACGCC AAGGAAAAGT TCAAAATCAA TCCCAAACAG TGCCGGGTTA TCTATGAGAT TCTTCGGTTG GGAGTCACCG ATCTTAAAAA TGAAACCGAG TACCGCAAGT ACCGGGTAGA GGTGAAGAAA CGACTGAATC TGAACGACAG TAAGAACCGT CGTGAGCTGA TGCGGCTGCA GAAGAAGGGA GTGGATGTAA GTGGAGCTTT TAGTATCCTT CCGTCGCTTG AGGACAGAAT AGAGCAGCTG CACGCTGCTT ACAAGGTAAG ATTGTTGAAT TCCATTTGAT TCAAACTTCA TAACTGATGT GTTTCTCATA CCTTGCAGGA AGTGGAAAAA GTATACCACC AGGTGTTGAA GAAGATAAAT TCTATTCGCG AGTAAGCAAT GATCCATGTAGATTTCCTGA TACATCTCTC TTTCTAGAAG GAGTCTATTA CATCTCATAT GTTTCTGTTTAAGGGTCCTT TGTTTTAAAT TCTTAATTTG TACTATTAAA CTCTATGTTA TGCGTAAATAAAGCACAAGT GTATGAAAAA TTThe coding sequence of SEQ ID NO:9 is indicated in bold text.

[0100] In some embodiments, the HAT1 sequence is Aedes aegypti HAT1 and comprises a nucleotide sequence of SEQ ID NO:9 or a nucleotide sequence having at least 95% sequence identity to the nucleotide sequence of SEQ ID NO:9. In some embodiments, the HAT1 sequence comprises a nucleotide sequence having at least 70%, 75%, 80%, 85%,90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the nucleotide sequence of SEQ ID NO:9. In some embodiments, the HAT1 sequence is Aedes aegypti HAT1 and comprises the coding sequence of SEQ ID NO:9 or a nucleotide sequence having at least 95% sequence identity to the coding sequence of SEQ ID NO:9. In some embodiments, the HAT1 sequence comprises a coding sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the coding sequence of SEQ ID NO: 9.

[0101] The amino acid sequence of the protein encoded by the Aedes aegypti HAT1(SEQ ID NO:9) is GenBank Accession No. XP_001651817. 1, which is hereby incorporated by reference in its entirety, and is set forth below as SEQ ID NOTO:MTAI SSLQNF VTDALECTQF RLIRDDADFE DESVAFHPEM AHQI FGEQES I FGYRDLQID VCFAASSLDI YFNIKYSKKV DEVCSEGVKP DDVEKALAEI VEDGCYYTNL EEYKKWKAK TESFKPFGEK VDEFQI SSGT ADGGTRTFEV YVSDVNDKEF LKFHSRLESF SFWFIDAFSR VEHDPLWLFF MVYEKYSNNN NETRYATAGY FTVYQYFSYP QFIRPRVSQI LVLPPFQKLG IASRLIEETV FKYFVPKENV ADITYEQPTD IVQHIRSVSD AKRCMTLPSF AKEHLLAGFSKSMLRDAKEK FKINPKQCRV IYEILRLGVT DLKNETEYRK YRVEVKKRLN LNDSKNRREL MRLQKKGVDV SGAFSILPSL EDRIEQLHAA YKEVEKVYHQ VLKKINSIRE

[0102] In some embodiments, the HAT1 sequence is Aedes aegypti HAT1 and encodes an amino acid sequence of SEQ ID NOTO or an amino acid sequence having at least 95% sequence identity to the amino acid sequence of SEQ ID NOTO. In some embodiments, the HAT1 sequence comprises an amino acid sequence having at least 70%, 75%, 80%, 85%, 90%, 95%, 96%, 97%, 98%, or 99% (or any number or range therein) sequence identity to the amino acid sequence of SEQ ID NOTO.

[0103] In some embodiments, the arthropod is genetically modified with a heterologous polynucleotide comprising a promoter expressed during germline development. In some embodiments, this promoter is operably linked to an inhibitory poly nucleotide such as an inhibitory RNA (e.g, an shRNA). In some embodiments, the promoter is expressed duringspermatogenesis. In some embodiments, the promoter is a Nanos (“nos”) promoter. In some embodiments, the promoter is a Vasa or a Maternal Triple Driver (“MTD”) promoter. In some embodiments, a two-part system is used in which the first promoter is a germline specific promoter (e.g., Nanos, Vasa or MTD) expressing a Gal4 or Gal4:VP16 driver and the Gal4 or Gal4:VP16 driver drives expression from an upstream activation sequence to express the inhibitory polynucleotide.

[0104] In some embodiments, a promoter is operably linked to a histone deacetylase to increase expression of the histone deacetylase. In some embodiments, the histone deacetylase is expressed during germline development.

[0105] In some embodiments, the expression of an arthropod histone acetyltransferase is reduced using an inhibitory polynucleotide. In some embodiments expression of the endogenous histone acetyltransferase is reduced by about 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, 90% or 95% (or any number or range therein) or 100% of the expression of endogenous histone acetyltransferase in an arthropod cell that is not transformed with an inhibitory polynucleotide targeting an endogenous histone acetyltransferase. Expression levels of HAT can be determined by any method known in the art, for example by qRT-PCR.

[0106] In some embodiments, the male arthropods comprising an inhibitory polynucleotide targeting a HAT gene have reduced numbers of offspring when mated to a non-endosymbiont infected female or a female with a non-rescuing endosymbiont relative to a female with a rescuing endosymbiont. In some embodiments, when mated to a non- endosymbiont infected female, the male arthropods comprising an inhibitory polynucleotide targeting a HAT gene have a median embryonic hatch rate of about 0%, 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, or 90% (or any number or range therein) of the median embryonic hatch rate when mating with to a female with a rescuing endosymbiont. In some embodiments, viable offspring are reduced by at least 5%, 10%, 15%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, 90%, 95% or 100% (or any number or range therein).

[0107] In some embodiments, the expression or activity of the histone acetyltransferase can be reduced using genome editing. Genome editing is a type of genetic engineering in which DNA is inserted, replaced, or deleted, or any combination thereof, froma genome using artificially engineered nucleases, or “molecular scissors.” The nucleases typically create double-stranded breaks (“DSBs”) at desired locations in the genome and harness the cell’s endogenous mechanisms to repair the induced break by processes of homology dependent repair (“HDR”) or nonhomologous end-joining (“NHEJ”). Any method of genome editing may be used in the embodiments of the present disclosure.

[0108] CRISPR / Cas type RNA-guided endonucleases provide an efficient system for inducing genetic modifications in genomes of many organisms. Non-limiting examples of genome editing nucleases include Casl, CaslB, Cas2, Cas3, Cas4, Cas5, Cas6, Cas7, Cas8, Cas9 (also known as Csnl and Csxl2), CaslO, Casl2a (Cpfl), Csyl, Csy2, Csy3, Csel, Cse2, Cscl, Csc2, Csa5, Csn2, Csm2, Csm3, Csm4, Csm5, Csm6, Cmrl, Cmr3, Cmr4, Cmr5, Cmr6, Csbl, Csb2, Csb3, Csxl7, Csxl4, CsxlO, Csxl6, CsaX, Csx3, Csxl, Csxl5, Csfl, Csf2, Csf3, Csf4, Cpfl, CasX, CasY, Mad7, or homologs, modified versions, and endonuclease inactive versions thereof. An example of a fusion protein to Cas9 is a cytidine deaminase-Cas9 fusion protein used in cytidine base editing to mutate nucleotides in target genes without generating double-strand breaks as described in Komor et al., “Programmable Editing of a Target Base in Genomic DNA without Double-Stranded DNA Cleavage,” Nature 533:420-424 (2016), which is hereby incorporated by reference in its entirety. The use of CRISPR guide RNA in conjunction with CRISPR / Cas technology to target RNA is also described in Wiedenheft et al., “RNA-Guided Genetic Silencing Systems in Bacteria and Archaea,” Nature 482:331-338 (2012); Zhang et al., “Multiplex Genome Engineering Using CRISPR / Cas Systems,” Science 339:819-23 (2013); and Gaj et al., “ZFN, TALEN, and CRISPR / Cas-based Methods for Genome Engineering,” Cell 31 :397-405 (2013), each of which is hereby incorporated by reference in its entirety.

[0109] There are typically two distinct components to a CRISPR system, a guide RNA (“gRNA”) and a genome editing endonuclease. The gRNA uses a CRISPR RNA (“crRNA”) comprising a DNA targeting segment that can be engineered to contain a complementary stretch of nucleotide sequence (e.g, at least 10 nucleotides) to target a DNA site for binding and subsequent modification by CRISPR genome editing nuclease. The length of a crRNA may range from about 15 nucleotides to about 60 nucleotides. The crRNA can be chemically synthesized and can also be engineered to include a ribonucleotide analog or a modified form thereof, or an analog of a modified form, or non-natural nucleosides.

[0110] Depending on the genome editing nuclease used, the gRNA can also comprise a trans-activating crRNA (“tracrRNA”). Such is the case with Cas9, for example. ThetracrRNA is a small RNA sequence that forms a binding handle used by the CRISPR protein. The tracrRNA can be chemically synthesized and can also be engineered to include a ribonucleotide analog or a modified form thereof, or an analog of a modified form, or nonnatural nucleosides.10111] The term gRNA also includes single guide RNAs (“sgRNA”), which combine the targeting specificity of the crRNA with the scaffolding properties of the tracrRNA. sgRNAs can be synthesized or expressed as a continuous RNA transcript. Alternatively, two- part gRNAs can be assembled from two separate RNAs by combining a crRNA-containing RNA with a tracrRNA. In the sgRNA or the two-part gRNA, crRNA and tracrRNA are present either in their native form, or a modified form. Either type of gRNA may be about 60 nucleotides to about 120 nucleotides long. These gRNAs can be chemically synthesized and can also be engineered to include a ribonucleotide analog or a modified form thereof, or an analog of a modified form, or non-natural nucleosides.

[0112] When the gRNA and the gene editing endonuclease are introduced into the cell, the genomic target sequence can be modified or permanently disrupted to create a loss-of- function mutation(s).

[0113] In some embodiments, the expression of an arthropod histone deacetylase is increased by overexpression. In some embodiments expression of the histone deacetylase is increased by about 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, 90%, 100% (or any number or range therein) or greater thanl00% of the expression of endogenous histone deacetylase in an arthropod cell or organism that is not transformed. Expression levels of HD AC can be determined by any method known in the art, for example by qRT-PCR.

[0114] In some embodiments, the male arthropods comprising an inhibitory polynucleotide targeting a HAT gene have reduced numbers of offspring when mated to a non-endosymbiont infected female or a female with a non-rescuing endosymbiont relative to a female with a rescuing endosymbiont. In some embodiments, when mated to a non- endosymbiont infected female, the male arthropods comprising an inhibitory polynucleotide targeting a HAT gene have a median embryonic hatch rate of about 0%, 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, or 90% (or any number or range therein)of the median embryonic hatch rate when mating with to a female with a rescuing endosymbiont.

[0115] In some embodiments, the male arthropod comprising an inhibitory polynucleotide targeting a HAT gene or overexpressing a HD AC gene have reduced histone H3 and / or H4 acetylation levels in sperm in comparison to a wild-type male arthropod. In some embodiments, the male arthropod comprising an inhibitory' polynucleotide targeting a HAT gene or overexpressing a HD AC gene have reduced histone H3 and / or H4 acetylation levels during sperm development in comparison to a non-contacted male arthropod. In some embodiments, histone H3 and / or histone H4 acetylation levels are reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2, 2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold. In some embodiments, the H3 and / or histone H4 acetylation levels are reduced by at least 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, the H3 acetylation levels are reduced by at least 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, H4 acetylation levels are reduced by at least 5%, 10%, 20%, 30%, 40%, 50%, 60%, 70%, 80%, 90%, 100% or any number or range therein. In some embodiments, H3 and / or histone H4 acetylation levels are reduced by at least 20%. In some embodiments, H3 and / or histone H4 acetylation levels are reduced by at least 40%. In some embodiments, histone H3 acetylation is reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2, 2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold. In some embodiments, histone H4 acetylation is reduced by at least 1, 1.1, 1.2, 1.3, 1.4, 1.5, 1.6, 1.7, 1.8, 1.9, 2.0, 2.2, 2.4, 2.6, 2.8, 3, 3.5, 4, 4.5, 5, 6, 7, 8, 9, 10, 20, 30, 40, or 50-fold, or any number or range therein, or more than 50-fold.

[0116] In some embodiments, the histone H2B acetylation levels during sperm development stages are not reduced by the inhibitory polynucleotide targeting a HAT gene or by overexpression of a HD AC gene. In some embodiments, the histone H3 and / or histone H4 acetylation levels are reduced during post-meiotic sperm development by the inhibitory polynucleotide targeting a HAT gene or by overexpression of a HD AC gene. In some embodiments, the histone H2B acetylation levels are not reduced during post-meiotic sperm development by the inhibitory polynucleotide targeting a HAT gene or by overexpression of a HD AC gene.Methods of Controlling Arthropod Populations -Population Suppression

[0117] One aspect of the present disclosure relates to a method of population suppression of arthropods. This method involves contacting a population of male arthropods with ahistone acetyltransferase (HAT) inhibitor, where said contacting (i) decreases the ability of the male arthropods to produce viable offspring upon mating with female arthropods lacking a bacterial endosymbiont in comparison to mating with female arthropods having a rescuing bacterial endosymbiont or in comparison to uncontacted male arthropods; or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with female arthropods having anon-rescuing bacterial endosymbiont in comparison to uncontacted male arthropods; and releasing the population of male arthropods into a target population of arthropods lacking a rescuing bacterial endosymbiont under conditions effective to reduce the target population.

[0118] Another aspect of the present disclosure relates to a method of population suppression of arthropods. This method involves providing a genetically engineered arthropod, said arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod (i) decreases the ability of the male arthropod to produce viable offspring upon mating with a female arthropod lacking a bacterial endosymbiont in comparison to mating with a female arthropod having a rescuing bacterial endosymbiont or in comparison to wild-type male arthropods; or (ii) decreases the ability of the male arthropod to produce viable offspring upon mating with a female arthropod having a non-rescuing bacterial endosymbiont in comparison to wild-type male arthropods; and releasing the genetically engineered arthropod into a target population of arthropods under conditions effective to reduce the target population.

[0119] A method of controlling arthropod populations is the sterile insect technique (SIT) (Knipling, “Possibilities of Insect Control or Eradication Through the Use of Sexually Sterile Males,” J. Econ. Entomol. 48, 459-462 (1955), which is hereby incorporated by reference in its entirety). SIT is the use of sterile males to suppress populations of insects. SIT works by periodic controlled releases of vast numbers of sterile male insects into wild populations. In principle, these sterile males outnumber and outcompete wild males for matings with wild females. If a female mates with a sterile male she will lay eggs that do not hatch. If the proportion of sterile males consistently exceeds the proportion of fertile malesthen each new generation's reproduction is suppressed. As the wild population numbers dwindle, SIT becomes more and more effective creating a negative feedback loop that ultimately eradicates the species in an area. One major advantage of SIT population suppression versus traditional insecticide treatment is that it is species specific and environmentally safe.

[0120] Described herein is the use of a small chemical histone acetyltransferase inhibitor and / or use of genetic approaches to reduce expression of histone acetylase or to increase expression of histone deacetylase to result in reduced histone acetylation during sperm development in arthropod males for use in the methods and compositions described herein. In some embodiments, the methods and compositions of the disclosure are used for application of SIT. In some embodiments, achieving a perfect 100% sterility is not entirely necessary for application of SIT (Dame, et al. “Historical applications of induced stenlization in field populations of mosquitoes,” Malaria J. 8: 1-10 (2009), which is hereby incorporated by reference in its entirety).

[0121] As disclosed herein, male fertility can be modulated by contacting a male arthropod with a HAT inhibitor such that said contacting causes a reduction in the ability of the male arthropods to produce viable offspring with female arthropods lacking a bacterial endosymbiont or with females having a non-rescuing endosymbiont in comparison to female arthropods having a rescuing bacterial endosymbiont. Additional exemplary methods of the disclosure involve genetically engineered arthropods comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod causes a reduction in the ability of the male arthropods to produce viable offspring with arthropod females lacking a bacterial endosymbiont or with females having a non-rescuing endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont.

[0122] In some embodiments, male insects can be separated from female insects, delivered to the target site, and released for mating with wild females to eradicate a pest population. Alterations to the system can be made to optimize effectiveness of the controlled population suppression.

[0123] In the case of the transgenic modifications, the transgene can be used as a tracking marker distinguishing the modified sterile insects from wild insects. Thus, the presence of the inhibitory polynucleotide or overexpression construct would provide anadditional tool to monitor and characterize the spread or incompatibility of the released populations or other populations of insects containing the inhibitory polynucleotide or overexpression construct. These markers can be detected by means such as standard polymerase chain reaction. Furthermore, because the inhibitory polynucleotide or overexpression construct described underlie the reproductive barriers induced by wild strains of Wolbachia in insects, these markers can be used for determining and assessing mating compatibilities of any intraspecies insect strains in general. Thus, commercial testing, research, and reproductive compatibility assessment by characterization of these inhibitory polynucleotides in conjunction with Wolbachia can be used as a pest management tool for agricultural companies seeking to eradicate or monitor the spread of a particular pest.Methods of Controlling Arthropod Populations: Population Replacement

[0124] Another method for controlling pest and disease vector populations is a Population Replacement Strategy (PRS). In some embodiments, the goal of PRS is to replace wild pest or vector populations with those that are not competent to function as pests or vectors of human disease (Sinkins, “ Wolbachia and Cytoplasmic Incompatibility in Mosquitoes,” Insect Biochem. Mol. Biol. 34:723-729 (2004); Dobson et al., “Wolbachia- Based Strategies to Control Insect Pests and Disease Vectors,” AsPac J. Mol. Biol. Biotechnol, 17:55-63 (2009), each of which is hereby incorporated by reference in its entirety).

[0125] In some embodiments, Population Replacement is dependent on two pieces of technology: 1) a beneficial trait that is desired in the target arthropod; and 2) a genetic drive mechanism to spread the desired trait through the arthropod population (Sinkins and Gould, “Gene Drive Systems for Insect Disease Vectors,” Nat. Rev. Genet. 7:427-35 (2006), which is hereby incorporated by reference in its entirety).

[0126] The technology described herein addresses the second problem by utilizing chemical treatment, inhibitory polynucleotides, or overexpression of histone deacetylase to affect histone acetylation that enhances cytoplasmic incompatibility (CI), which is a natural genetic drive mechanism used by various, unrelated bacterial infections (e.g., Wolbachia and Cardinium endosymbionts). The technology disclosed herein is effective without infection of the male arthropod by the endosymbiont.

[0127] Another aspect of the present disclosure relates to a method of population replacement of target arthropods. This method involves contacting a population of male arthropods with a histone acetyltransferase (HAT) inhibitor, where said contacting (i) increases the ability of the male arthropods to produce viable offspring upon mating witharthropod females having a rescuing bacterial endosymbiont in comparison to arthropod females lacking a rescuing bacterial endosymbiont, or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a nonrescuing bacterial endosymbiont or in comparison to uncontacted males. The method further involves either (i) introducing the male arthropods into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, where said introducing replaces the target population lacking the rescuing endosymbiont or (ii) introducing a population of female arthropods having a rescuing bacterial endosymbiont and the male arthropods into a target population of arthropods, where said introducing replaces the target population of arthropods.

[0128] A further aspect of the present disclosure relates to a method of population replacement of target arthropods. This method involves providing a genetically engineered arthropod, the arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase where expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod (i) increases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a rescuing bacterial endosymbiont in comparison to arthropod females lacking a rescuing bacterial endosymbiont, or (ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont or in comparison to wild-type males. The method further involves either (i) introducing the male arthropods into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, where said introducing replaces the target population lacking the rescuing endosymbiont or (ii) introducing a population of female arthropods having a rescuing bacterial endosymbiont and the male arthropods into a target population of arthropods, where said introducing replaces the target population of arthropods.

[0129] In some embodiments, the target population comprises arthropod females lacking a bacterial endosymbiont.Methods of Controlling Arthropod Populations: Genetic Drive

[0130] Genetic drive involves the introduction of a specific trait into a population through the release of insects with a desired trait. These insects carry a “drive” gene that biases inheritance in favor of the desired trait, leading to its rapid spread throughout the population. In some embodiments, the methods and compositions described herein can serveas genetic drivers to propagate desirable genes or traits within targeted arthropod populations by leveraging CI as a mechanism for gene spread.Arthropods and Inf ectious Disease Vectors

[0131] The methods and compositions disclosed herein are useful for control of arthropod (for example, insects) pests and disease vectors, such as mosquitoes transmitting the Dengue fever and Zika viruses.

[0132] In some embodiments, the arthropod is an insect. In some embodiments, the insect is selected from the group consisting of a mosquito, planthopper, rootworm, beetle, cricket, leafhopper, aphid, fly, ant, wasp, cockroach, termite, looper, caterpillar, and moth. In some embodiments, the arthropod is a mosquito or a fly. In some embodiments, the mosquito is selected from the genera consisting of^etfey Culex and Anopheles. In some embodiments, the mosquito is an Aedes mosquito. In some embodiments, the mosquito is an Anopheles mosquito. In some embodiments, the mosquito is a Culex mosquito. In some embodiments, \\Q Aedes mosquito species is selected from the group consisting of Aedes cilbopictus. Aedes aegypti and Aedes polynesiensis . In some embodiments, the Anopheles mosquito species is Anopheles gambiae. In some embodiments, the Culex mosquito species is Culex pipiens. In some embodiments, the male and female arthropods are the same species.

[0133] Since histone acetylation is a key epigenetic histone modification to regulate gene expression and reprogramming of insect spermiogenesis, a similar chemically -induced CI approach can be applied more broadly for population suppression or replacement of diverse insect vectors / pest populations including but not limited to: Aedes aegypti, Anopheles gambiae, and Culex mosquitoes and agriculture pests spanning Nilaparvata lugens, Diabrotica virgifera, Popillia japonica, Anabrus simplex, Empoasca fabae, Aphis glycines, Rhagoletis indifferens, Drosophila suzukii, ants such as Dasymutilla occidentalis, Pseudomyrmex gracilis, cockroaches such as Periplaneta americana, Supella longipalpa, termites such as Infraorder Isoptera, Coptotermes formosanus, Marginitermes hubbardi, cattle dcVRhipicephalus (Boophilus) microplus, caterpillar Chrysodeixis includens), moth (Plutella xylostellaf etc.

[0134] In some embodiments, disclosed herein are methods for controlling or reducing populations of insects that transmit human or veterinary pathogens. In some embodiments, disclosed herein are methods for replacing a population of arthropods that transmit human or veterinary pathogens with a replacement arthropod population that is infected with a genetically modified bacteria (for example Wolbachia) that reduces the ability of the insect totransmit the pathogen. In some embodiments, the pathogen is selected from dengue virus, Zika virus, a malaria parasite (Plasmodium genus) , West Nile virus, yellow fever virus, chikungunya virus, Japanese encephalitis, St. Louis encephalitis and Western and Eastern Equine Encephalitis viruses.

[0135] In some embodiments, disclosed herein are methods for controlling or reducing populations of insects that transmit trypanosomes including African sleeping sickness, Chagas disease, and Nagana. In some embodiments, the pathogen is Trypanosoma cruzi. In some embodiments, the pathogen is Trypanosoma brucei. In some embodiments, the insect is of the genus Glossina. In some embodiments, the insect is Glossina morsitans. In some embodiments, the insect is a Tsetse fly. In some embodiments, the insect is a kissing bug. In some embodiments, the insect is of the genus Rodnius. In some embodiments, the insect is Rhodnius prolixus.

[0136] In some embodiments, disclosed herein are methods for controlling or reducing populations of arthropods that transmit rickettsioses and pathogens within Anaplasmatacea including Rickettsias rickettsii, africae, parkeri, sibirica, conorii, slovaca, peacockii, philipii, rickettsii Hlp2, heilongjiangensis, japonica, montanensis, massiliae, rhipicephali, amblyommii, helvetica, monacensis, buchneri, hoogstralli, felis, akari, australis, canadensis, prowazekii, typhi, bellii. In some embodiments, the arthropod is a tick. In some embodiments, the arthropod is a tick of the genera Amblyomma, Ixodes, or Rhipicephalus . In some embodiments, the disease is epidemic typhus. In some embodiments, the disease is scrub typhus. In some embodiments, the disease is an Ehrlichiosis. In some embodiments, the pathogen is of the genus Ehrlichia. In some embodiments, the pathogen is of the genus Anaplasma. In some embodiments, the pathogen is of the genus Orientia. In some embodiments, the arthropod is a chigger of the genus Leptotrombidium. In some embodiments, the arthropod is a louse of the genus Pediculus. In some embodiments, the arthropod is a flea of the genus Pulex.

[0137] In some embodiments, the disclosure is useful for controlling sandflies that transmit leishmaniasis. In some embodiments, the insect is of the genus Phlebotomus . In some embodiments, the pathogen is of the genus Leishmania. In some embodiments, the pathogen is Leishmania donovani, Leishmania infantum, or Leishmania Chagasi.

[0138] In some embodiments, the insect is of various aphids including: Acyrthosiphon kondoi, Brevicoryne brassicae, Rhopalosiphum maidis, Aphis gossypii, Aphis craccivora, Myzus persicae, Rhopalosiphum padi, Acyrthosiphon pisum, Rhopalosiphum rufiabdominalis,Metopolophium dirhodum, Aphis glycine, Therioaphis trifolii, Lipaphis erysimi, Rhopalosiphum padi.

[0139] In some embodiments, the disclosure is useful for controlling the armyworm agricultural pest species including Leucania convecta, Spodoptera exempta, Spodoptera Mauritia, Spodoptera exigua, Mythimna separate, Leucania stenographer.

[0140] In some embodiments, the disclosure is useful for controlling pests of beans and beets. In some embodiments, the insect is either the Bean fly (Ophiomyia phaseoil), the Bean leafroller (Omiodes diemenalis). the Bean looper or Mocis (Mods alterna), the Bean podborer (Maruca vitrata), the Bean spider mite (Tetranychus ludeni), the Beet webworm (Spoladea recurvalis), the Large Brown bean bug (Riptortus serripes), the Small Brown bean bug (Melanacanthus scutellaris).

[0141] In some embodiments, the disclosure is useful for controlling the Blue oat mite (Penthaleus major). In some embodiments, the disclosure is useful for controlling the Brown flea beetle (Chaetocnema sp.). In some embodiments, the disclosure is useful for controlling the Brown mirid (Creontiades pacificus). In some embodiments, the disclosure is useful for controlling the Brown shield bug (Dictyotus caenosus). In some embodiments, the disclosure is useful for controlling the Brown wheat mite (Petrobia latens). In some embodiments, the disclosure is useful for controlling the Bruchid, Cowpea (Callosobruchus maculatus).

[0142] In some embodiments, the disclosure is useful for controlling pests of Com including: the Com aphid (Rhopalosiphum maidis), and the Com earworm (Helicoverpa armigera).

[0143] In some embodiments, the disclosure is useful for controlling pests of cotton including the Cotton aphid (Aphis gossypii), Cotton bollworm Helicoverpa armigera), the Cotton harlequin bug (Tectocoris diophthalmus), the Cotton leafhopper (Amrasca terraereginae), the Cotton leafperforator (Bucculatrix gossypii), the Cotton looper (Anomis flava), the Cottonseed bug (Oxycarenus luctuosus), the Cotton seedling thrip (Thrips tabaci),the Cotton tipworm (Crocidosema plebejana), and the Cotton webspinner (Achyra affinitalis).

[0144] In some embodiments, the disclosure is useful for controlling the Diamondback moth (Plutella xylostella). In some embodiments, the disclosure is useful for controlling the Dried fruit beetle (Carpophilus spp.). In some embodiments, the disclosure is useful for controlling the Eastern false wireworm (Pterohelaeus spp.). In some embodiments, the disclosure is useful for controlling the Etiella moth (Etiella behrii). In some embodiments, thedisclosure is useful for controlling the False wireworm (Pterohelaeus and Gonocephalum spp.). In some embodiments, the disclosure is useful for controlling the Flea beetles, Brown and Redheaded (Chaetocnema and Nisosira sp.). In some embodiments, the disclosure is useful for controlling the Flower beetle (Carpophilus spp .

[0145] In some embodiments, the disclosure is useful for controlling various Grasshoppers and locusts including the Grasshopper, Wingless (Phaulacridium vittatum), the Locust, Australian plague (Chortoicetes terminifera), the Locust, Migratory (Locusta migratori ), the Locust, Yellow- winged (Gastrimargus musicus), the Locust, Spur-throated (Austracris (Noamdacris) guttulos ).

[0146] In some embodiments, the disclosure is useful for controlling the Greenhouse whitefly (Trialeurodes vaporariorum). In some embodiments, the disclosure is useful for controlling the Green peach aphid (Myzus persicae). In some embodiments, the disclosure is useful for controlling the Green mirid (Creontiades dilutus). In some embodiments, the disclosure is useful for controlling the Green vegetable bug (Nezara viriduld). In some embodiments, the disclosure is useful for controlling the Green stink bug (Plautia af finis). In some embodiments, the disclosure is useful for controlling the Grey cluster bug (Nysius clevelandensis). In some embodiments, the disclosure is useful for controlling the Helicoverpa species (armigera and punctigera).

[0147] In some embodiments, the disclosure is useful for controlling planthoppers. In some embodiments, the insect is the small brown planthopper (Laodelphax striatellus). In some embodiments, the disclosure is useful for preventing the transmission of crop diseases like Rice White Stripe Virus. In some embodiments, the disclosure is useful for controlling vectors of plant pathogens.

[0148] In some embodiments, the disclosure is useful for controlling the Jassids and various leafhoppers including the Leafhopper, cotton (Amrasca terraereginae), the Leafhopper, lucerne (Austroasca alfalfas), the Leafhopper, maize (Cicadulina bimaculata), the Leafhopper, vegetable (Austroasca viridigrisea).

[0149] In some embodiments, the disclosure is useful for controlling the Loopers including the Looper, Brown pasture (Ciampa arietaria), the Looper, Castor oil (Achaea janata), the Looper, Cotton (Anomis flava), the Looper, Sugarcane (Mods frugalis), the Looper, Soybean (Thysanoplusia orichalced), the Looper, Tobacco (Chrysodeixis argentifera), the Looper, Vegetable (Chrysodeixis eriosoma).

[0150] In some embodiments, the disclosure is useful for controlling various Thrip pests including the Onion Thrip (Thrips tabaci), the Cotton seedling Thrip (Thr ps tabaci), the Maize Thrip (Frankliniella williamsi), the Plague Thrip Thrips imaginis), the tobacco Thrip (Thrips tabaci), the Tomato Thrip (Frankliniella schultzei), the Western flower Thrip (Frankliniella orientalis) In some embodiments, the disclosure is useful for controlling various Mite pests including the Mite, Bean spider (Tetranychus ludeni), Mite, Brown wheat (Petrobia latens), Mite, Blue oat (Penthaleus major), Mite, Peanut (Paraplonobia spp.), Mite, Redl egged earth (Halotydeus destructor), Mite, Strawberry spider (Tetranychus Iambi), and the Two-spotted mite (Tetranychus urticae).

[0151] In some embodiments, the disclosure is useful for controlling various whitefly pests including the Greenhouse whitefly (Trialeurodes vaporariorum), the Silverleaf whitefly (Bemisia tabaci biotype B and Australian native AN), and the Silverleaf whitefly (Bemisia tabaci biotype Q).

[0152] In some embodiments, the present disclosure is useful for controlling various fruit pests. In some embodiments, the arthropod is from the genera Drosophila. In some embodiments, the arthropod is Drosophila suzukii. Drosophila suzukii, commonly called the spotted-wing drosophila, is a vinegar fly closely related to Drosophila melanogaster . Unlike its vinegar fly relatives who are primarily attracted to rotting or fermented fruit, D. suzukii attacks fresh, ripe fruit by laying eggs under the soft skin. The larvae hatch and grow in the fruit, destroying the fruit's commercial value. The pest particularly (but not limited to) infests cherries, apples, apricots, persimmons, tomatoes, blueberries, grapes, nectarines, pears, plums, peaches, figs, raspberries and strawberries. Although D. suzukii is native to Southeast Asia, the fruit pest has recently invaded North and Central America as well as Europe, where it is expanding rapidly. Effective management of this pest is a challenge owing to the wide host range and short generation time. Therefore, monitoring and controlling D. suzukii is of great economic importance. However, traps and baits containing for instance apple cider vinegar, which are typically used for attracting vinegar flies such as D. melanogaster, are less efficient for attracting and trapping D. suzukii. In some embodiments the fly is D. simulans. In some embodiments, the insect is the Mexican Fruit Fly (Anastrepha ludens). In some embodiments, the insect is the Mediterranean Fruit Fly (Ceratitis capitata). In some embodiments, the insect is of the genus Anastrepha, Bactrocera, or Ceratitis. In some embodiments, the insect is a tephritid. In some embodiments, the disclosure is useful for controlling various other agricultural pests including: the red-houldered leaf beetle (Monolepta australis), Nativebudworm (Helicoverpa punctigera), Native whitefly (Bemis ia tabaci), Northern armyworm (Mythimna separata), Oat aphid (Rhopalosiphum padi), Onion thrip (Thrips tabaci), Pale cotton stainer bug (Dysdercus sidae), Pea aphid (Acyrthosiphon pisum), Pea blue butterfly (Lampides boeticus), Peanut mite (Paraplonobia sppl), Peanut scarab (Heteronyx sppl), Pea weevil (Bruchus pisorum), Pinkspotted bollworm (Pectinophora scutigera), Plague thrip (Thrips imaginis), Podsucking bugs (Nezara viridula), Redbanded shield bug (Piezodorus oceanicus , Redheaded flea beetle (Nisotra sp.), Redlegged earth mite (Halotydeus destructor), Redshouldered leaf beetle (Monolepta australis), Rice root aphid (Rhopalosiphum ruflabdominalis), Rose grain aphid (Metopolophium dirhodum), Rough bollworm (Earias huegeliana), Rutherglen bug (Nysius vinitor), Seed harvesting ants (Pheidole spp.), Scarab, Black sunflower (Pseudoheteronyx sp.), Scarab, Peanut (JPG, 20.4KB) (Heteronyx sp.), Shoot flies (Atherigona sp.), Silverleaf whitefly (Bemisia tabaci biotype B and Australian native AN), Silverleaf whitefly (Bemisia tabaci biotype Q), Sitona weevil (Sitona discoideus), Solenopsis mealybug (Phenacoccus solenopsis), Sorghum midge (Stenodiplosis sorghicola), Sorghum head caterpillar (Cryptoblabes adoceta), Soybean leafminer (Porphyrosela aglaozona), Soybean looper (Thysanoplusia orichalcea), Soybean moth (Aproaerema simplexella), Spotted alfalfa aphid (Therioaphis trifolii), Spur-throated locust (Austracris (Nomadacris) guttulosa), Strawberry spider mite (Tetranychus Iambi , Swarming leaf beetle (Rhyparida spp.), Tortrix (Epiphyasa postvittana), True wireworm (Agrypnus spp.). Vegetable weevil (Listroderes difficilis). Weed web moth (Achyra affinitalis), Whitegrub (Heteronyx spp. , Wingless cockroaches (Calolampra spp.), Wireworm, False (Pterohelaeus and Gonocephalum spp.), Wireworm, True (Agrypnus spp.), Yellow peach moth (Conogethes punctiferalis). In some embodiments, the insect is Heteronychus arator. In some embodiments, the insect is of the genus Amnemus. In some embodiments, the insect is of the genus Pheidole. In some embodiments, the disclosure is useful for controlling the Black field cricket (Teleogryllus commodus, T. oceanicus, Lepidogryllus parvulus), the Black field earwig (Nala lividipes), the Black leaf beetle (Rhyparida nitida), the Black sunflower scarab (Pseudoheteronyx sp.). In some embodiments, the disclosure is useful for controlling the Cowpea bruchid (Callosobruchus macidatus). In some embodiments, the disclosure is useful for controlling the Cricket, Black field (Teleogryllus commodus, T. oceanicus, Lepidogryllus parvulus). In some embodiments, the disclosure is useful for controlling the Crop mirid (Sidnia kinbergi). In some embodiments, the disclosure is useful for controlling the Cutworm (Agrotis spp.). In some embodiments, the disclosure is useful for controlling the Cabbagemoth (Plutella xylostella). In some embodiments, the disclosure is useful for controlling the Castor oil looper (Achaeajanata). In some embodiments, the disclosure is useful for controlling the Click beetle (Agrypnus spp.). In some embodiments, the disclosure is useful for controlling the Clover spnngtail (Sminthurus viridis) In some embodiments, the disclosure is useful for controlling the Cluster caterpillar (Spodoptera liturd). In some embodiments, the disclosure is useful for controlling the Cockroach, Wingless (Calolampra spp. . In some embodiments, the disclosure is useful for controlling the Common grass blue butterfly (Zizina labradus). In some embodiments, the disclosure is useful for controlling the Legume webspinner (Omiodes diemenalis). In some embodiments, the disclosure is useful for controlling the Light brown apple moth (Epiphyas postvittana). In some embodiments, the disclosure is useful for controlling Mods trifasdata. In some embodiments, the disclosure is useful for controlling Pantydia spp . In some embodiments, the disclosure is useful for controlling the Lucerne crownborer (Zygrita diva). In some embodiments, the disclosure is useful for controlling the Lucerne flea (Sminthurus viridis). In some embodiments, the disclosure is useful for controlling the Lucerne leafhopper (Austroasca alfalfae). In some embodiments, the disclosure is useful for controlling the Lucerne leafroller (Merophyas divulsana). In some embodiments, the disclosure is useful for controlling the Lucerne seed wasp (Bruchophagus roddi). In some embodiments, the disclosure is useful for controlling the Lucerne seed web moth (Etiella behrii).

[0153] In some embodiments, the disclosure is useful for controlling forestry and wildlife pests such as the emerald ash borer. In some embodiments, the insect is of the genus Agrilus or specifically Agrilus planipennis. In some embodiments, the disclosure is useful for pests of trees and lumber.

[0154] In some embodiments, the arthropods contacted with a HAT inhibitor are used to produce male arthropods to control and / or to replace a population of target arthropods (for example, replacement of a target population with arthropods that are less susceptible to infectious agents or have a reduced capacity to transmit an infectious agent (for example, dengue virus or Zika virus). Exemplary infectious agents encompass a broad range of pathogens, including but not limited to, bacteria such as Borrelia burgdorferi, which causes Lyme disease and Rickettsia rickettsia which causes Rocky Mountain Spotted Fever; viruses like Dengue, Zika, and Chikungunya, West Nile Virus, and Eastern Equine Encephalitis (EEE); and parasites such as Plasmodium species responsible for malaria.

[0155] In some embodiments, the arthropods include, but are not limited to, Aedes aegypti, Anopheles gambiae, Culex mosquitoes and agriculture pests spanning Nilaparvata lugens, Diabrotica virgifera, Popillia japonica, Anabrus simplex, Empoasca fabae, Aphis glycines, Rhagoletis indifferens, Drosophila suzukii, ants such as Dasymutilla occidentalis, Pseudomyrmex gracilis, cockroaches such as Periplaneta americana, Supella longipalpa, termites such as Infraorder Isoptera, Coptotermes formosanus, Marginitermes hubbardi, cattle tick Rhipicephalus (Boophilus) microplus, caterpillar (Chrysodeixis include ns). and moth (Plutella xylostella), etc.

[0156] In some embodiments, the arthropods include, but are not limited to Aedes aegypti, Anopheles gambiae, Culex, Nilaparvata lugens, Diabrotica virgifera, Popillia japonica, Anabrus simplex, Empoasca fabae, Aphis glycines, Rhagoletis indifferens, Drosophila suzukii, Dasymutilla occidentalis, Pseudomyrmex gracilis, Periplaneta americana, Supella longipalpa, Infraorder Isoptera, Coptotermes formosanus, Marginitermes hubbardi, Rhipicephalus (Boophilus) microplus, Chrysodeixis includens, and Plutella xylostella.Long Noncoding RNA Control of CI

[0157] Another aspect of the present disclosure relates to a genetically modified arthropod. The arthropod comprises a bacterial endosymbiont encoding a cytoplasmic incompatibility factor, a promoter operably linked to an inhibitory RNA molecule targeting a long non-coding RNA, where expression of the inhibitory RNA molecule in a male arthropod causes a reduction in fertility and / or the ability of the male arthropod to produce viable offspring in comparison to an endosymbiont-infected male arthropod lacking the inhibitory RNA molecule.

[0158] Another aspect of the invention relates to a method for controlling a population of target arthropods. This method involves providing a genetically modified arthropod comprising a bacterial endosy mbiont encoding two cytoplasmic incompatibility factors and a promoter operably linked to an inhibitory RNA molecule targeting a long non-coding RNA, where expression of the inhibitory RNA molecule in a male arthropod causes a reduction in fertility and / or the ability of the male arthropod to produce viable offspring in comparison to a male arthropod lacking the inhibitory RNA molecule. A genetically modified male arthropod is released amongst a population of target arthropods under conditions effective to reduce the population of target arthropods.

[0159] A further aspect of the present disclosure relates to a method for replacing a population of target arthropods. This method involves providing a genetically modified replacement arthropod comprising a promoter operably linked to an inhibitory RNA molecule targeting a long non-coding RNA. The replacement arthropod is infected with a bacterial endosymbiont encoding cytoplasmic incompatibility factors, and the replacement arthropod is released amongst a population of target arthropods, where said release of the replacement arthropod reduces the population of target arthropods.

[0160] As described herein, cytoplasmic incompatibility factor proteins CifA and CifB, encoded by prophage WO of the endosymbiont Wolbachia, alter long non-coding RNA (IncRNA) and DNA during sperm development to establish a paternal-effect embryonic lethality known as cytoplasmic incompatibility (CI). CifA is a ribonuclease (RNase) that depletes a spermatocyte IncRNA important for the histone-to-protamine transition of spermiogenesis. Both CifA and CifB are deoxyribonucleases (DNases) that elevate DNA damage in late spermiogenesis. IncRNA knockdown using genetic modification enhances CI, and mutagenesis links IncRNA depletion and subsequent sperm chromatin integrity changes to embryonic DNA damage and CI. Hence, prophage proteins interact with eukaryotic macromolecules during gametogenesis to create a symbiosis that is fundamental to insect evolution and vector control.

[0161] Also described herein, is the enhancement of CI using a combination of Wolbachia infection and IncRNA-engineering in arthropod genomes to provide a versatile and sustainable approach for population suppression, replacement, and trait propagation in arthropod populations. Central to this innovation is the regulated expression of IncRNA at a specific stage of spermiogenesis.

[0162] Mills et al., “RNA from a Simple-Tandem Repeat is Required for Sperm Maturation and Male Fertility in Drosophila melanogasterf eLife 8:e48940 (2019), which is hereby incorporated by reference in its entirety) used the Bag of marbles (Bam)-GAL4:VP16 driver to express AAGAG short hairpin (sh)RNA and deplete AAGAG IncRNA in spermatocytes in males devoid of Wolbachia. A 72% reduction in AAGAG IncRNA expression resulted in 100% male sterility with abnormal histone-to-protamine transition resulting in no mature sperm production, limiting this approach for application in population suppression or replacement. However, it was surprisingly shown herein that expressing AAGAG shRNA earlier in germline and spermatogonia using a weaker nos-Gal4:VP16 driverdid not make males fully sterile. This discovery enables new applications of population suppression or replacement as described herein.

[0163] The relevance of AAGAG IncRNA-mediated partial male sterility with Wolbachia-induced CI has not been tested before. InZ). melanogaster , wild type CI levels induced by ‘wMel’ strain of Wolbachia are -70% embryonic lethality. Notably, the IncRNA- mediated sterility and Wolbachia-conierred CI both impair the histone-to-protamine transition of sperm development. As described herein, knocking down IncRNA at a moderate level (less than 72%) in the presence of wMel should increase wild type CI levels by increasing embryonic lethality.

[0164] By integrating IncRNA transgene with Wolbachia infection in transgenic arthropods, the present disclosure facilitates applications including Controlled Population Suppression, Population Replacement, and Genetic Drive for Trait Propagation as discussed supra.

[0165] In Controlled Population Suppression, transgenic arthropods engineered with Wolbachia-targeted, down-regulated IncRNA can be deployed in controlled releases to rapidly suppress pest populations. The augmented CI induced by the Wolbachia and knocked-down IncRNA enhances the effectiveness of the “incompatible insect technique”, leading to a rapid suppression of arthropod vector population.

[0166] In Population Replacement with long noncoding RNA, Wolbachia-cwying transgenic arthropod males, equipped with Wolbachia-tergeted, down-regulated IncRNA, will effectively mitigate CI -induced embryonic mortality upon mating with non-engineered IFo / Z>oc / zza-carrying arthropod females in natural habitats. This mating strategy ensures the inheritance of maternally -transmitted Wolbachia in the offspring. Consequently, through sustained high levels of CI, the wild arthropod vector population will be rapidly replaced by Wolbachia-carrying individuals, thereby diminishing their disease transmission capability.

[0167] In Genetic Drive for Trait Propagation, the transgenic arthropods can serve as genetic drivers to propagate desirable genes or traits within targeted arthropod populations by leveraging CI as a mechanism for gene spread.

[0168] In some embodiments, the compositions and methods disclosed herein provide a paradigm shift in controlling commercially valuable arthropod insects and pest species, offering a comprehensive and integrated approach that leverages both microbial and host genetic factors to enhance CI, mitigating the risk of CI 'aning and ensuring its continued efficacy.

[0169] Also disclosed herein is the use of long non-coding RNA targets of the CI genes to genetically modify arthropods in order to modulate sterility of male arthropods and / or to replace a population of target arthropods

[0170] In some embodiments, the arthropod endosymbiont comprises a cytoplasmic incompatibility factor that targets long non-coding RNAs. In some embodiments, the endosymbiont is from the genus Wolbachia. In some embodiments, the endosymbiont is from the genus Cardinium.

[0171] In some embodiments, the genetically modified arthropods are used to produce male arthropods to control and / or to replace a population of target arthropods (for example, replacement of a target population with arthropods that are less susceptible to infectious agents or have a reduced capacity to transmit an infectious agent (for example, dengue vims or Zika virus). Exemplary infectious agents are discussed supra and encompass a broad range of pathogens, including but not limited to, bacteria such as Borrelia burgdorferi, which causes Lyme disease and Rickettsia rickettsia which causes Rocky Mountain Spotted Fever; viruses like Dengue, Zika, and Chikungunya, West Nile Virus, and Eastern Equine Encephalitis (EEE); and parasites such as Plasmodium species responsible for malaria.

[0172] Long non-coding RNAs (IncRNA) are transcribed from DNA tandem repeats (satellites) that are present in most eukaryotic species and are frequently expressed as RNA transcripts. In Drosophila melanogaster , AAGAG is one of the most abundant simple repeats, comprising ~5% of the genome (Lohe and Brutlag, “Multiplicity of Satellite DNA sequences in Drosophila melanogaster f Proc. Natl. Acad. Sci. USA 83(3):696-700 (1986), which is hereby incorporated by reference in its entirety). High levels of long non-coding RNA transcribed from the AAGAG repeat are expressed in primary spermatocytes originating from the 2R, 3R and X heterochromatic satellite regions and is specifically not generated from the Y chromosome. AAGAG IncRNA expression is necessary for completing spermatogenesis and male fertility in Drosophila melanogaster, in part by promoting the histone-protamine transition during sperm development.

[0173] Nucleic acid molecules that reduce abundance of long non-coding RNAs include inhibitory RNAs. As discussed supra, non-limiting examples of inhibitor RNAs include antisense RNAs or RNAi, such as short interfering RNAs (siRNA), short hairpin RNAs (shRNA), and micro interfering RNAs (miRNA).

[0174] In some embodiments, an arthropod is genetically modified to comprise a heterologous polynucleotide encoding an inhibitory RNA targeting a long non-coding RNA.In some embodiments, the inhibitory polynucleotide targets a long non-coding RNA comprising AAGAG(n) repeats. In some embodiments, the number (n) of AAGAG repeats in the long noncoding RNA is 32. In some embodiments, the inhibitory' polynucleotide targets a Scrambled long non-coding RNA. In some embodiments the IncRNA is Drosophila melanogaster IncRNA.

[0175] In some embodiments, the inhibitory polynucleotide comprises repeats of AAGAG. In some embodiments, the inhibitory polynucleotide comprises at least two repeats of AAGAG. In some embodiments, the inhibitory polynucleotide comprises at least three repeats of AAGAG. In some embodiments, the inhibitory polynucleotide comprises at least four repeats of AAGAG. In some embodiments, the inhibitory polynucleotide comprises 21 nucleotides of a repeat of any part of AAGAG.

[0176] In some embodiments, the inhibitory polynucleotide rs an shRNA. An exemplary shRNA inhibitory polynucleotide used to reduce expression of an AAGAG(n) long non-coding RNA is SEQ ID NO: 11, as shown below. The capital letters in SEQ ID NO: 11 and 12 indicate the sequence directed to the target long non-coding RNA and its complementary sequence, which form a double-stranded RNA when expressed in the arthropod. shRNA sequence to AAGAG(n) (SEQ ID NO: 11): ctagcagtGAAGAGAAGAGAAGAGAAGAGtagttatattcaagcataCTCTTCTCTTCTCTTCTCT TCgcgIn some embodiments, the sequence of the inhibitory polynucleotide comprises SEQ ID NOT E

[0177] An exemplary shRNA inhibitory polynucleotrde used to reduce expression of a Scrambled long non-coding RNA is SEQ ID NO: 12 as follows: shRNA sequence to Scrambled (SEQ ID NO: 12): ctagcagtGAGAGAAAAAGGGAAAGAAGGtagttatattcaagcataCCTTCTTTCCCTTTTTCTC TCgcgIn some embodiments, the sequence of the inhibitory polynucleotide comprises SEQ ID NO:12.

[0178] Other arthropod long non-coding RNAs are suitable for use in the embodiments of the present disclosure. Such long non-coding RNAs can be isolated from developmental tissue (such as cells undergoing spermatogenesis) from arthropods and sequenced using standard techniques. shRNA can be designed to the sequence of these longnon-coding RNAs. Besides Drosophila, IncRNAs role in reproduction have also been reported in a range of diverse insects spanning the orders Diptera, Lepidoptera, Coleopteria, and Hymenoptera. Aedes albopictus (Belavilas-Trovas et al., “A Species-Specific IncRNA Modulates the Reproductive Ability of the Asian Tiger Mosquito,” Front. Bioeng. Biotechnol. 10:8857672022, which is hereby incorporated by reference in its entirety), Bombyx mori (Wu et al., “Systematic Identification and Characterization of Long Non-Coding RNAs in the Silkworm, Bombyx mori,” PLoS ONE 11(1): e01471472016 (2016), which is hereby incorporated by reference in its entirety), Zeugodacus cucurbitae (Li et al., “Inc94638 is a Testis-Specific long non-coding RNA Involved in Spermatozoa Formation in Zeugodacus cucurbitae (Coquillett),” Insect Mol. Biol. 30, 605-614 (2021), which is hereby incorporated by reference in its entirety) and in mammals (Sahlu et al., “Long Noncoding RNAs: New Insights in Modulating Mammalian Spermatogenesis,” Anim. Sci. Biotechnol. Feb 28; 11 : 16 (2020), which is hereby incorporated by reference in its entirety).

[0179] Arthropod pests are discussed supra. For example, Aedes albopictus is a vector of dengue and chikungunya, and Z. cucurbitae is an agricultural pest attacking crops such as cucurbits and tomatoes. Furthermore, stable introduction and spread of Wolbachia into the brown planthopper, Nilaparvata lugens enabled by high levels of CI is proposed to block the vector-mediated rice virus transmission and limit the crop damage. Thus, Wolbachia and IncRNA mediated augmented CI can be useful in this context.

[0180] Since IncRNAs are important to regulate the gene expression and epigenetic reprogramming of insect spermiogenesis, a similar IncRNA depletion approach can be applied more broadly for population suppression or replacement of diverse insect vectors / pest populations including but not limited to: Aedes aegypti, Aedes albopictus, Anopheles gambiae, and Culex mosquitoes and agriculture pests spanning Nilaparvata lugens, Diabrotica virgifera, Popillia japonica, Anabrus simplex, Empoasca fabae, Aphis glycines, Rhagoletis indifferens, Drosophila suzukii, and Z. cucurbitae as non-limiting examples.

[0181] The following mosquito species share 100% similarity to the Drosophila AAGAG IncRNA repeat sequence: Anopheles gamble, Anopheles stephensi, Anopheles ziemanni, Anopheles arabiensis, Anopheles coluzzi, Anopheles coustani, Anopheles aquasalis, Culex pipiens, and Culex quinquefasciatus .

[0182] In many organisms (e.g., humans, mice, and insects such as Drosophila, mosquitoes), the switch from a histone- to a protamine-based chromatin structure is a crucial characteristic feature of sperm maturation. Since Wolbachia inhabit -50% of the arthropodspecies, the disclosure relates to transgenic method of augmented CI upon intermediate knockdow n (reduced expression) of this IncRNA in a Wolbachia-infected target species. In some embodiments, knock down of IncRNA expression is targeted to IncRNA(s) involved in the histone-to-protamine transition of arthropod sperm development.

[0183] In some embodiments, the expression of an arthropod long non-coding RNA is reduced using an inhibitory polynucleotide. In some embodiments expression of the endogenous long non-coding RNA is reduced by about 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, 90% or 95% (or any number or range therein) or 100% of the expression of endogenous long non-coding RNA in an arthropod cell that is not transformed with an inhibitory polynucleotide targeting an endogenous long non-coding RNA. Expression levels of IncRNA can be determined by any method known in the art, for example by qRT-PCR.

[0184] As shown herein (e.g, FIGs. 15A-D), to decouple the AAGAG relevance to CI from male sterility, the nos-Ga!4:VP16 driver was used to drive expression of a short hairpin RNA (“shRNA”) directed to AAGAG and knockdown AAGAG RNA in both wMel-infected (AAGAG_KD+) and uninfected (AAGAG_KD-) wild type lines. Strikingly, it was found that wMel-carrying AAGAG_KD+ males induced approximately three-fold more CI (median embryonic hatching = 11.8%) upon matings with uninfected (wMel-) females compared to control males (Scramble_KD+) with a randomized A and G content that induced intermediate levels of CI (median embryonic hatching = 35.6%) (FIGs. 14A-B). These findings supported a causal interaction between Wolbachia and AAGAG IncRNA depletion on CI.

[0185] In some embodiments, the male arthropods comprising an endosymbiont and an inhibitory polynucleotide targeting a long non-coding RNA have reduced numbers of offspring when mated to a non-endosymbiont infected female relative to non-endosymbiont infected males or relative to endosymbiont infected males not having an inhibitory polynucleotide targeting a long non-coding RNA when mated to a non-endosymbiont infected female. In some embodiments, when mated to a non-endosymbiont infected female, the male arthropods comprising an endosymbiont and an inhibitory polynucleotide targeting a long non-coding RNA have a median embryonic hatch rate of about 1%, 2%, 3%, 4%, 5%, 6%, 7%, 8%, 9%, 10%, 11%, 12%, 13%, 14%, 15%, 16%, 17%, 18%, 19%, 20%, 25%, 30%, 35%, 40%, 45%, 50%, 55%, 60%, 70%, 80%, or 90% (or any number or range therein) of the median embryonic hatch rate of non-endosymbiont infected males or of the median embryonichatch rate of endosymbiont infected males not having an inhibitory polynucleotide targeting a long non-coding RNA.Methods of Controlling Arthropod Populations Using Long Noncoding RNA

[0186] As discussed supra, in some embodiments, one method of controlling arthropod populations is the sterile insect technique (SIT) (Knipling, “Possibilities of Insect Control or Eradication Through the Use of Sexually Sterile Males,” J. Econ. Entomol. 48:459- 462 (1955), which is hereby incorporated by reference in its entirety).

[0187] Three major processes are necessary for the implementation of SIT: 1) a method of sterilization; 2) a method of sex separation; and 3) a method of dispersal. In some embodiments, the present disclosure relates to the first point and represents a unique method of controlling male fertility. In some embodiments, male fertility can be modulated by reducing the levels of long non-coding RNA targeted by the endosymbiont CI mechanism. A range of levels of long non-coding RNA can be achieved (e.g., by varying the promoter used to drive the inhibitory polynucleotide targeting the long non-coding RNA) with greater reductions in IncRNA further increasing CI.

[0188] Also described herein is the use of an endosymbiont’s cytoplasmic incompatibility genes with an inhibitory polynucleotide targeting a long non-coding RNA transgenically inserted into a pest insect for the purposes of controlled population suppression. In some embodiments, the inhibitory polynucleotide is used to induce high levels of CI nearing 90-100% sterility in D. melanogaster . In some embodiments, the inhibitory polynucleotide is used for application of SIT. Achieving a perfect 100% sterility is not entirely necessary for application of SIT (Dame et al., “Historical Applications of Induced Sterilisation in Field Populations of Mosquitoes,” Malaria J. 8 (2009), which is hereby incorporated by reference in its entirety).

[0189] After preparation, male insects could then be separated from female insects, delivered to the target site, and released for mating with wild females to eradicate a pest population. Alterations to the system can be made to optimize effectiveness of the controlled population suppression. As mentioned previously, these changes might include alterations of gene regulatory sequences driving the inhibitory polynucleotide production.

[0190] The unique difference with the methods disclosed herein is the fact that the targeted expression of the inhibitory polynucleotide effectively inhibit embryonic development and hatching of eggs. Evidence for this is provided, e.g., in FIG. 15C.

[0191] Once released, the transgenic insertion can be used as a tracking marker distinguishing the modified sterile insects from wild insects. Thus, the presence of the inhibitory polynucleotide would provide an additional tool to monitor and characterize the spread or incompatibility of the released populations or other populations of insects containing the inhibitory polynucleotide. These markers can be detected by means such as standard polymerase chain reaction. Furthermore, because the inhibitory polynucleotide described underlie the reproductive barriers induced by wild strains of Wolbachia in insects, these markers can be used for determining and assessing mating compatibilities of any intraspecies insect strains in general. Thus, commercial testing, research, and reproductive compatibility assessment by characterization of these inhibitory polynucleotides in conjunction with Wolbachia can be used as a pest management tool for agricultural companies seeking to eradicate or monitor the spread of a particular pest.Methods of Controlling Arthropod Populations: Population Replacement Using Long Noncoding RNA

[0192] Another method for controlling pest and disease vector populations is a Population Replacement Strategy (PRS) as discussed supra. Population Replacement is dependent on two pieces of technology: 1) A beneficial trait that is desired in the target arthropod; 2) A genetic drive mechanism to spread the desired trait through the arthropod population (Sinkins and Gould, “Gene Drive Systems for Insect Disease Vectors,” Nat. Rev. Genet. 7:427-35 (2006), which is hereby incorporated by reference in its entirety).

[0193] The technology described herein addresses the second problem by utilizing inhibitory polynucleotides that enhance cytoplasmic incompatibility (CI), which is a natural genetic drive mechanism used by various, unrelated bacterial infections (e.g., Wolbachia and Cardinium endosymbionts).

[0194] Disclosed herein are methods for population replacement. Some current uses of a PRS rely on one factor, such as an infection by Wolbachia pipientis. to provide both the beneficial trait and the genetic drive mechanism required for population replacement. The use of inhibitory polynucleotides to enhance CI, however, is an improvement to this approach.

[0195] [1] A genetically modified arthropod, said arthropod comprising:(i) a bacterial endosymbiont encoding a cytoplasmic incompatibility factor; and(ii) a promoter operably linked to an Inhibitory RNA molecule targeting a long noncoding RNA;wherein the expression of the inhibitory' RNA molecule in a male arthropod causes a reduction in fertility and / or the ability of the male arthropod to produce viable offspring in comparison to an endosymbiont-infected male arthropod lacking the Inhibitory RNA molecule.

[0196] [2] The genetically modified arthropod of [1], wherein the long non-codingRNA is reduced by 50% or more compared to a non-genetically modified arthropod.

[0197] [3] The genetically modified arthropod of [1] or [2], wherein the viable offspring are reduced by at least 20%.

[0198] [4] The genetically modified arthropod of any one of [l]-[3], wherein the arthropod is an insect.

[0199] [5] The genetically modified arthropod of [4], wherein the insect is selected from the genera consisting of Aedes, Culex, Drosophila and Anopheles.

[0200] [6] The arthropod of any one of [l]-[5] , wherein the bacterial symbiont isWolbachia.

[0201] [7] A method for controlling a population of target arthropods, comprising:(a) providing a genetically modified arthropod comprising:(i) a bacterial endosymbiont encoding cytoplasmic incompatibility factors; and(ii) a promoter operably linked to an inhibitory RNA molecule targeting a long non-coding RNA; wherein the expression of the inhibitory RNA molecule in a male arthropod causes a reduction in fertility and / or the ability of the male arthropod to produce viable offspring in comparison to a male arthropod lacking the inhibitory RNA molecule; and(b) releasing the genetically modified male arthropods amongst a population of target arthropods under conditions effective to reduce the population of target arthropods.

[0202] [8] A method for replacing a population of target arthropods, comprising:(i) providing a genetically modified replacement arthropod comprising: a promoter operably linked to an inhibitory RNA molecule targeting a long non-coding RNA;(ii) infecting the population of replacement arthropods with a bacterial endosymbiont encoding a cytoplasmic incompatibility factor; and(iii) releasing the replacement arthropods amongst a population of target arthropods, wherein said release of the replacement arthropods reduces the population of target arthropods.

[0203] The following examples are intended to exemplify the practice of embodiments of the disclosure but are by no means intended to limit the scope thereof.EXAMPLESExample 1 - Methods for Examples 2-5

[0204] Fly rearing and strains. D. melanogaster stock yfy* (BDSC 1495) (wMel+) was maintained on a 12-hour light / dark cycle at 25°C and 70% relative humidity on 50 ml of standard fly food media composed of agar, cornmeal, yeast, molasses, ethanol-dissolved tegosept, and propionic acid. IFo / Zw / zza-uninfected (wMel-) fly line was previously generated (LePage et al., “Prophage WO Genes Recapitulate and Enhance Wolbachia-induced Cytoplasmic Incompatibility,” Nature 543:243-247 (2017), which is hereby incorporated by reference in its entirety) by feeding tetracycline-treated food (20 pg / ml in 50 ml of fly media) for three generations followed by two rounds of rearing on standard food media. Before using the lines for experimental purposes, their infection status was confirmed by PCR using Wolb_F and Wolb_R3 primers (Casiraghi et al., “Phylogeny of Wolbachia pipientis Based on gltA, groEL and ftsZ Gene Sequences : Clustering of Arthropod and Nematode Symbionts in the F Supergroup, and Evidence for Further Diversity in the Wolbachia Tree,” Microbiology (Reading) 15 l(Pt 12):4015-4022 (2005), which is hereby incorporated by reference in its entirety).

[0205] Chemical preparation and feeding. For chemical feeding assays, a stock solution of 28.69 mM anacardic acid (AA) (Merck Millipore, #172050) in dimethyl sulfoxide (DMSO) (Sigma, #D8418) solvent was prepared as previously described (Gartner et al., “Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment,” Journal of Visualized Experiments 11 :(91):51868 (2014), which is hereby incorporated by reference in its entirety). In 10 ml standard fly food vials, 52.56 pl stock solution (AA+DMSO) was added to reach the desired final concentration (150 pM) per vial. The same amount of DMSO solvent alone was mixed with standard food to create a ‘control’ chemical treatment condition in each experiment. To setup chemical -treated flies for conducting CI assays, wMel- paternal grandmothers were first mass-reared in standard 50 ml food bottles. Virgin females were collected and aged to 11 days before mating with wMel- males. Males and females in a 4:4 ratio were transferred into 8 oz bottles each affixed with a grape-juice agar plate smeared with yeast. Following incubation at 25°C for 24h, grape plates were discarded and replaced with freshly smeared ones and bottles wereincubated again for 24h. The grape plates with eggs on them were collected and incubated for additional 24h to allow egg to larvae development. A total of 50 larvae at LI - L2 stage were manually picked and gently placed into AA-treated and control food vials each. Since AA is reportedly light-sensitive, the chemicals were mixed in the food under minimum light exposure the same morning of larvae transfer. After mixing, the vial trays were covered with aluminum foil and stored in the dark until the larvae transfer process started. In parallel, wMel- and wMel- flies were setup to use as positive and negative CI controls, respectively, by rearing them on the same standard fly food media lacking the chemicals. Food was manually churned using a spatula and larvae were manually picked and transferred to the churned food vials similar to chemical-mixed food to avoid any confounding effect.

[0206] Hatch rates. Male flies hatching between 0-4 hours from AA- and DMSO- treated food vials were collected and used in hatch rate (HR) assay to ensure higher CI penetrance. HR set up was used as described previously (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-induced Cytoplasmic Incompatibility in Drosophila melanogaster,” mBio 10(6):e01879- 1 (2019), which is hereby incorporated by reference in its entirety). Briefly, a male and female pair was placed in an 8 oz, round bottom, polypropylene Drosophila stock bottle with a yeast-smeared grape juice agar plate affixed. The bottles were placed in a 25°C incubator overnight to allow mating. The following day, these plates were discarded and replaced with new grape juice agar plates with fresh yeast smear. After an additional 24 hours, the plates were removed, and the total number of embryos were counted. The embryo plates were then incubated for 36 hours at 25°C before the number of unhatched embryos were counted. Any plates with less than 20 embryos laid were discarded from the analyses. Significant differences (p < 0.05) were determined by pairwise Mann Whitney U tests or by a Kruskal-Wallis test and Dunn multiple test correction in GraphPad Prism 10.

[0207] Immunofluorescence. Sibling males from the above HR assay (0-4 hours old) were collected for testes dissection in ice-cold 1 x PBS solution. Tissues were fixed in 4% formaldehyde and processed for immunostaining as per our previous study (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). Briefly, samples were washed, blocked in BSA, and incubated with primary antibodies overnight at 4°C. Tissues were then incubated with secondary antibodies for 4 hours at room temperature in the dark. Tissues were then washed and mounted on slides using a DAPLcontaining Vectashield medium. Imaging was performedusing Zeiss LSM880 confocal microscope under constant exposure settings across treatment groups. Images processing and quantification was performed using ImageJ software.

[0208] Histone-specific antibodies used were rabbit polyclonal a) anti-Acetyl-Histone H3 (EMD Millipore, Cat. No. 06-599, 1:500 dilution) recognizing acetylation on K9 and K14 lysine residues, b) anti-acetyl-Histone H4 (EMD Millipore, Cat. No. 06-598, 1 :5000 dilution) recognizing acetylation of K5, K8, KI 2, and KI 6 residues, and c) anti -Histone H3 (EMD Millipore, Cat. No. 06-755, 1: 100 dilution). Core histones were detected with mouse monoclonal anti-Histone antibody (EMD Millipore, Cat. No. F152.C25.WJJ, 1: 1200 dilution). Anti-mouse alexa488 and anti-rabbit alexa594 conjugated secondary' antibodies (Vector Laboratories) were used at 1 :500 dilutions. Rabbit polyclonal anti-Histone H4 antibody (EMD Millipore, Cat. No. 07-108) were also tested to detect H4 histone retention in multiple replicates, however, the antibody did not work.

[0209] Mature sperm isolation, CMA3 staining, and quantification. Sperm from seminal vesicles of 0-4 hours old sibling HR males were extracted and processed for CMA3- based staining assay as described previously (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety ). Fluorescence quantification was performed by scoring fluorescent pixels in arbitrary units (A.U.) within individual sperm head using ImageJ software. The calculated fluorescence intensity per sperm head was graphed. Statistical significance (p < 0.05) was determined by a pairwise comparison using Mann-Whitney test in GraphPad Prism 10.Example 2 - Wolbachia Reduce Histone Acetylation During Sperm Development

[0210] Symbiotic relationships between arthropod hosts and bacteria have sparked interest and success in leveraging microorganisms to control pests and disease vectors. As arthropod-symbiont relationships are molecularly disentangled, approaches that bypass the symbionts to replicate traits directly in a symbiont-free arthropod will serve as adjuncts or new alternatives to current vector control measures.

[0211] During I). melanogaster spermiogenesis, DNA-bound histones undergo various posttranslational modifications (PTMs) for removal and replacement by protamines for a tight chromatin organization (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol.20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). Histone acetylation is a prerequisite PTM for histone detachment (Grunstein, “Histone Acetylation inChromatin Structure and Transcription,” Nature 389:349-352 (1997), which is hereby incorporated by reference in its entirety) as inhibition of H3 and H4 acetylation blocks the histone-to-protamine switch and consequently, the chromatin remains histone-bound (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster ” Syst. Biol. Reprod. Med. 56:44-61 (2010) and Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster,” PLoS One 13(9):e0203622 (2018), each of which is hereby incorporated by reference in its entirety). Because D. melanogaster Wolbachia (wMel) and transgenic- expression of the CI proteins cause abnormal core histone retention in CI males (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety), it was first tested if retention is linked to reduced histone acetylation (Bannister & Kouzarides, “Regulation of Chromatin by Histone Modifications,” Cell Res. 21 :381-395 (2011) and Bohm et al., “Nucleosome Accessibility Governed by the Dimer / Tetramer Interface,” Nucleic Acids Res. 39:3093-3102 (2011), each of which is hereby incorporated by reference in its entirety). In wMel+ CI males, there was a significant reduction in H3 acetylation (H3ac) in all stages of sperm development (fold change reduction in spermatocytes: 1.8; young elongating: 9.8; early canoe: 5.9; late canoe: 14.1) relative to uninfected, wMel- negative control males that do not cause CI (FIGs. 1B-C). Control males, as expected, showed H3ac in primary spermatocytes followed by signal waning in round onion spermatids and then reappearing in young elongating and early canoe spermatids (Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster ” PLoS One 13(9):e0203622 (2018), which is hereby incorporated by reference in its entirety)- Faint disappearing H3ac signals in control late canoe spermatids correlated with the typical histone removal during this stage (FIG. IB). While H4ac signal intensity levels did not differ among CI and non-CI males across all stages (FIG. ID, FIG. 6), there was a significant reduction in the number of early canoe spermatid bundles with H4ac signals in wMel+ males compared to the negative control (FIG. IE, FIG. 7). Altogether, the data suggest that both H3 and H4 acetylation are reduced / inhibited in wild type CI males.

[0212] During sperm development in Drosophila melanogaster, DNA-bound histones undergo various posttranslational modifications (PTMs) for removal and replacement byprotamines to tightly package the chromatin into a classic, needle-shaped sperm. Histone acety lation is considered a prerequisite PTM for histone detachment as inhibition of H3 and H4 histone acetylation blocks the histone-to-protamine switch and consequently, the chromatin DNA remains histone-bound. In / ), melanogaster, wild type CI levels of embryonic mortality induced by ‘wMel’ strain of Wolbachia are -70%. We previously showed that wMel Wolbachia cause abnormal retention of histones in CI males Here, it was found that the abnormal histone retention in wMel+ males was due to significantly reduced H3 acety lation (H3ac) in all stages of sperm development relative to non-CI causing wMel- males. Moreover, a reduced number of early developing spermatid bundles with H4ac signals in wMel+ males compared to the negative control were identified. Altogether, these results indicated that both H3 and H4 acetylation are reduced / inhibited in CI males.Example 3 - Small Molecule Inhibitor Reduces Histone Acetylation in Aposymbiotic Males to Recapitulate CI

[0213] Next, it was tested whether reduced H3 and H4 acetylation is causal to CI as measured by a decrease in embryos hatching into larvae. Anacardic acid (AA) is a small molecule compound that inhibits histone acetyltransferase (HAT) enzyme activity from invertebrates to vertebrates (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster ” Syst. Biol. Reprod. Med. 56:44-61 (2010), Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster ” PLoS One 13(9):e0203622 (2018), Sun et al., “Inhibition of Histone Acetyltransferase Activity by Anacardic Acid Sensitizes Tumor Cells to Ionizing Radiation,” FEBSLett. 580:4353-4356 (2006), and Balasubramanyam et al., “Small Molecule Modulators of Histone Acetyltransferase p300,” J. Biol. Chem. 278:19134- 19140 (2003), each of which is hereby incorporated by reference in its entirety). Crucially in D. melanogaster, AA-exposed in vitro cultures of intact pupal testes and sperm cysts deplete histone H3 and H4 acetylation that in turn inhibits the histone-to-protamine (H-P) exchange during spermiogenesis (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster,” Syst. Biol. Reprod. Med. 56:44-61 (2010), which is hereby incorporated by reference in its entirety). Using males that lack Wolbachia or CI proteins expression, it was tested if AA treatment of wMel-uninfected (wMel-) D. melanogasterrecapitulates CI upon inhibiting HAT activity, reducing histone acetylation, and altering H-P transition.

[0214] To do this, wMel- 1stand 2ndinstar larvae were fed 150 pM AA in 10 ml standard food vials to assimilate the chemical impact from early stages of sperm development, since meiotic divisions commence in the testes of 3rdinstar larvae (Gartner et al., “Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment,” Journal of Visualized Experiments 11 :(91):51868 (2014), which is hereby incorporated by reference in its entirety). DMSO alone (the solvent in which AA was dissolved) mixed in standard fly food vials was used as a negative control group. Remarkably, AA-treated wMel- males recapitulated reduced embryonic hatching (34.3%), similar to wMel+ wild type CI (27.6%), which were rescued to high levels of embryonic hatching upon crossing to wMel+ females (78.3%) (FIG. 2B). Notably, rescue indicates that reduced embryonic hatching is bona fide CI rather than a chemically-conferred sterility artifact. DMSO-control treated males showed high embryonic hatching when crossed with both w Mel- and wMel+ females (median = 81% and 81.95%, respectively). Some variation in embryonic hatch rates occurred in both AA and DMSO crosses, presumably due to off-target impacts of DMSO.

[0215] Similar to wild type wMel+ males, AA-treated CI males showed significantly reduced H3ac testes signal in all post-meiotic sperm development stages (fold change reduction in young elongating: 1.45; early canoe: 3.27; late canoe: 5.0) compared to DMSO testes (FIG. 3 A). Similarly, the characteristic pattern of H4ac was detected in all post-meiotic stages of DMSO-testes, whereas in sharp contrast, no to low H4ac signals were detected in AA-testes from round spermatids till the elongating, early canoe spermatid nuclei stages (fold change in round onion: 18.1; young elongating: 12.3; early canoe: 16.4) (FIG. 3C-D). Both H3ac and H4ac levels did not differ in spermatocytes of AA- and DMSO-treated testes during the early stages of spermatogenesis (FIGs. 3A, 3C). This could be due to the larvae having already developed spermatocytes before undergoing AA feeding, thereby mitigating the chemical impact on HAT activity at this stage. By chemically-depleting histone acetylation levels, it was demonstrated that the AA treatment can engineer wMel- males to recapitulate CI at levels comparable to wild type wMel+ induced CI.Example 4 - CI Sperm from Treated Aposymbiotic Males Develop with Abnormal H3 Histone Retention and Protamine Deficiency

[0216] HATs can acetylate all four core histones H2A, H2B, H3, and H4 with stronger preferences for H3 and H4 (Dancy & Cole, “Protein Lysine Acetylation by p300 / CBP,” Chem. Rev. 115:2419-2452 (2015), which is hereby incorporated by reference in its entirety). They do so by transferring an acetyl group from acetyl-Coenzyme A (acetyl-CoA) to the s- amino group of lysine side chains. As the lysine-rich histone tails are positively charged and interact with the negatively charged nucleosomal DNA, HAT-mediated acetylation weakens histone tail-DNA interactions (Bannister & Kouzarides, “Regulation of Chromatin by Histone Modifications,” Cell Res. 21 : 381 —395 (2011), which is hereby incorporated by reference in its entirety), causing histones to be removed. Since reduced / inhibited H3ac and H4ac levels in AA-treated aposymbiotic, Ci-inducing males was observed, it was hypothesized that reduced acetylation results in a lack of acetylated H3 and H4 for their timely removal from the chromatin.

[0217] Immunostaining of dissected testes with H2B and H3 antibodies demonstrated that H3 remains attached to the maturing sperm chromatin in CI males. H3 signals were consistently detected in Ci-inducing, AA-treated wMel- and untreated wild type wMel+ males until late elongating and needle-stages of spermiogenesis, whereas signals disappeared in negative control groups, as expected (FIGs. 4A-C). The H2B signals, however, did not vary between CI and non-CI groups (FIGs. 8A-B). The signals vanished at the late canoe stage, suggesting that nucleosomal removal of H2A-H2B dimer is unaffected. Since histone elimination occurs in a sequential manner with H2A-H2B dimer removal followed by H3 and H4 tetramer removal (Bohm et al., “Nucleosome Accessibility Governed by the Dimer / Tetramer Interface,” Nucleic Acids Res. 39:3093-3102 (2011), which is hereby incorporated by reference in its entirety), it was concluded that H2B (and possibly H2A) histones dimers are removed normally from both CI and non-CI chromatin independently of the H3 (and possibly H4) tetramers consistent with reduced H3 and H4 acetylation, respectively. The data also specifies H3 as the specific histone type retained in chromatin of developing CI sperm.

[0218] Moreover, the mature sperm isolated from seminal vesicles of AA-treated CI males exhibited a 1.26-fold significant increase in CMA3 intensity, consistent with protamine deficiency relative to DMSO-treated sperm (FIG. 4C). The data align with our previous studies in wMel+ CI males in which abnormal histone retention leads to protamine deficiency Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrityto Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5):e3001584 (2022) and Kaur et al., “The Mechanism of Cytoplasmic Incompatibility is Conserved in Wolbachia-beahng Aedes aegypti Mosquitoes Deployed for Arbovirus Control,” PLoSBiol 22:e3002573 (2024), each of which is hereby incorporated by reference in its entirety). Altogether, these results specify that anacardic acid inhibits the enzymatic activity of histone acetyltransferase to disrupt the histone-to-protamine transition underpinning abnormal chromatin integrity of developing sperm.Example 5 - Discussion of Examples 1-4

[0219] The symbiosis between Wolbachia and arthropods is established as the most common in the animal world with central roles in vector-home disease control (Walker et al., “The wMel Wolbachia Strain Blocks Dengue and Invades Caged Aedes aegypti Populations,” Nature 476:450-455 (2011) and Ross & Hoffmann, “Continued Susceptibility of the wMEL Wolbachia Infection in Aedes aegypti to Heat Stress Following Field Deployment and Selection,” Insects 9(3): 78 (2018), each of which is hereby incorporated by reference in its entirety) and insect reproductive isolation (Bordenstein et al., “JTb / iacA / a-induced Incompatibility Precedes Other Hybrid Incompatibilities in Nasonia,” Nature 409:707-710 (2001, Breeuwer & Werren, “Microorganisms Associated with Chromosome Destruction and Reproductive Isolation Between Two Insect Species,” Nature 346(6284):558-60 (1990), Jaenike et al., “Asymmetrical Reinforcement and Wolbachia Infection in Drosophila f PLoS Biol. 4: 1852-1862 (2006) and Shoemaker et al., "'Wolbachia and the Evolution of Reproductive Isolation Between Drosophila recens and Drosophila subquinariaf Evolution (N Y) 53: 1157-1164 (1999), each of which is hereby incorporated by reference in its entirety).

[0220] Furthermore, the symbiosis is an archetype of how Wolbachia' s prophage proteins hijack animal reproductive biology to shape reproductive phenotypes such as male killing and cytoplasmic incompatibility (Shropshire et al., “Symbiont-Mediated Cytoplasmic Incompatibility : What Have We Learned in 50 Years?,” Elife 9:e61989 (2020), which is hereby incorporated by reference in its entirety). Deciphering the molecular mechanism of these adaptations has been a long-sought goal not only to understand the nature of a widespread symbiosis, but to directly engineer functional traits for positive human health outcomes. Here, key molecular switch that recapitulates CI and its defining cellular mechanism in aposymbiotic males via inhibition of histone acetylation - a post-translational epigenetic modification was unlocked. These data reveal that aposymbiotic males treated with a small molecule inhibitor, anacardic acid (AA), recapitulate rescuable CI upon developing CIhallmarks of reduced histone acetylation, histone retention, and protamine deficiency during spermiogenesis.

[0221] Histone acetylation is linked to transcription and chromatin reorganization as it causes chromatin to transition from tightly packed heterochromatin to loosely packed euchromatin states (Grunstein, “Histone Acetylation in Chromatin Structure and Transcription,” Nature 389:349-352 (1997) and Richards & Elgin, “Epigenetic Codes for Heterochromatin Formation and Silencing: Rounding Up the Usual Suspects,” Cell 108:489- 500 (2002) each of which is hereby incorporated by reference in its entirety), rendering DNA more accessible to transcription factors, enzymes, and other proteins involved in transcription. Long non-coding RNA (IncRNAs) often collaborate with enhancer and promoter regions to mediate histone modifications, including acetylation, and activate target genes for successful chromatin organization (Hah et al., “Transcripts Mark Active Estrogen Receptor Binding Sites, “ Genome Res 23: 1210-1223 (2013), Zhu et al., “Predicting Enhancer Transcription and Activity from Chromatin Modifications,” Nucleic Acids Res. 41 : 10032-10043 (2013), Lai et al., “Activating RNAs Associate with Mediator to Enhance Chromatin Architecture and Transcription,” Nature 494:497-501 (2013), each of which is hereby incorporated by reference in its entirety). It was recently showed in Drosophila spermatocytes that wMel Wolbachia and CifA mediate depletion of a IncRNA that is essential for a normal H-P exchange (Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety). Specifically, IncRNA depletion leads to retained histones in developing spermatids, increased protamine deficiency in mature sperm, and enhancement of wild type CI (Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety). Here, in the absence of Wolbachia and IncRNA depletion, a chemical disruption of the downstream process of histone acetylation and H-P exchange recapitulated the CI trait.

[0222] There is also increasing evidence that histone acetylation is involved in DNA damage repair (Dhar et al., “The Tale of a Tail: Histone H4 Acetylation and the Repair of DNA Breaks,” Philosophical Transactions of the Royal Society B: Biological Sciences 372(1731):20160284 (2017), Hunt et al., “Histone Modifications and DNA Double-strand Break Repair After Exposure to Ionizing Radiations,” Radiat. Res. 179:383-392 (2013), each of which is hereby incorporated by reference in its entirety). As chromatin structure regulatesthe accessibility of DNA to a variety of different factors, H3 and H4 acetylation can modulate double-stranded DNA break repair by recruiting DNA repair enzymes or by facilitating homologous recombination, respectively (Horikoshi et al., “Pre-existing H4K16ac Levels in Euchromatin Drive DNA Repair by Homologous Recombination in S-phase,” Commun. Biol. 2:253 (2019), which is hereby incorporated by reference in its entirety). In addition, HAT- mediated H3 and H4 acetylation can facilitate the recruitment of non-homologous end-joining proteins to DNA damage sites (Ogiwara et al., “Histone Acetylation by CBP and p300 at Double-strand Break Sites Facilitates SWI / SNF Chromatin Remodeling and the Recruitment of Non-homologous End Joining Factors,” Oncogene 30:2135-2146 (2011), which is hereby incorporated by reference in its entirety). Consequently, the reduced acetylation is consistent with enhanced DNA damage in developing CI sperm during spermiogenesis (Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383:1111-1117 (2024), which is hereby incorporated by reference in its entirety).

[0223] Once fertilization occurs, paternal chromatin undergoes chromatin remodeling such as protamines from the paternal chromatin are removed and replaced by maternal histones to decondense and activate the chromatin of the developing embryo (McLay & Clarke, “Remodelling the Paternal Chromatin at Fertilization in Mammals,” Reproduction 125:625-633 (2003), which is hereby incorporated by reference in its entirety). Interestingly, postfertilization delays in maternal H3.3 histone deposition occur in CI embryos (Landmann et al., “JFb / tocto-mediated Cytoplasmic Incompatibility is Associated with Impaired Histone Deposition in the Male Pronucleus,” PLoS Pathog. 5(3) : e 1000343 (2009), which is hereby incorporated by reference in its entirety). It is proposed here that the delay can be due to preloaded paternal H3 (and H4) histones causing altered paternal epigenome information that results in chromatin remodeling errors and mistiming of maternal H3.3 deposition. The symbiotic embryos may possibly rescue Ci-derived paternal chromatin errors by establishing similar maternal chromatin modifications during oogenesis due to Wolbachia or transgenic expression of the cifA rescue gene, as previously proposed (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5) :e3001584 (2022), Kaur et al., “The Mechanism of Cytoplasmic Incompatibility is Conserved in Wolbachia-bearmg Aedes aegypti Mosquitoes Deployed for Arbovirus Control,” PLoS Biol. 22:e3002573 (2024), and Dhar et al., “The Tale of a Tail: Histone H4 Acetylation and the Repair of DNA Breaks,” PhilosophicalTransactions of the Royal Society B: Biological Sciences 372(1731 ):20160284 (2017), each of which is hereby incorporated by reference in its entirety).

[0224] Overall, these data strongly support the Host Modification (HM) model of CI in which CI can be achieved by modifying a pre-fertilization, host factor without Wolbachia or its effector proteins CifA and CifB. Finally, it is noteworthy that some arthropods are refractory to Wolbachia, natural CI strength varies in Wolbachia-canying arthropods, and some Wolbachia strains do not cause CI in vectors (Mercot & Charlat, "Wolbachia Infections in Drosophila melanogaster and D. simulans: Polymorphism and Levels of Cytoplasmic Incompatibility,” Genetica 120:51-59 (2004), which is hereby incorporated by reference in its entirety). Therefore, chemically reprogramming reproductive epigenetics in the absence of Wolbachia or Cif expression provides a new gateway to engineer this symbiotic trait crucial to arthropod biology and vector control programs.Example 6 - Wolbachia Deplete Histone Acetylation During Sperm Development to Cause CI

[0225] During Drosophila melanogaster spermiogenesis, DNA-bound histones on immature spermatids undergo various posttranslational modifications (PTMs) for removal and replacement by protamines to form a tightly condensed mature sperm chromatin (Rathke et al., “Chromatin Dynamics During Spermiogenesis,” Biochim. Biophys. Acta Gene Regal. Meeh. 1839: 155-168 (2014), which is hereby incorporated by reference in its entirety) (FIGs. 1 A-E). Histone acetylation is a prerequisite PTM for histone removal (Grunstein, M., “Histone Acetylation in Chromatin Structure and Transcription,” Nature 389:349-352 (1997), which is hereby incorporated by reference in its entirety) as inhibition of histone H3 and H4 acetylation blocks the histone-to-protamine switch and consequently, the chromatin remains histone-bound (Awe & Renkawitz-Pohl. “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster f Syst. Biol. Reprod. Med. 56:44-61 (2010) and Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster,” BLoS One 13(9):e0203622 (2018), each of which is hereby incorporated by reference in its entirety). Because D. melanogaster Wolbachia (wMel) and transgenic expression of the CI proteins, CifA and CifB, cause abnormal core histone retention in CI spermatids (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrify to Establish Cytoplasmic Incompatibility,” PLoS Biol. 220(5): e3001584 (2022), which is hereby incorporated by reference in its entirety), we hypothesized that the histone retention is linked to reduced histone acety lation.

[0226] In symbiotic CI males, a significant reduction in H3 acetylation (H3ac) in all stages of sperm development (fold change reduction in spermatocytes: 1.8; young elongating: 9.8; early canoe: 5.9; late canoe: 14.1) relative to uninfected, wMel- negative control males that do not cause CI (FIG. 1A-C) was identified. Control males, as expected, showed H3ac in primary spermatocytes followed by signal waning in round onion spermatids and then reappearance in young elongating and early canoe spermatids (Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility in Drosophila melanogaster ” PLoS One 13(9):e0203622 (2018), which is hereby incorporated by reference in its entirety). Faint disappearing H3ac signals in control late canoe spermatids correlated with the typical histone removal during this stage (FIGs. 1B- C). H4ac signal intensity levels did not differ among CI and non-CI males (FIG. ID, FIG. 6), but the number of early canoe spermatid bundles with H4ac signals in wMel+ CI males compared to the negative control was 1.75-fold, significantly reduced (FIG. IE, FIG. 7). Altogether, the data specify that Wolbachia deplete either the level and / or prevalence of H3 and H4 acetylation in wild type CI males.Example 7 - Transcriptional Silencing of Histone Acetyltransferase by RNAi Enhances Wild Type CI

[0227] To test if reduced histone acetylation was causal to CI, RNAi-mediated knockdown of the histone acetyltransferase (Hail) that acetylates all four core histones H2A, H2B, H3, and H4 with stronger preferences for H3 and H4 was carried out. HAT1 is described in Parthun, M.R., “Histone Acetyltransferase 1: More Than Just an Enzyme ." Biochimica et Biophysica Acta (BBA) - Gene Regulatory Mechanisms 1819:256-263 (2012) and Varga et al., “Hatl Acetylates Histone H4 and Modulates the Transcriptional Program in Drosophila embryogenesis,” Sci. Rep. 9(1): 17973 (2019), each of which is hereby incorporated by reference in its entirety. HAT1 RNAi line BDSC#42488 and HDAC1 RNAi line VDRC#30600 from the Bloomington Drosophila Stock Center and the Vienna Drosophila Resource Center, respectively, were used.

[0228] Hatl knockdown males (Hat IRNAi ) were generated using paternal grandmothers aged 5 days old that normally induce incomplete / intermediate levels of wild type CI (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-induced Cytoplasmic Incompatibility in Drosophila melanogaster ” mBio 10(6):e01879-19 (2019),which is hereby incorporated by reference in its entirety) to assess if CI strength increases upon knockdown otHatl. CI levels were measured by standard assays that calculate the percentage of embryos hatching into larvae CI (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-induced Cytoplasmic Incompatibility in Drosophila melanogaster ” mBio 10(6): e01879- 19 (2019), which is hereby incorporated by reference in its entirety).

[0229] The sense and antisense nucleotide sequences used in the RNAi construct targeting Drosophila melanogaster HAT1 are: 21bp_seq_sense nucleotide sequence CAGAAGAATGTTGTGGACATA (SEQ ID NO: 1), and 21bp_seq_antisense nucleotide sequence TATGTCCACAACATTCTTCTG (SEQ ID NO:2). The sense and antisense nucleotide sequences used in the RNAi construct targeting Drosophila melanogaster HDAC1 are: 21bp_seq_sense nucleotide sequence GTCTAGAGAGGCGTTCTATACCACCGA (SEQ ID NO:30), and 21bp_seq_antisense nucleotide sequence AGAATTCATGTGCAGCTTAAAATCGGG (SEQ ID N0:31).

[0230] As shown in FIG. 9A, HatlRNAi+ males induced approximately 4.5-fold higher CI (median embryonic hatching = 5.08%) relative to control wild type wMel+ males (median embryonic hatching = 22.5%).

[0231] HAT activity is reversable by histone deacetylases (HD AC) that catalyze removal of acetyl groups from histones (Thiagalingam et al., “Histone Deacetylases: Unique Players in Shaping the Epigenetic Histone Code ” Ann N Y Acad Sci. 983:84-100 (2003), which is hereby incorporated by reference in its entirety). HDAC1 knockdown males (HDAClRNAi+) were generated using paternal grandmothers aged 11-13 days old that typically induce very strong levels of wild type CI (Layton et al., “Paternal Grandmother Age Affects the Strength of Wolbachia-induced Cytoplasmic Incompatibility in Drosophila melanogaster ” mBio 10(6): e01879- 19 (2019), which is hereby incorporated by reference in its entirety ) to assess if CI strength decreases upon knockdown of HDAC1.

[0232] Notably, CI levels were suppressed in HDAC1RNAI+ males (median embryonic hatching = 20.0%) compared to wild type wMel+ males (median embryonic hatching = 3.25%, FIG. 9B). These findings specify a causal link between Wolbachia- mediated reduced histone acetylation and strong CI, a phenomenon that can be reversed by downregulating histone deacetylation.Example s - Small Molecule Inhibitor of Histone Acetylation Recapitulates CI and Cell Biological Marks in Aposymbiotic Males

[0233] Next, it was tested whether engineering the histone acetylation pathway in the absence of Wolbachia recapitulates CI. Anacardic acid (AA) is a small molecule compound that inhibits HAT enzyme activity to acetylate H3 and H4 across the animal kingdom (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogasterf Syst. Biol. Reprod. Med. 56:44-61 (2010), Hundertmark et al., “Nejire / dCBP-mediated Histone H3 Acetylation During Spermatogenesis is Essential for Male Fertility7in Drosophila melanogaster f PLoS One 13(9):e0203622 (2018), Sun et al., “Inhibition ofHistone Acetyltransferase Activity by Anacardic Acid Sensitizes Tumor Cells to Ionizing Radiation,” FEBSLett. 580:4353-4356 (2006) and Balasubramanyam et al., “Small Molecule Modulators ofHistone Acetyltransferase p300,” J. Biol. Chem. 278: 19134-19140 (2003), each of which is hereby incorporated by reference in its entirety).

[0234] Aposymbiotic (vt Mel -) 1st and 2nd instar larvae were fed 150 pM AA in 10 ml standard food vials to assimilate the chemical impact during the early stages of sperm development, since meiotic divisions commence in the testes of 3rd instar larvae (Gartner et al., “Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment,” Journal of Visualized Experiments (91):51868 (2014), which is hereby incorporated by reference in its entirety). DMSO alone (the solvent in which AA was dissolved) mixed in standard fly food vials was used as a negative control (FIG. 2A). Notably, AA-treated, wMel- males recapitulated reduced embryonic hatching (34.3%), similar to wMel+ wild type CI (27.6%). The embryonic inviability was in turn rescued to high levels of embryonic hatching upon crossing both male types to wMel+ females (FIG. 2B). This rescue indicated that the AA-mediated reduced embryonic hatching is a recapitulation of CI rather than a chemically-conferred sterility artifact. Negative control DMSO-treated, wMel- males showed high embryonic hatching when crossed with both vt’Mel- and w’Mel+ females (median = 81% and 81.95%, respectively). Some variation in embryonic hatch rates occurred in both AA and DMSO crosses, presumably due to off-target impacts of DMSO.

[0235] Histone acetylation depletion in wild type wMel+ males and AA-treated wMel- males was further verified using the aforementioned antibodies and quantification metrics. Both groups showed significantly reduced H3ac signal in all post-meiotic sperm stages (fold change reduction in young elongating: 1.45; early canoe: 3.27; late canoe: 5.0) relative toDMSO (FIGs. 3A-B). Similarly, the characteristic pattern of H4ac was detected in all post- meiotic stages of DMSO-testes, whereas in sharp contrast, no to low H4ac signals were detected in AA-testes from round spermatids till the elongating, early canoe spermatid nuclei stages (fold change in round onion: 18.1; young elongating: 12.3; early canoe: 16.4) (FIGs. 3C-D). Both H3ac and H4ac levels did not differ in spermatocytes of AA- and DMSO-treated testes during the early stages of spermatogenesis (FIGs. 4A-D). This could be due to the larvae having already developed spermatocytes before undergoing AA feeding (Gartner et al., "Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection, Imaging, and Pharmacological Treatment,” Journal of Visualized Experiments (91):51868 (2014), which is hereby incorporated by reference in its entirety), thereby mitigating the chemical impact on HAT activity at this stage. By chemically-depleting histone acetylation levels, it was demonstrated that the AA treatment can engineer wMel- males to recapitulate CI at levels comparable to wild type wMel+ induced CI.Example 9 - CI Sperm from Treated Aposymbiotic Males Exhibit H3 Histone Retention and Protamine Deficiency

[0236] HAT-mediated acetylation weakens the interaction between histone tails and negatively charged DNA, causing nucleosomal histones to be removed (Bannister & Kouzarides, “Regulation of Chromatin by Histone Modifications,” Cell Res. 21 :381-395 (2011), which is hereby incorporated by reference in its entirety). Since reduced / inhibited H3ac and H4ac levels in AA-treated, Ci-inducing aposymbiotic males was observed, it was next hypothesized that the treatment causes H3 and H4 histone retention in both CI and AA- treated developing sperm. Moreover, since histone elimination occurs in a sequential manner with H2A-H2B dimer removal followed by H3 and H4 tetramer removal (Bohm et al., “Nucleosome Accessibility Governed by the Dimer / Tetramer Interface,” Nucleic Acids Res. 39:3093-3102 (2011), which is hereby incorporated by reference in its entirety), H2B retention was also monitored.

[0237] Immunostaining of dissected testes with H2B and H3 antibodies demonstrated that H3 remains attached to the maturing sperm chromatin in CI males (FIGs. 4A-C), whereas H2B is properly removed (FIGs. 8A-B). H3 signals were consistently detected in AA-treated wMel- and untreated wild type wMel+ testes until late elongating and needle-stages of spermiogenesis, whereas signals disappeared in negative control groups, as expected (FIGs. 4A-B, FIGs. 5A-B). The H2B signals, however, did not vary between CI and non-CI groups (FIGs. 8A-B), and they vanished at the late canoe stage, suggesting that nucleosomal removalof H2A-H2B dimer is unaffected. It was concluded that H2B (and possibly H2A) histones are removed normally from both CI and non-CI chromatin, whereas H3 (and possibly H4) histones are retained, which is consistent with the observed, reduced H3 and H4 acetylation.

[0238] Finally, the mature sperm isolated from seminal vesicles of AA-treated CI males exhibited a 1.26-fold significant increase in CMA3 intensity, consistent with protamine deficiency relative to DMSO-treated sperm (FIG. 4C). The data align with our previous studies in wMel+ CI males in which CI results in both histone retention and protamine deficiency (Kaur et al., “The Cif Proteins From Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 2 20(5):e3001584 (2022) and Kaur et al., “The Mechanism of Cytoplasmic Incompatibility is Conserved in Wolbachia-^Qa.m^ Ae des aegypti Mosquitoes Deployed for Arbovirus Control,” PLoS Biol. 22:e3002573 2024, each of which is hereby incorporated by reference in its entirety). Altogether, these results specify that anacardic acid inhibits the enzy matic activity of histone acetyltransferase to disrupt the histone-to-protamine transition underpinning abnormal chromatin integrity of developing CI sperm.Example 10 - Discussion of Examples 6-9

[0239] The symbiosis between Wolbachia and arthropods is common and widespread, with central roles in vector-borne disease control (Walker et al., “The wMel Wolbachia Strain Blocks Dengue and Invades Caged Aedes aegypti Populations,” Nature 476:450-455 (2011) and Ross & Hoffmann, “Continued Susceptibility of the wMEL Wolbachia Infection in Aedes aegypti to Heat Stress Following Field Deployment and Selection,” Insects 9(3): 78 (2018), each of which is hereby incorporated by reference in its entirety) and arthropod reproductive isolation (Bordenstein et al., “IFo / bac / i / a-induced Incompatibility Precedes Other Hybrid Incompatibilities in Nasonia,” Nature 409:707-710 (2001), Breeuwer & Werren, “Microorganisms Associated with Chromosome Destruction and Reproductive Isolation Between Two Insect Species,” Nature 346(6284):558-60 (1990), and Shoemaker et al., “Wolbachia and the Evolution of Reproductive Isolation Between Drosophila recens and Drosophila subquinariaf Evolution 53: 1157-1164 (1999), each of which is hereby incorporated by reference in its entirety). Furthermore, it is an exemplar of how prophage- associated enzymes in an endosymbiont hijack animal reproductive biology to cause major impacts on sex ratios and sexual reproduction (Shropshire et al., “Symbiont-mediated Cytoplasmic Incompatibility: What Have We Learned in 50 Years?,” Elife 9:e61989 (2020), which is hereby incorporated by reference in its entirety). Deciphering the key molecularswitch that turns on symbiotic adaptations has been a long-sought goal in the field, not only to understand the nature of a widespread symbiosis, but to directly engineer functional traits for positive human health outcomes. As shown herein, inhibition of histone acetylation- a crucial post-translational epigenetic modification - is unlocked as a master regulator to engineer CI independent of the bacteria or its genes. The data in the present disclosure most notably reveal that aposymbiotic males treated with a small molecule inhibitor, anacardic acid (AA), recapitulate rescuable CI upon developing cellular hallmark signatures of CI, namely reduced histone acetylation, abnormal histone retention, and protamine deficiency during spermiogenesis.

[0240] Histone acetylation is linked to transcription and chromatin reorganization as it causes chromatin to transition from tightly packed heterochromatin to loosely packed euchromatin states (Grunstein, “Histone Acetylation in Chromatin Structure and Transcription,” Nature 389:349-352 (1997) and Shoemaker et al., “Wolbachia and the Evolution of Reproductive Isolation Between Drosophila recens and Drosophila subquinaria ” Evolution 53: 1157-1164 (1999), which is hereby incorporated by reference in its entirety), rendering DNA more accessible to transcription factors, enzymes, and other proteins involved in transcription. Specifically, long non-coding RNA (IncRNAs) often collaborate with enhancer and promoter regions to mediate histone modifications, including acetylation, and activate target genes for successful chromatin organization (Hah et al., “Enhancer Transcripts Mark Active Estrogen Receptor Binding Sites,” Genome Res. 23: 1210— 1223 (2013), Zhu et al., “Predicting Enhancer Transcription and Activity From Chromatin Modifications,” Nucleic Acids Res. 41: 10032-10043 (2013), and Lai et al., “Activating RNAs Associate with Mediator to Enhance Chromatin Architecture and Transcription,” Nature 494:497-501 (2013), each of which is hereby incorporated by reference in its entirety). As described in Examples 14-23 infra, wMel Wolbachia and the Ci-causing CifA enzyme mediate depletion of a IncRNA that initiates changes in the histone-to-protamme exchange, driving histone retention on developing spermatids, protamine deficiency in mature sperm, and enhancement of wild type CI in D. melanogaster spermatocytes (Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal- effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety). Here, in the absence of Wolbachia and IncRNA depletion, a chemical disruption of histone acetylation and the downstream H-P exchange processes recapitulated the CI trait.

[0241] There is also increasing evidence that histone acetylation is involved in DNA damage repair (Dhar et al., “The Tale of a Tail: Histone H4 Acetylation and the Repair of DNA Breaks,” Philosophical Transactions of the Royal Society B: Biological Sciences 372(1731):20160284 (2017) and Hunt et al., “Histone Modifications and DNA Double-strand Break Repair After Exposure to Ionizing Radiations,” Radiat. Res. 179:383-392 (2013), which is hereby incorporated by reference in its entirety). As chromatin structure regulates the accessibility of DNA to a variety of different factors, H3 and H4 acetylation can modulate double-stranded DNA break repair by recruiting DNA repair enzymes or by facilitating homologous recombination, respectively (Horikoshi et al., “Pre-existing H4K16ac Levels in Euchromatin Drive DNA Repair by Homologous Recombination in S-phase,” Commun. Biol. 2:253 (2019) , which is hereby incorporated by reference in its entirety)- In addition, HAT- mediated H3 and H4 acetylation can facilitate the recruitment of non-homologous end-joining proteins to DNA damage sites (Ogiwara et al., “Histone Acetylation by CBP and p300 at Double-strand Break Sites Facilitates SWI / SNF Chromatin Remodeling and the Recruitment of Non-homologous End Joining Factors,” Oncogene 30:2135-2146 (2011), which is hereby incorporated by reference in its entirety). Consequently, the reduced acetylation is consistent with enhanced DNA damage by CifB enzyme in developing CI spermatids (Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383:1111-1117 (2024), which is hereby incorporated by reference in its entirety).

[0242] Once fertilization occurs, paternal chromatin undergoes chromatin remodeling such as protamines are removed from the paternal chromatin and replaced by maternal histones to decondense and activate the chromatin of the developing embryo (McLay & Clarke, “Remodelling the Paternal Chromatin at Fertilization in Mammals,” Reproduction 125:625-633 (2003), which is hereby incorporated by reference in its entirety). Interestingly, postfertilization delays in maternal H3.3 histone deposition occur in CI embryos45. It is proposed that the delay can be due to preloaded paternal H3 (and H4) histones causing altered paternal epigenome information that results in chromatin remodeling errors and mistiming of maternal H3.3 deposition. The symbiotic embryos may possibly rescue Ci-derived paternal chromatin errors by establishing similar maternal chromatin modifications during oogenesis due to Wolbachia or transgenic expression of the rescue factor CifA, as previously proposed (Kaur et al., “The Cif Proteins From Wolbachia Prophage WO Modify' Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 2 20(5): e3001584 (2022),Kaur et al., “The Mechanism of Cytoplasmic Incompatibility' is Conserved in Wolbachia- beanng Aedes aegypti Mosquitoes Deployed for Arbovirus Control,” PLoS Biol 22:e3002573 2024), and Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), each of which is hereby incorporated by reference in its entirety).

[0243] Overall, these data support, in an unprecedented manner, the Host Modification model of CI in which CI can be achieved by modifying a pre-fertilization, host factor without Wolbachia or its effector proteins CifA and CifB, Finally, it is noteworthy that natural CI strength vanes in Wolbachia-canying arthropods, some arthropods are refractory to Wolbachia, and some Wolbachia strains do not cause CI in vectors (Mercot & Charlat, “Wolbach a Infections in Drosophila melanogaster and D. simulans: Polymorphism and Levels of Cytoplasmic Incompatibility,” Genetica 120(l-3):51-9 (2004), which is hereby incorporated by reference in its entirety). Therefore, chemically reprogramming reproductive epigenetics in the absence of Wolbachia or Cif expression provides a new gateway to engineer this symbiotic trait crucial to arthropod biology and vector control programs.Example 11 - Methods for Examples 6-10

[0244] Fly rearing and strains. D. melanogaster stocks yhv* (BDSC 1495), UAS-HatlRNAi (BDSC 42488), UAS-HDACIRNAi (V30600), and Pasa-Ga / 4 (II, from Dr. Yukiko Yamashita, University of Michigan) were maintained on a 12-hour light / dark cycle at 25°C and 70% relative humidity on 50 ml of standard fly food media composed of agar, cornmeal, yeast, molasses, ethanol-dissolved tegosept, and propionic acid. Wolbachia-uninfected ylw* (wMel-) fly line was previously generated48 by feeding tetracycline-treated food (20 pg / ml in 50 ml of fly media) for three generations followed by two rounds of rearing on standard food media. Before using the lines for experimental purposes, their infection status was confirmed by PCR using Wolb_F and Wolb_R3 primers (Casiraghi et al., “Phylogeny of Wolbachia pipientis Based on gltA , groEL and ftsZ Gene Sequences : Clustering of Arthropod and Nematode Symbionts in the F Supergroup , and Evidence for Further Diversity in the Wolbachia Tree,” Microbiology (Reading) 15 l(Pt 12):4015-4022 (2005), which is hereby incorporated by reference in its entirety).

[0245] Chemical preparation and feeding. For chemical feeding assays, a stock solution of 28.69 mM anacardic acid (AA) (Merck Millipore, #172050) was prepared in dimethyl sulfoxide (DMSO) (Sigma, #D8418) solvent as previously described (Gartner et al., “Ex vivo Culture of Drosophila Pupal Testis and Single Male Germ-line Cysts: Dissection,Imaging, and Pharmacological Treatment,” Journal of Visualized Experiments 11 :(91):51868 (2014), which is hereby incorporated by reference in its entirety). In 10 ml standard fly food vials, 52.56 pl stock solution AA+DMSO) was added to reach the desired final concentration (150 pM) per vial. The same amount of DMSO solvent alone was mixed with standard food to create a ‘control’ chemical treatment condition in each experiment. To setup chemical -treated flies for conducting CI assays, wMel- paternal grandmothers were first mass-reared in standard 50 ml food bottles. Virgin females were collected and aged to 11 days before mating with wMel- males. Males and females in 4:4 ratio were transferred into 8 oz bottles each affixed with a grape-juice agar plate smeared with yeast. Following incubation at 250C for 24h, grape plates were discarded and replaced with freshly smeared ones and bottles were incubated again for 24h. The grape plates with eggs on them were collected and incubated for additional 24h to allow egg to larvae development. A total of 50 larvae at LI and L2 stage were manually picked and gently placed into AA-treated and control food vials each. Since AA is reportedly light-sensitive, the chemicals were mixed in the food under minimum light exposure the same morning of larvae transfer. After mixing, the vial trays were covered with aluminum foil and stored and processed in dark until the larvae transfer process started. In parallel, we also setup wMel+ flies to use as positive CI control by rearing them on the same standard fly food media lacking the chemicals. Food was manually churned using a spatula and larvae were manually picked and transferred to the churned food vials similar to chemical-mixed food to avoid any confounding effect.

[0246] Hatch rates. Male flies hatching between 0-8 hours were collected and used in hatch rate (HR) assays to ensure higher CI penetrance. HR set up was used as described previously. Briefly, a male and female pair was placed in an 8 oz, round bottom, polypropylene Drosophila stock bottle with a yeast-smeared grape juice agar plate affixed. The bottles were placed in a 25°C incubator overnight to allow mating. Following day, these plates were discarded and replaced with new grape juice agar plates with fresh yeast smear. After an additional 24 hours, the plates were removed, and the total number of embryos were counted. The embryo plates were then incubated for 36 hours at 25°C before the number of unhatched embryos were counted. Any plates with less than 20 embryos laid were discarded from the analyses. Significant differences (p < 0.05) were determined by pairwise Mann Whitney U tests or by a Kruskal-Wallis test and Dunn multiple test correction in GraphPad Prism 10.

[0247] Immunofluorescence. Sibling males from the HR assay (0-4 hours old) were collected for testes dissection in ice-cold 1 x PBS solution. Tissues were fixed in 4% formaldehyde and processed for immunostaining as per our previous study. Briefly, samples were washed, blocked in BSA, and incubated with primary' antibodies overnight at 4°C. Tissues were then incubated with secondary antibodies for 4 hours at room temperature in the dark. Tissues were then washed and mounted on slides using a DAPI-containing Vectashield medium. Imaging was performed using Zeiss LSM880 confocal microscope under constant exposure settings across treatment groups. Images processing and quantification was performed using ImageJ software.

[0248] Histone-specific antibodies used were rabbit polyclonal a) anti-Acetyl-Histone H3 (EMD Millipore, Cat. No. 06-599, 1:500 dilution) recognizing acetylation on K9 and K14 lysine residues, b) anti-acetyl-Histone H4 (EMD Millipore, Cat. No. 06-598, 1 :5000 dilution) recognizing acetylation of K5, K8, KI 2, and KI 6 residues, and c) anti -Histone H3 (EMD Millipore, Cat. No. 06-755, 1: 100 dilution). Core histones were detected with mouse monoclonal anti-Histone antibody (EMD Millipore, Cat. No. F152.C25.WJJ, 1: 1200 dilution). Anti-mouse alexa488 and anti-rabbit alexa594 conjugated secondary' antibodies (Vector Laboratories) were used at 1 :500 dilutions. Rabbit polyclonal anti-Histone H4 antibody (EMD Millipore, Cat. No. 07-108) were also tested to detect H4 histone retention in multiple replicates, however, the antibody did not work.

[0249] Mature sperm isolation, CMA3 staining, and quantification. Sperm from seminal vesicles of 0-4 hours old sibling HR males were extracted and processed for CMA3- based staining assay as described previously (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5) :e3001584 (2022), which is hereby incorporated by reference in its entirety). Fluorescence quantification was performed by scoring fluorescent pixels in arbitrary units (A.U.) within individual sperm head using ImageJ software. The calculated fluorescence intensity per sperm head was graphed. Statistical significance (p < 0.05) was determined by a pairwise comparison using Mann-Whitney test in GraphPad Prism 10.Example 12 - Effect of HAT Inhibitor Treatment and CI in Drosophila simulans

[0250] Experiments were conducted to determine if histone retention during late spermatogenesis a cause of CI across Drosophila species. The hypothesis that feeding larvae with anacardic acid mixed into standard fly food medium will inhibit the activity of histone acetyltransferase enzy me in the elongating stages of spermatogenesis to block the histone-to-protamine chromatin switch, causing histones to be retained on chromatin rather than removed, and recapitulate CI was tested.

[0251] In Drosophila, the paternal genome is condensed into a protamine-based chromatin structure (Awe & Renkawitz-Pohl, ‘‘Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster,” SystBiol Reprod Med 56:44-61 (2010), which is hereby incorporated by reference in its entirety). This condensing process occurs by replacing the histone-based chromatin with protamines during spermiogenesis and is necessary for the formation for fertile sperm (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster,” SystBiol Reprod Med 56:44-61 (2010), which is hereby incorporated by reference in its entirety). This transition occurs during the early to late canoe nuclei stage and is initially marked by an increase in H3 and H4 acetylation by its corresponding histone acetyltransferase (HAT) (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster,” SystBiol Reprod Med 56:44-61 (2010), which is hereby incorporated by reference in its entirety). Anacardic acid (AA) is an inhibitor of HAT and has been shown to block the histone to protamine switch in Drosophila pupal testes, leaving the sperm with a histone-bound chromatin. Importantly, AA does not induce apoptosis nor is it cytotoxic to the host since post-switch cysts develop normally (Awe & Renkawitz-Pohl, “Histone H4 Acetylation is Essential to Proceed from a Histone- to a Protamine-based Chromatin Structure in Spermatid Nuclei of Drosophila melanogaster,” SystBiol Reprod Med 56:44-61 (2010), which is hereby incorporated by reference in its entirety).

[0252] Males expressing transgenic cifAB (nos>cifA;B) and wild type Wolbachia (M’Mel+) abnormally retain histones (Kaur et al., “The Cif Proteins From Wolbachia Prophage WO Modify Sperm Genome Integrify to Establish Cytoplasmic Incompatibility,” PLoS Biol 2 20(5):e3001584 (2022), Kaur et al., “The Mechanism of Cytoplasmic Incompatibility is Conserved in WolbachiaWtQwmg Aedes aegypti Mosquitoes Deployed for Arbovirus Control,” PLoS Biol 22:e3002573 2024), and Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), each of which is hereby incorporated by reference in its entirety). As described in Examples 1-11, the wMel+ and cifAB phenotype was mimicked in wMel- flies to artificially create the sperm with histone retention ability to cause CI (see FIG. 2B).

[0253] These results indicate that AA can chemically program rescuable CI in D. melanogaster system. AA was tested to determine if it could also induce similar changes in a different host system: D. simulans in the absence of its native Wolbachia strain ’w Ri'. If CI could be recapitulated using AA treated w RiT males that can be rescued using wRi infected females, it would be concluded that an impaired histone-to-protamine transition is a conserved feature across host systems necessary for the establishment of CI.

[0254] As shown in FIG. 10, an initial test was conducted for a wRi hatch rate test using wild type wRi carrying D. simulans males and chemically treated n R.iT males using AA and dimethylsulfoxide (DMSO) (lx; 150 pM concentration). In the AA crosses, complete CI (0% embryonic hatching in the 150pM AA wRiT x wRiT cross) and rescue (50% embryonic hatching in the 150pM AA H RIT x H RI+ cross) were obtained. In the wild type CI and rescue control crosses we were able to cause complete CI (0% embryonic hatching in the wRi+ x wRiT cross) and rescue (80% embryonic hatching in the wRi+ x u Ri+ cross), respectively. In this experiment, hatching was not observed in the two DMSO crosses, likely because a higher concentration was mistakenly used. The results of this experiment are shown in FIG. 10, with a cutoff point of less than 20 eggs.

[0255] The number of adults eclosed was also tracked to determine the effects of chemical treatment on survivability (FIGs. 11A-B). Calculations were made of survivability for each sex (FIG. 11 A) and total individuals (FIG. 11B). Specifically, the calculations were made by: (A) dividing the number of emerged males and females by 25 (50% of the total number of larvae added to each vial, assuming there is a 1 : 1 maleTemale sex ratio), then multiplying by 100 to convert the value into a percentage; and (B) dividing the total number of emerged adults by 50 (total number of larvae added to each vial), then multiplying by 100 to convert the value into a percentage.

[0256] Males and females had comparable survival within each treatment group (FIG. 11A; ANOVA. p = 0.73). Indeed, an average of 47.2% (± 15.9% SD) males and 46.8% (± 12.7% SD) females survived on diets with 150 pM AA. Similarly, 51% (± 18.8% SD) of males and 49% (± 15.2% SD) of females survived, on average, to adulthood when reared on diets treated with 150 pM DMSO. No differences in survivorship were found across chemical treatments (FIG. 1 IB; Mann Whitney U-test, p = 0.17), albeit a higher concentration of DMSO was used in this experiment. Specifically, on average, 47% (± 8.9% SD) and 50% (± 15.4% SD) of total flies survived in the AA and DMSO treatments, respectively. Thesefindings indicate that AA doesn’t induce additional toxicity to host fitness / survivorship than DMSO alone.

[0257] In a second experiment, three lower concentrations of AA, including 112.5 pM (0.75x), 75 pM (0.50x), and 37.5 pM (0.25x) were tested, because the AA rescue obtained in the first experiment was 30% lower than the control rescue cross. The results of the second experiment using lower AA concentrations are shown in FIG. 12. Here, a cutoff point of 10 eggs was used because very few females laid more than this amount of eggs. In crosses using 112.5 pM AA, complete CI (0% embryonic hatching) and rescue (-57% embryonic hatching) were obtained. Similar results were obtained when using 37.5 pM AA: only 5% embryonic hatching in the CI cross that was rescuable to some degree (-35% embryonic hatching). However, when using 75 pM AA, chemically induced CI was not rescuable, this was unexpected given there was rescue to some degree using higher 112.5 pM dose. The DMSO rescue controls produced 35%, 30%, and 90% embry onic hatching when using 112.5 pM, 75 pM, and 37.5 pM concentrations, respectively, overcoming the prior issue by using the correct DMSO concentrations. However, repeating the assay with more sample size will strengthen the statistical comparison across groups.

[0258] Once again, the number of adults eclosed was tracked to determine the effects of chemical treatment on survivability as shown in FIGs. 13A-B.

[0259] In this experiment, the percent of males and females that survived was similar within each treatment (FIG. 13A; ANOVA, p = 0.75). Furthermore, the percent of adults that eclosed was comparable when comparing chemicals at the same concentration (FIG. 13B; Bonferroni’s correction, / ? > 0.05). For example, an average of 54.2% (± 15.2% SD) and 54% (19.0% SD), of flies survived when reared on food containing 112.5 pM AA and DMSO, respectively. While it was found that the chemical concentration had an significant overall effect on fly survivorship (FIG. 13B; GLM with a beta error distribution, p < 0.05), pairwise comparisons were not significantly different between concentrations of the same chemical (FIG. 13B; Bonferroni’s correction, p > 0.05). For instance, mean percent survival varied by less than 6% across all AA concentrations. It is suggested that these patterns were driven by the unusually high mean percent survival in the 37.5 pM DMSO, as the only significant pairwise comparison was between 37.5 pM DMSO and 75 pM AA (Bonferroni’s correction, / ?< 0.05). Indeed, the mean percent survival was 56.1% (± 10.9% SD), 56.8% (± 12.6% SD), and 54% (± 19% SD) for the 37.5 pM, 75 pM, and 112.5 pM DMSO treatments, respectively.Taken together, these findings indicate that varying AA had no additional toxic effect on host fitness / survivorship beyond DMSO. Furthermore AA-induced CI was also shown in Z). simulans.Prophetic Example 13 - Anacardic Acid-induced CI in Aedes aegypti mosquitoes without Wolbachia

[0260] Experiments will be conducted to determine if feeding wMelM- Aedes aegypti larvae with Anacardic Acid (AA) would inhibit the activity of Histone acetyltransferase (HAT) enzyme in the elongating stages of spermatogenesis to block the histone to protamine chromatin switch and recapitulate CI.

[0261] - wMelM+ and wMelM- Aedes aegypti mosquito colonies will be reared in theMcGraw Lab insectary at the Orchard Rd. All colonies and lines will be maintained at temperature of 25 ± 2° C, relative humidity of 75 ± 5%, and a 12-h light / dark cycle.

[0262] Infection by Wolbachia was confirmed using the following protocol. Single individuals were collected in 2ml Eppendorf tube from both Hoff wMel (wMelM+) and Hoff WT (wMelM-) Ac des aegypti mosquito lines and DNA was isolated. PCR screening was performed using primers (Table 1) amplifying a fragment of the WSP gene of wMelM Wolbachia and with primers amplifying a fragment of ribosomal protein S17 (RPS17) from Ae. aegypti (305 bp). The RPS17 primers were designed to act as a control for the presence of amplifiable DNA or any PCR inhibition in the reaction. The PCR program was 98°C for 3 min, 35 cycles of 98°C for 30 s, 60°C for 30 s and 72°C for 40 s and ended with 72°C for 5 min.Table 1. PCR Primers for Detection of Wolbachia Infection

[0263] An initial experiment was conducted. The protocol for setting up fathers and mothers to conduct cytoplasmic incompatibility assay was as follows. An initial AA dose (lx = 0.0234 pg / ml) was used in mosquitoes. A report by Pinto et al. suggested that this concentration induced minimum larvicidal activity (10-30%) (Pinto et al, “Larvicidal Activity, Aquatic and in vivo Toxicity of Anacardic Acid Loaded-Zein Nanoparticles,” Journal of Drug Delivery Science and Technology 63: 102513 (2021), which is hereby incorporated byreference in its entirety). However, when tested at this concentration of AA, no mortality was observed because out of total larvae transferred in AA-mixed water (550), 50.18% adult males hatched ((276 / 550)* 100). The rest were females.

[0264] Increased doses of AA will be tested to determine their impact on males' ability to induce CI. In the next experiment, the dose of AA will be increased by 2.5 and 5 times, including Dose 1 (25x) = 0.585 pg / ml and Dose 2 (5 Ox) = 1.17 pg / ml. Note that Pinto et al. showed 100% larval mortality using a 0.1875 pg / ml dose of AA. Higher doses remain to be tested.

[0265] Anacardic acid is made by dissolving lOmg AA powder in 10 ml of DMSO to make 1 mg / ml stock concentration. Pipette 292.5 pl of 1 mg / ml stock AA solution and dissolve in 500ml of water to make working concentration = 0.585 pg / ml. 585.0 pl of 1 mg / ml stock AA solution will be pipetted and dissolved in 500ml of water to make working concentration = 1.17 pg / ml. The stock DMSO concentration is 14.1M or l. lg / ml. The concentration of DMSO that goes into AA solution above is Cl x VI = C2 x V2. l. lg / ml x 292.5 pl = C2 x 500ml C2 = 643.5 pg / ml. Thus, to use equal DMSO concentration in control treatments, 292.5 pl of 1.1 g / ml stock DMSO solution will be pipetted and dissolved in 500ml of water to make dose 1 working concentration = 643.5 pg / ml. 585.0 pl of 1.1 g / ml stock DMSO solution will be pipetted and dissolved in 500ml of water to make dose 2 working concentration = 1287.0 pg / ml.

[0266] Father setup timeline: Egg papers from both wMelM- and wMelM+ lines will be placed in DI water and allowed to develop into LI and L2 larvae. wMelM- LI and L2 larvae (100 total) will be transferred into 500ml of dose 1 AA- vs. DMSO- treated water containers (n=5). In parallel, wMelM- LI and L2 larvae (100 total) will be transferred into 500ml of dose 2 AA- vs. DMSO- treated water containers (n=5). In parallel, wMelM+ LI and L2 larvae (100 total) will be transferred into 500ml of untreated water container (n=5). Developed pupae will be transferred from above water containers to square-shaped tupperware with 100 ml., of water to fit into metal cages. Collect hatched males and also count them. Perform mating in 10: 10 (males: females) group setting in large 60oz paper cups. Leave a cotton ball soaked in 10% sucrose solution on top of the cup lid and cover with a small petri dish. Let the mosquitoes mate for 6 days. Cotton will be changed every 3 days. Post-mating, collect only females and release them in big metal cages for blood-feeding (BF) in group. Let them starve for a day before BF. BF females in the morning. Post BF, give them 1-2 hrs to just relax and settle down. Then after, sort BF females from non-BF ones (BF females have a red,bloated belly compared to a flat, non-bloated ones from non-BF females), and put them in 60oz paper cups in a group of 10 to lay eggs for 5 days. For egg laying, a small (20ml capacity) plastic cup filled with 5ml of room temperature water and a strip of filter paper will be added inside the 60oz cup. Collect egg papers and count total eggs. Dry them for 2 days. Place the eggs papers in deoxygenated water containing 0.25 g / L yeast extract for 24 hours to hatch. Count total hatched eggs (that have a clearly detached cap) and calculate % embryonic hatching to measure CI levels. Determine the statistical significance (p<0.05) by a Kruskal- Wallis test and Dunn’s multiple test correction in GraphPad Prism.

[0267] Mother setup timeline -without any treatment: Hatch egg papers from wMelM- line in DI water container and let them develop into larvae and pupae. Developed pupae will be transferred from above water container to square-shaped tupperwares with 100 mL of water to fit into metal cages. Collect hatched virgin females and use them to mate with fathers above in a 10: 10 group setting.Table 2. CI hatch rate crossings

[0268] It is expected that wMelM-; AA males will induce CI similarly or comparably to that of the positive control wMelM+ (dH2O) resulting in reduced embryonic hatching. A direct correlation between the AA dose used in males to the CI levels induced is expected to be obtained. In comparison to wMelM-; AA males, it is expected that wMelM-;DMSO males will not or only slightly have reduced embryonic hatching as DMSO is a control chemical solvent used to dissolve AA.Example 14 - Methods for Examples 15-23

[0269] Fly rearing and strains. D. melanogaster stocks b * (BDSC #1495), nos- GAL4:VP16 (BDSC #4937), UAS transgenic (TG) lines homozygous for cifA, cifB, cifA;B,and cif mutants (Barr, J. J., “Missing a Phage: Unraveling Tripartite Symbioses within the Human Gut,” mSystems 4(3):e00105-19 (2019) and Weinert et al., “The Incidence of Bacterial Endosymbionts in Terrestrial Arthropods,” Proceedings of the Royal Society B: Biological Sciences 282(1807):20150249 (2015), each of which is hereby incorporated by reference in its entirety), AAGAG and Scramble RNAi knockdown fly lines (Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility in Drosophila simulansf Genetics 126(4): 933-48 (1990), which is hereby incorporated by reference in its entirety) were maintained at 12: 12 light: dark at 25°C and 70% relative humidity on 50 ml of standard cornmeal- and molasses-based food medium. Lines without Wolbachia were previously generated (Barr, J. J., “Missing a Phage: Unraveling Tripartite Symbioses within the Human Gut,” mSystems 4(3):e00105-19 (2019), which is hereby incorporated by reference in its entirety) through tetracycline treatment for three generations. Wolb_F and Wolb_R3 primers were used to confirm symbiont presence before conducting experiments. Virgin flies were collected and stored at room temperature.

[0270] In vitro nuclease assay. Codon optimization, gene synthesis, cloning, and protein expression / purification were outsourced to GenScript Biotech (New Jersey, USA). Briefly, cifA and cifB genes were codon-optimized for translation and expression in E. coli, de novo synthesized, cloned into pGS-21a expression vector, and transformed into E. coli BL21 (DE3) competent cells. Cultures were grown in 1 L TB medium containing ampicillin and were incubated in 37°C at 200rpm. Once cell density reached to 1.2 O.D. at 600 nm, 0.5 mM isopropyl-b-D-thiogalactoside (IPTG) was introduced for induction at 15°C for 16 hours and then centrifuged at 8000 g for 20 min.

[0271] For CifA, cell pellets were lysed on ice using buffer A (50 mM Tris, 150 mM NaCl, 1 mM TCEP, pH 8.0) by sonication. The lysate was clarified by centrifugation at 12000 g for 30 min at 4°C. The clarified extracts of target CifA proteins carrying His tags at the C- terminus were filtered with a 0.45 pm filter and loaded onto Ni-nitrilotriacetic acid (NTA) agarose resin preequilibrated in buffer A. The column was then washed with buffer B (50 mM Tris, 150 mM NaCl, pH 8.0) 10 column volumes (CV) to wash away impurities from the target proteins. The protein was eluted using buffer B supplemented with 20 / 50 / 300 mM imidazole. All proteins were sterilized by passing through 0.22 pm filter before being stored in aliquots. The concentration was determined by BCA™ protein assay with BSA as a standard. SDS-PAGE and Western blot were used to confirm protein purity and molecular weight.

[0272] For CifB, cell pellets were lysed on ice using buffer A (50 mM Tris, 150 mM NaCl, pH 8.0). The lysate was clarified by centrifugation at 12000 g for 30 min at 4°C. The inclusion body pellet was solubilized in denature buffer C (7M Gu-HCl, 50 mM Tris-HCl,150 mM NaCl, pH 8.0) by sonication. The cell precipitate was spun down at 13,000 rpm for 30 min at 4°C, then the supernatant including target CifB proteins carrying His-GST tag at the N- terminus was filtered with a 0.45 pm filter and loaded onto NTA agarose resin preequilibrated in buffer A. The column was then washed with buffer D (8 M Urea, 50 mM Tris-HCl, pH 8.0) 10 CV to wash away impurities from the target proteins. The protein was eluted using buffer D supplemented with 20 / 50 / 300 mM imidazole. All proteins were sterilized by passing through 0.22 pm filter before being stored in aliquots. The concentration was determined by BCA™ protein assay with BSA as a standard. SDS-PAGE and Western blot were used to confirm protein purity and molecular weight.

[0273] Additionally, mass spectrometry-based protein identification was performed to ensure no contaminant nuclease from the E. coll expression system was co-purified. Briefly, 20 pg of purified proteins (CifA, CifAAC, CifB AD, and CifB2;AD) were prepared for analysis using Suspension trap technology (Brucker & Bordenstein, ‘‘Speciation by Symbiosis,” Trends Ecol Evol 27:443-51 (2012), which is hereby incorporated by reference in its entirety) using the manufacturer protocol. The resulting peptides were analyzed by a 70-minute data- dependent LC-MS / MS analysis. Briefly, peptides were auto sampled onto a 200 mm by 0. 1 mm (Jupiter 3 micron, 300A), self-packed analytical column coupled directly to an LTQ (ThermoFisher) using a nanoelectrospray source and resolved using an aqueous to organic gradient. A single full-scan mass spectrum followed by 5 data-dependent tandem mass spectra (MS / MS) was collected throughout the run and dynamic exclusion was enabled to minimize the acquisition of redundant spectra. The Resulting MS / MS spectra were searched via SEQUEST against a database containing the expressed proteins, an E. colt background proteome and reversed version for each of the entries. Identifications were filtered and collated at the protein level using Scaffold Proteome Software.

[0274] In vitro nuclease activity assays were performed as previously described (5). For DNase and RNase activity measures, IpM of individual Cif proteins were incubated in a reaction buffer containing 20 mM Hepes (pH 8.0), 5 mM MgCh, 2.5% sucrose, 150 mM NaCl, 0.001% Triton X-100, and 2 mM DTT with 500 nM single-stranded (ss) Cy5-labeled DNA [70-mer: Cy5- GCAATTCGATCGTTGACATCTCGCGTGCTCGGTCAATCGGCAGATGCGGAGTGAAGTTCCAACGTTCGGC-3] (SEQ ID N0:17) as previously used (Caputo et al., "A Bacterium Against the Tiger: Preliminary Evidence of Fertility Reduction After Release of Aedes albopictus Males with Manipulated Wolbachia Infection in an Italian Urban Area,” PestManag. Sci. 76: 1324- 1332 (2020), which is hereby incorporated by reference in its entirety); 15nM double-stranded (ds) 154bp PCR purified fragment of rp49 gene (Zheng et al., “Incompatible and Sterile Insect Techniques Combined Eliminate Mosquitoes,” Nature 572:56-61 (2019), which is hereby incorporated by reference in its entirety); or lOOnM of synthetic RNA [45-mer: 5’ GGGUCAACGUGGGCAAAGAUGUCCUAGCAAGCCAGAAUUCGGCAG -3’] (SEQ ID NO:18) generated by Sigma. In reactions where CifA was co-present with CifB, 10 pM CifA was used as previously described (Caputo et al., “A Bacterium Against the Tiger: Preliminary Evidence of Fertility Reduction After Release of Aedes albopictus Males with Manipulated Wolbachia Infection in an Italian Urban Areaf Pest Manag. Sci. 76: 1324-1332 (2020), which is hereby incorporated by reference in its entirety). All reactions were carried out at 25°C for 120 min and quenched by adding EDTA to a final concentration of 100 mM unless otherwise noted. Samples were run in 10% TBE polyacrylamide urea gels at 180 V for 60 min. For reactions using Cy5-labeled ssDNA, gels were imaged on the Odyssey CLx imaging system. Unlabeled dsDNA and ssRNA sample gels were post-stained with GelRed (Biotium) stain and imaged with Alpha innotech imager.

[0275] In situ TUNEL assay. Terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling (TUNEL) assays are based on detecting single- and double-stranded DNA nicking and fragmentation, which are characteristic of apoptotic cells (Nazni et al., “Establishment of Wolbachia Strain wAlbB in Malaysian Populations of Aedes aegypti for Dengue Control,” Current Biology 29(24):4241-4248 (2019), which is hereby incorporated by reference in its entirety). To perform the assay and detect sperm DNA fragmentation, we first set up the flies as previously described (Weinert et al., “The Incidence of Bacterial Endosymbionts in Terrestrial Arthropods,” Proceedings of the Royal Society B: Biological Sciences 282(1807):20150249 (2015), which is hereby incorporated by reference in its entirety). Briefly, virginity-controlled wild type (wMel+ and wMel-) and TG ( s-Gal4:VP I 6) females were aged 9-11 days and mated with males (Indriani et al., “Reduced Dengue Incidence Following Deployments of IFo / ftac / zza-Infected Aedes aegypti in Yogyakarta, Indonesia,” Gates Open Res. 4:50 (2020), which is hereby incorporated by reference in its entirety). We used -Ga!4:VP 16 line since it was previously shown to drive cifA;B expression sufficient to induce near-complete embryonic death (Shropshire et al., “Initiative,Symbiont-mediated Cytoplasmic Incompatibility : What Have We Learned in 50 Years?,” Elife 1-51 (2020), which is hereby incorporated by reference in its entirety). We collected <8 hours old wild type and TG Gal4-UAS males as the young, hatched males induce strong CI levels (LePage et al., “Prophage WO Genes Recapitulate and Enhance Wolbachia-induced Cytoplasmic Incompatibility,” Nature 543:243-247 (2017) and Beckmann et al., “A Wolbachia Deubiquitylating Enzyme Induces Cytoplasmic Incompatibility,” Nat Microbiol 2: 17007 (2017), each of which is hereby incorporated by reference in its entirety), anesthetized on ice to stop their movement, and dissected whole testes in ice-cold IX PBS solution. Dissected tissues were treated with 2 mM dithiothreitol for 45 mm at room temperature, followed by fixation in 2% paraformaldehyde on ice for 15 min. After washing in IX PBS for 2 min, samples were permeabilized in 0.1% TritonX-100 in sodium citrate (10 mg sodium citrate, 10 ml Triton, 10 ml milliQ H2O) for 2 min on ice. After washing in IX PBS for 2 min, samples were incubated with 50 ml mix of 5 ml enzy me and 45 ml labeling solution (TUNEL In situ Cell Death Detection Kit, Fluorescein, Cat. No. 11 684 795 910 from Roche) for 1.5 h at 37°C in a dark humid chamber. After washing in IX PBS for 2 min, samples were finally incubated with 50 mL of DAPI staining solution (0.2 mg / ml), mounted on a glass slide, squashed wi th a coverslip, and stored overnight at 4°C. Imaging was performed using green fluorescence filter excited at 488 nm laser for TUNEL and blue at 359 nm for DAPI stain at lOOx magnification in All-in-one Keyence BZ-X700 fluorescence microscope. Image exposure settings were kept constant throughout the treatment groups and images were analyzed using ImageJ software. The total number of sperm bundles and TUNEL-positive bundles with damaged DNA were manually counted per testes. A complete H-P replacement involves stage-specific histone modifications (Shropshire et al., “One Prophage WO Gene Rescues Cytoplasmic Incompatibility in Drosophila melanogaster,” Proc Natl Acad Sci U SA 1 15:4987— 4991 (2018), which is hereby incorporated by reference in its entirety). Histone H4 acetylation (AcH4), for example, occurs in early canoe (EC) stage nuclei and disappears in late canoe (LC) stage when protamines are deposited (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). TUNEL-based DNA breaks are more pronounced in LC spermatids (Yu et al., “Enzyme Function Prediction Using Contrastive Learning,” Science 379(6639): 1358-1363 (2023), which is hereby incorporated by reference in its entirety). To discriminate between EC and LC spermatids, we labeled histones using the AcH4 antibody.To combine TUNEL and antibody staining, we followed previously established protocols (Yu et al., “Enzyme Function Prediction Using Contrastive Learning,” Science 379(6639): 1358— 1363 (2023) and Murray et al., “Sequence-specific Cleavage of RNA by Type II Restriction Enzymes,” Nucleic Acids Res. 38:8257-8268 (2010), each of which is hereby incorporated by reference in its entirety). The experiment was performed with three independent biological replicates and samples were blind-coded for the first run.

[0276] Fluorescent in situ hybridization (FISH) assay. AAGAG IncRNA expression peaks from spermatogonia to primary spermatocyte stage during Drosophila spermatogenesis (Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility in Drosophila simiilans." Genetics 126(4):933-48 (1990), which is hereby incorporated by reference in its entirety). Knocking AAGAG RNA down using spermatocyte-specific Bam- Gal4 driver (Chen et al., “A Wolbachia Nuclease and its Binding Partner Provide a Distinct Mechanism for Cytoplasmic Incompatibility,” Proc. Natl. Acad. Sci. USA 116:22314-22321 (2019), which is hereby incorporated by reference in its entirety) catastrophically altered the histone-to-protamine transition and rendered males fully infertile, whereas expressing it earlier in germline and spermatogonia using «ov-Gal4:VP16 driver did not(Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility in Drosophila simulans,” Genetics 126(4): 933-48 (1990), which is hereby incorporated by reference in its entirety). To decouple AAGAG relevance to CI from male sterility, we used ms-Gal4: VP 16 driver to knockdown AAGAG RNA in both IVo / bachia-mfecled (AAGAG_KD+) and uninfected (AAGAG_KD-) wild type lines. Virgin <8 hr old males were dissected to isolate testes as described above. Samples were then fixed in 3.7% formaldehyde in IX PBS followed by washing three times for 10 min each in IX PBS with 0.03% Triton-X 100 (PBS-T). PBST was replaced with absolute ethanol and samples were permeabilized overnight under constant agitation at 4°C. Next day, tissues were washed in freshly prepared washing buffer (2X saline sodium citrate SSC, 10% deionized formamide, nuclease-free water) for 5 mins at RT. Samples were then hybridized overnight by incubating in dark chamber at 37°C in 50 pl hybridization buffer (0.1% dextran sulfate, 2X SSC, 10% deionized formamide, and nuclease-free water) containing 0.5 pl of 3 pmol of AAGAG RNA probe as described previously (Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility in Drosophila simulans ” Genetics 126(4): 933-48 (1990), which is hereby incorporated by reference in its entirety). Post-hybridization, tissues were rinsed quickly in washing buffer at room temperature (twice for 5 min each), followed by two washes for 30 min each at 37°C. After washing, tissues wereequilibrated in IX PBS with 0.01% Triton-X 100 for 5 min at room temperature and then mounted on glass slide using Vectashield DAPI-containing medium. Samples were imaged using Zeiss LSM880 confocal microscope using constant exposure settings across treatment groups. Images were processed using ImageJ software and analyzed using “Analyze particles” function to measure the total area and integrated density for calculating RNA transcript signal intensity in spermatocytes. Each individual spermatocyte per testes was quantified during the image processing. Location of representative spermatocytes is shown in FIG. 21 with circled white periphery in individual channels. The experiment was performed with two independent biological replicates and samples were blind-coded for the first run.

[0277] Embryonic DNA damage assay. Embryos were staged according to D. melanogaster embryogenesis (Sun et al., “The CinB Nuclease from w No Wolbachia Is Sufficient for Induction of Cytoplasmic Incompatibility in Drosophila "' mBio 13 (2022), which is hereby incorporated by reference in its entirety). Embry os were collected from grape juice agar plates and dechori onated in 2.8% sodium hypochlorite solution, subsequently fixed in a 1 :1 mixture of heptane and 4% paraformaldehyde (PFA) diluted in IX phosphate buffered saline (PBS) rotating for 20 minutes. Heptane and PFA were allowed to separate before the bottom PFA phase were carefully removed. Methanol was added to the remaining heptane, and the tube was shaken vigorously for 20 seconds before the embryos settled to the bottom and solution was removed. Embryos were washed in methanol and rehydrated through an ethanol series (95, 70, 50 and 35%) and then washed in PBT (PBS containing 0.1% Triton X- 100). After treatment with a blocking buffer (PBT containing 3% bovine serum albumin) for 60 min at room temperature, the embryos were incubated with mouse anti-pH2Av primary antibody (1 :1000 DSHB #UNC93-5.2.1) at 4°C overnight, washed three times in PBT and incubated with secondary antibody (1:500 Thermo Fisher Scientific, Goat anti-mouse Alexa Fluor 594 conjugate) at room temperature for 4 hours. Antibodies were diluted in the blocking buffer. Stained embryos were washed three times in PBT, mounted in DAPI-containing Vectashield mounting media, and observed under Zeiss LSM 880 confocal microscope. The experiment was performed with two independent biological replicates and samples were blind-coded for the first run.

[0278] Hatch rate assays. Male siblings from TUNEL and FISH assays were used to measure CI hatch rate levels as previously described (7). Briefly, males and females were paired in 8 oz bottles affixed with a grape-juice agar plate smeared with yeast. Bottles were incubated at 25°C for 24 hours at which time the plates were replaced with freshly smearedplates and again stored for 24 hours. Plates were then removed from bottles, and the numbers of eggs on each plate were counted. Any crosses with fewer than 25 eggs laid were discarded from the count. After another 30 hours incubation at 25°C, the remaining unhatched eggs were counted. The percent of eggs hatched into larvae was calculated by dividing the number of hatched eggs by the total egg count and multiplying by 100.

[0279] In silico prediction analysis. Protein sequence alignment of CifB orthologs from different Wolbachia strains (NCBI accession number, CifB T1 wMel - WP-010962721.1, T1 w Pip - WP_012481788.1, T4 M’Pip - WP_007302979.1) and CifA T1 H’Mel (WP 010962721.1) were performed using the MUSCLE plugin (Lindsey et al., “Evolutionary Genetics of Cytoplasmic Incompatibility Genes cifA and cifB in Prophage WO of Wolbachia ” Genome Biol Evol 10:434-451 (2018), which is hereby incorporated by reference in its entirety) in Geneious Prime v2021.0.3 (Wang et al., “Crystal Structures of Wolbachia CidA and CidB Reveal Determinants of Bacteria-induced Cytoplasmic Incompatibility and Rescue,” Nat Commiin 13(1): 1608 (2022), which is hereby incorporated by reference in its entirety)- Secondary structure predictions of CifA and CifB protein sequences were made using the PSIPRED Protein Sequence Analysis Workbench program (Singleton et al., “Crystal Structure of RecBCD Enzyme Reveals a Machine for Processing DNA Breaks,” Nature 432 (7014): 187-93 (2004), which is hereby incorporated by reference in its entirety). We manually curated the presence of the QxxxY motif based on its localization within a region of predicted a-helices in the nuclease domains in CifB and throughout the length of CifA proteins.

[0280] Statistical analysis. All statistical analyses were performed using GraphPad Prism 9 software. While comparing in situ TUNEL data between two groups, we used a two- tailed, non-parametric Mann-Whitney U-test. For comparisons between more than two data sets, we used a non-parametric Kruskal-Wallis one-way analysis of variance test followed by a Dunn’s multiple correction. This allowed robust testing between all data groups while correcting for multiple test biases. For CI hatch rate assays, statistical significance was determined by Kruskal -Wallis and Dunn’s multiple correction tests. All P-values are reported in Tables S1-S4 and raw data files related to each experiment are included in the supplementary files of Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety.Example 15 - Prophage Proteins of an Insect Symbiont Modulate Sperm Noncoding RNA and DNA to Kill Embryos

[0281] The extent to which bacteriophage proteins in symbiotic bacteria modulate eukaryotic macromolecules has largely been overlooked, except for some virulence proteins and observations of physical interactions within eukaryotic cells (Barr, “Missing a Phage: Unraveling Tripartite Symbioses within the Human Gut,” mSystems 4(3):e00105-19 (2019), which is hereby incorporated by reference in its entirety). A prophage-containing bacterial symbiont, Wolbachla pipte iis. inhabits the reproductive tract of approximately half of all arthropod species worldwide (Weinert et al., “The Incidence of Bacterial Endosymbionts in Terrestrial Arthropods,” Proceedings of the Royal Society B: Biological Sciences 282(1807):20150249 (2015), which is hereby incorporated by reference in its entirety) and selfishly alters host reproduction to increase the relative number of symbiotic females that transmit the bacteria to the next generation (Hoffmann et al., “Factors Affecting the Distribution of Cytoplasmic Incompatibility inDrosophila simulans,” Genetics 126(4):933-48 (1990), which is hereby incorporated by reference in its entirety). Cytoplasmic incompatibility (CI) is the most commonly studied reproductive alteration that impacts arthropod evolution (Brucker & Bordenstein, “Speciation by Symbiosis,” Trends Ecol Evol 27:443-51 (2012), which is hereby incorporated by reference in its entirety) and major vector control efforts for population suppression and replacement strategies (Caputo et al., “A Bacterium Against the Tiger: Preliminary Evidence of Fertility Reduction After Release of Aedes albopictus Males with Manipulated Wolbachia Infection in an Italian Urban Area,” PestManag Sci 76:1324- 1332 (2020), Zheng et al., “Incompatible and Sterile Insect Techniques Combined Eliminate Mosquitoes,” Nature 572:56-61 (2019), Nazni et al., “Establishment of Wolbachia Strain wAlbB in Malaysian Populations of Aedes aegypti for Dengue Control,” Current Biology 29(24): 4241-4248 (2019), and Indriani et al., “Reduced Dengue Incidence Following Deployments of Wolbachia-Infected Aedes aegypti in Yogyakarta, Indonesia,” Gates Open Res. 4:50 (2020), each of which is hereby incorporated by reference in its entirety).Specifically, CI results in embryonic death when symbiotic males mate with aposymbiotic females. Nullification of death, and thus rescue of CI, occurs when transmitting females and their eggs harbor the same strain of Wolbachia (Shropshire et al., “Initiative, Symbiont- mediated Cytoplasmic Incompatibility : What Have We Learned in 50 Years?,” Elife 1-51 (2020), which is hereby incorporated by reference in its entirety).

[0282] CI is caused by male germline expression of two cytoplasmic incompatibility factor genes, cifA and cifB, encoded by prophage WO in the u Mel strain of Wolbachia fromDrosophila melanogaster (LePage et al., “Prophage WO Genes Recapitulate and Enhance Wolbachia-mduced Cytoplasmic Incompatibility,” Nature 543:243-247 (2017) and Beckmann et al., “A Wolbachia Deubiquitylating Enzy me Induces Cytoplasmic Incompatibility,” Nat. Microbiol. 2: 17007 (2017), each of which is hereby incorporated by reference in its entirety); rescue occurs by expression of cifA alone in ovaries (Shropshire et al., “One Prophage WO Gene Rescues Cytoplasmic Incompatibility in Drosophila melanogaster,” Proc. Natl. Acad. Sci. USA 115:4987-4991 (2018), which is hereby incorporated by reference in its entirety). We recently established that the wMel CifA and CifB proteins invade developing sperm nuclei and alter the abundance of histone and protamine nucleoproteins essential for fertility in Drosophila melanogaster (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). However, the action of the Cif proteins raise the question of how they incipiently modulate developing sperm macromolecules to impair nucleoprotein composition. Here, we show that the Cif proteins modify long non-coding RNA and DNA content of developing sperm early in spermatogenesis to trigger a developmental cascade that results in sperm chromatin integrity changes and induction of CI. Our findings link arthropod reproductive biology and prophage-facilitated bacterial modulation of sperm RNA and DNA macromolecules.Example 16 - CifA and CifB Are In Vitro Nucleases

[0283] To resolve CifA’s molecular function, we used a machine learning tool called CLEAN (Yu et al., “Enzyme Function Prediction Using Contrastive Learning,” Science 379(6639): 1358-1363 (2023), which is hereby incorporated by reference in its entirety). CifA was identified as a site-specific type II restriction endonuclease (p<0.001) whose family of enzymes cleave DNA and strands of RNA-DNA heteroduplexes (Murray et al., “Sequencespecific Cleavage of RNA by Type II Restriction Enzymes,” Nucleic Acids Res 38:8257-8268 (2010), which is hereby incorporated by reference in its entirety). CifB variants are DNases with active PDDEXK (SEQ ID NO:20) nuclease domains (Chen et al., “A Wolbachia Nuclease and its Binding Partner Provide a Distinct Mechanism for Cytoplasmic Incompatibility,” Proc NatlAcadSci USA 116:22314-22321 (2019) and Sun et al., “The CinB Nuclease from w No Wolbachia Is Sufficient for Induction of Cytoplasmic Incompatibility in Drosophila ” mBio 13 (2022), each of which is hereby incorporated by reference in its entirety), and sequence homologs across CifB evolution broadly contain thesedomains (LePage et al., “Prophage WO Genes Recapitulate and Enhance Wo / bachia-mduced Cytoplasmic Incompatibility,” Nature 543:243-247 (2017), Beckmann et al., “A Wolbachia Deubiquitylating Enzyme Induces Cytoplasmic Incompatibility,” Nat Microbiol 2: 17007 (2017), and Lindsey et al., “Evolutionary Genetics of Cytoplasmic Incompatibility Genes cifA and cifB in Prophage WO of Wolbachia,'” Genome Biol Evol 10:434-451 (2018), each of which is hereby incorporated by reference in its entirety). Thus, we investigated prophage proteins function with in vitro enzymatic assays to systematically determine the ribonuclease (RNase) and deoxyribonuclese (DNase) activity of biochemically uncharacterized Cif proteins from wMel. We generated CifA and CifB proteins by recombinant expression (FIG. 14A) and incubated them with single-strand (ss)DNA, double-strand (ds)DNA, and ssRNA oligonucleotide substrates. Purified CifA cleaved ssDNA, dsDNA, and ssRNA substrates (FIG. 14B). Nuclease activity was halted in the presence of the chelating agent ethylenediaminetetraacetic acid (EDTA). Mass spectrometry of the purified proteins revealed small amounts of E. coli ribonuclease peptide in purifications of CifA (7 / 1470 total spectral count) and CifB (2 / 3150) (Data SI file of Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal-effect Lethality." Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety). However, 8-fold diluted CifA was able to degrade RNA in vitro (FIG. 19A), whereas CifB at the same dilution lacked any RNase activity (see below), supporting a causal link between the CLEAN predicted annotation for CifA and its in vitro RNase activity.

[0284] CifB was synthesized without its deubiquitinase (DUB) domain (hereafter called CUBAD) (FIG. 14A and FIG. 19A-B), because full length CifB is too large to express in recombinant E. coli (Wang et al., “Crystal Structures of Wolbachia CidA and CidB Reveal Determinants of Bacteria-induced Cytoplasmic Incompatibility and Rescue,” Nat Commun 13(1): 1608 (2022), which is hereby incorporated by reference in its entirety). CifB AD cleaved ssDNA and dsDNA substrates similarly to the control DNase enzyme but lacked RNase activity against a ssRNA oligonucleotide substrate (FIG. 14B). EDTA inhibited the DNase function as expected. Enzymatic activity decreased with lower enzyme concentrations and shorter incubation times (FIGs. 19A-B). Mass spectrometry showed no co-purified E. coli DNases were present that might confound the result (data SI file of Kaur et al., “Prophage Proteins Alter Long Noncoding RNA and DNA of Developing Sperm to Induce a Paternal- effect Lethality,” Science 383: 1111-1117 (2024), which is hereby incorporated by reference in its entirety). CifB possesses a QxxxY (SEQ ID NO: 19) motif within a region of predicteda -helices at the C-terminal nuclease domain (FIGs. 20A-B). This motif is characteristic of RecB-family nucleases with HsdR subunits of E. colt that contain a PDDEXK domain (SEQ ID NO:20) (Singleton et al., “Crystal Structure of RecBCD Enzyme Reveals a Machine for Processing DNA Breaks,” Nature 432(7014): 187-93 (2004), which is hereby incorporated by reference in its entirety). Mutating Q and Y residues impaired EcoR124I DNA cleavage, possibly by destabilizing the catalytic pocket or modulating the binding efficacy of this domain to the DNA (Sisakova et al., “A RecB-family Nuclease Motif in the Type I Restriction Endonuclease EcoR124I,” Nucleic Acids Res 36(12):3939-49 (2008), which is hereby incorporated by reference in its entirety). The presence of the QxxxY motif in CifB associates with its DNase activity (FIGs. 20A-B) as the mutant protein with alanine substitutions at the Q and Y residues (CUBQY) failed to cleave ss- and dsDNA (FIGs. 14A-B). CifA also possesses a QxxxY motif within a predicted a -helix (FIGs. 20A-B) however, the mutant protein (CifApy) maintained its dsDNA and ssRNA cleaving activity, although it lost its ssDNAase function (FIG. 14B), which indicates that the QxxxY motif may not be functionally essential to the CifA nuclease activity in vitro. These results show that CifB from wMel Wolbachia is a DNase, contrary to previous reports (Beckmann et al., “A Wolbachia Deubiquitylating Enzyme Induces Cytoplasmic Incompatibility,” Nat Microbiol 2: 17007 (2017), which is hereby incorporated by reference in its entirety) and that the QxxxY motif is necessary for its DNase function.Example 17 - Impact of Evolutionary-conserved Mutations on Cif s In Vitro Nuclease Properties

[0285] It was previously shown that mutations in highly conserved residues across CifA and CifB impact transgenic CI (Weinert et al., “The Incidence of Bacterial Endosymbionts in Terrestrial Arthropods,” Proceedings of the Royal Society B: Biological Sciences 282(1807):20150249 (2015), which is hereby incorporated by reference in its entirety), suggesting their essentiality to the CI mechanism. To investigate the link between CI and nuclease activity, nuclease activity was first tested in vitro and found that mutant proteins failed to reveal a pattern associated with CI. Non-CI inducing CifA? mutant with mutations in the Puf-farmly RNA-binding domain nicked both DNA and RNA substrates (FIGs. 28A-B) and Ci-inducing CifA4 with mutations in the STE transcription factor domain lacked ss-DNase activity but cleaved dsDNA and ssRNA (FIGs. 28A-B). Mutating residues across full CifB protein including both nuclease domains ablated CI and DNase activity in general. ssDNA cleavage was abolished by CifB substitutions in the NTND (CifB2;Ao) and CTND (CifB3;AD),whereas ds-DNase activity was ablated by Ciffi2;AD only (FIGs. 28A-B). Thus, the CifEh sites were crucial for both ssDNA and dsDNA cleavage, whereas CifBs sites were substrate specific. These data were semi-consistent with crystal structures of Cif proteins (Sisakova et al., “A RecB-family Nuclease Motif in the Type I Restriction Endonuclease EcoR124I,” Nucleic Acids Res. 36(12): 3939-49 (2008), which is hereby incorporated by reference in its entirety) that showed the NTND of T4 CifB from wPip strain was the main catalytic center, and the CTND may be involved in specific substrate binding. Truncated CifB were also generated with nuclease domains only (CifBAAAD, 277-796 aa) and the A-terminus alone (CUBANACAD, 1-276 aa) implicated in chromatin binding (Cenci et al., “Chromatin and Microtubule Organization During Premeiotic, Meiotic and Early Postmeiotic Stages of Drosophila melanogaster Spermatogenesis,” J. Cell Sci. 107:3521-3534 (1994), which is hereby incorporated by reference in its entirety) (FIG. 28A and FIG. 29). CUBAAAD did not cleave ssDNA and dsDNA (FIGs. 28A-B), suggesting the A-terminus plays a crucial role in nuclease activity, for instance, by affecting proper structural folding of the protein or binding to the target substrate. CU ANACAD also did not exhibit DNA cleavage, as expected (FIG. 29).Example 18 - Impact of Evolutionary-conserved Mutations on Cif s In Situ DNase Properties

[0286] To evaluate whether CI is a result of altered in situ DNase properties by Cif mutants, testes were stained with TUNEL in transgenic CifA and CifB mutants when expressed alone or together. Single and dual expression of CifB mutants with CifA generally ablated both spermatid DNase activity (FIGs. 24B and 24E) and CI (FIGs. 24C and 24F), suggesting that intact CifB was necessary for CI and spermatid DNA nicking. Most CifA mutants ablated CI and showed very few or no sperm bundles with enhanced DNA damage upon individual (FIGs. 24A) or dual expression with intact CifB (FIGs. 24D), confirming CifB’s DNase dependence on intact CifA, possibly by hampering CifB binding to the target substrate or to CifA. The CifAs mutant residues are near the binding interface of CifA and CifB (Sisakova et al., “A RecB-family Nuclease Motif in the Type I Restriction Endonuclease EcoR124I,” Nucleic Acids Res 36(12):3939-49 (2008), which is hereby incorporated by reference in its entirety) and notably failed to cause CI but rescue it (Weinert et al., “The Incidence of Bactenal Endosymbionts in Terrestrial Arthropods,” Proceedings of the Royal Society B: Biological Sciences 282(1807):20150249 (2015), which is hereby incorporated by reference in its entirety). Thus, binding of CifA and CifB may be important to CI but not rescue. Moreover, mutagenesis may affect the structural stability of the proteins (especially invitro vs. in situ) to render them incapable of binding / cleaving their nucleotide substrates in developing sperm. The mutant CifA4B did not cleave spermatid DNA in situ (FIG. 24D), yet it caused strong transgenic CI (FIG. 24F), suggesting that these substituted residues are mutually exclusive for DNA nicking and CI properties and may possess DNA versus RNA substrate-specificity consistent with their in situ ability to access and deplete the spermatocyte IncRNA (FIG. 18A) and impair the histone-protamine exchange during sperm maturation (FIGs. 18C-D), which is tied to the mechanistic pathway(s) launching CI.Example 19 - CifA-mediated Depletion of Long Non-coding RNA in Spermatocytes Strengthens CI

[0287] Having validated RNase and DNase properties of CifA and CifB in vitro, respectively, using transgenic and cytochemical approaches we tested the effect of Cifs in vivo against RNA and DNA during sperm development. In Drosophila, several mitotic divisions produce primary spermatocytes (spermatogenesis) before they enter meiosis to form elongating spermatids (spermiogenesis). Genes in primary spermatocytes are actively transcribed for spermiogenesis (Cenci et al., “Chromatin and Microtubule Organization During Premeiotic, Meiotic and Early Postmeiotic Stages of Drosophila melanogaster Spermatogenesis,'’ J. Cell Sci. 107:3521-3534 (1994), which is hereby incorporated by reference in its entirety). Long non-coding RNAs (IncRNAs) predominantly transcribed in Drosophila testes play a crucial role in chromatin organization during sperm development and male fertility (Chu et al., “Genomic Maps of Long Noncoding RNA Occupancy Reveal Principles of RNA-Chromatin Interactions,” Mol Cell 44:667-678 (2011) and Wen et al., “Critical Roles of Long Noncoding RNAs in Drosophila spermatogenesis,” Genome Res. 26: 1233-1244 (2016), each of which is hereby incorporated by reference in its entirety).IncRNA derived from AAGAG tandem repeats in heterochromatic regions are highly expressed in primary spermatocytes, and depletion results in a defective histone-to-protamine (H-P) transition during sperm chromatin organization (Mills et al., “RNA From a Simpletandem Repeat is Required for Sperm Maturation and Male Fertility in Drosophila melanogaster." Elife 8:e48940 (2019), which is hereby incorporated by reference in its entirety). We previously showed that wMel and transgenic cif expression impairs the H-P transition to induce CI (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). Thus, wehypothesized CifA might degrade AAGAG IncRNA. among others, owing to CifA’s RNase activity, and alter the H-P transition to establish CI.

[0288] <8-hour old wild type (wMel+ and wMel-) and transgenic c / / -expressing fly testes were dissected and the abundance of the AAGAG IncRNA in primary spermatocytes was measured using fluorescent in situ hybridization. Ci-inducing wMel+ and CifAB- expressing males had 1.2 - 2.4-fold less AAGAG IncRNA relative to their aposymbiotic counterparts (FIGs. 15A-D and FIG. 21). Upon individual expression, CifA caused a 1.3-fold reduction in AAGAG IncRNA relative to CifB (FIGs. 15A-B). Thus, CifB does not interfere with CifA’s ability to access spermatocyte RNA and cleave it in CI males. Notably, CifA harbors a bipartite nuclear localization signal (bNLS) essential for nuclear targeting, CI, and the H-P exchange (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety)- Mutant CifA with a deleted bNLS (CifAAbNLs) had significantly higher AAGAG RNA compared to CifA and CifAB lines (FIGs. 15A-B), corroborating the functional importance of CifA nuclear import in accessing CI targets.

[0289] To directly test the involvement of the IncRNA in CI, we used transgenic flies with RNAi -mediated knockdown of the AAGAG IncRNA (Mills et al., “RNA From a Simpletandem Repeat is Required for Sperm Maturation and Male Fertility in Drosophila melanogaster ” Elife 8:e48940 (2019), which is hereby incorporated by reference in its entirety). If the IncRNA is causal to CI, then depleting the AAGAG IncRNA in the presence of wMel should increase wild type CI. CI was assessed with a standard embryonic viability assay by measuring the percentage of embryos that hatch into larvae. wMel-carrying AAGAG knockdown males (AAGAG_KD+), induced approximately three-fold more CI (median embryonic hatching = 11.8%) upon matings with uninfected (wMel-) females compared with control males (Scramble_KD+) with a randomized A and G content that induced intermediate levels of CI (median embryonic hatching = 35.6%, FIG. 15C). These findings indicate causal interaction between Wolbachia and AAGAG IncRNA depletion on CI. Control AAGAG knockdown males without Wolbachia (AAGAG_KD-) did not recapitulate CI on their own (FIG. 22), indicating that AAGAG IncRNA depletion alone is not the sole cause of CI and must operate in conjunction with other CI modifications by the Cifs. Both CifA and CifB localize to spermatogonia and spermatocyte nuclei (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish CytoplasmicIncompatibility,” PLoS Biol. 20(5) :e3001584 (2022), which is hereby incorporated by reference in its entirety) thus, it is possible that Cifs act on AAGAG IncRNA and perhaps other spermatogenesis-related IncRNAs (Wen et al., “Cntical Roles of Long Noncoding RNAs in Drosophila spermatogenesis,” Genome Res. 26: 1233-1244 (2016) and Vedelek et al., “Analysis of Drosophila melanogaster Testis Transcriptome,” BMC Genomics 19(1):69 (2018), each of which is hereby incorporated by reference in its entirety) earlier during pre- meiotic spermatogenesis to prime cells and / or chromosomes to affect downstream, post- meiotic sperm chromatin organization. These findings are consistent with single-cell transcriptome sequencing in which Wolbachia alter fly gene expression in early stages of spermatogenesis, leading to downstream sperm defects (Dou et al., “Single-cell Transcriptome Sequencing Reveals Wolbachia-medmted Modification in Early Stages of Drosophila spermatogenesis,” Proceedings of the Royal Society B: Biological Sciences 290(1990): 20221963 (2023), which is hereby incorporated by reference in its entirety). Altogether, the role of CifA in depleting AAGAG IncRNA suggests CifA RNase activity is central to establishing the CI modification during early sperm development.Example 20 - CifA and CifB Enhance DNA Damage in Developing CI Spermatids

[0290] DNA damage and repair are typical features facilitating the H-P exchange during the canoe stage of spermiogenesis to tightly pack the sperm chromatin (Rathke et al., “Transition from a Nucleosome-based to a Protamine-based Chromatin Configuration During Spermiogenesis in Drosophila f J. Cell Sci. 120:1689-1700 (2007), which is hereby incorporated by reference in its entirety). However, in animals, excessive DNA breaks result in abnormal chromatin integrity , reduce male fertility, and cause embryonic inviability (Agarwal & Said, “Role of Sperm Chromatin Abnormalities and DNA Damage in Male Infertility,” Hum. Reprod. Update 9(4):331-45 (2003) and Hosen et al., “Oxidative Stress Induced Sperm DNA Damage, a Possible Reason for Male Infertility,” Iran J. Reprod. Med. 13(9): 525-32 (2015), each of which is hereby incorporated by reference in its entirety), suggesting a link to CI. Since Cifs alter the abundance of histone and protamine nucleoproteins (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol. 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety) and both Cifs catabolize DNA, we hypothesized that they promote spermatid DNA damage in elongating spermatids.

[0291] TUNEL staining was used to evaluate DNA fragmentation in wild type and transgenic D. melanogaster testes. TUNEL signals persisted at higher levels in Ci-inducingw’Mel- and cifAB late canoe spermatids, occasionally in needle spermatids than non-CI controls (FIGs. 16A-B and FIGs. 23A-B) with significantly higher number of damaged bundles (FIG. 16C), validating that Wolbachia and the Cif enzymes enhanced in situ DNA damage during elongation process. Single CifA and CifB also promoted spermatid DNA damage relative to the transgenic negative control (FIGs. 16A-C), confirming their in vitro enzymatic independence to cleave DNA. Notably, their individual impacts on DNA damage, as measured by corrected total cell fluorescence (CTCF) of the TUNEL signal (CifA median CTCF = 233. 12 and CifB median CTCF = 364.06) was approximately equivalent to the DNA damage.induced by dual expression of CifA and CifB (median CTCF = 506.42), indicating they additive effect (FIG. 16B). Moreover, CifA and CifB mutants that did not result in CI (Shropshire et al., “Evolution-guided Mutagenesis of the Cytoplasmic Incompatibility Proteins: Identifying CifA’s Complex Functional Repertoire and New Essential Regions in CifB,” PLoS Pathog 16(8):el008794 (2020), which is hereby incorporated by reference in its entirety) did not nick spermatid DNA in situ (FIGs. 24A-F) suggesting spermatid DNA damage contributes to CI phenotype. By co-staining TUNEL with Cif antibodies, we found that at the early canoe-stage, both CifA and CifB signals localized at the tip of spermatid heads with TUNEL-positive signals (FIG. 25). At the needle stage, damaged spermatids showed CifA localized toward the tail, and CifB in the sperm head remained undetectable due to poor antibody penetration in highly condensed nuclei, as shown previously (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety). Consequently, we propose that Cif presence in early spermatogonia, spermatocytes, onion spermatid nuclei, and early canoe-stage (Kaur et al., “The Cif Proteins from Wolbachia Prophage WO Modify Sperm Genome Integrity to Establish Cytoplasmic Incompatibility,” PLoS Biol 20(5):e3001584 (2022), which is hereby incorporated by reference in its entirety) prime the spermatid DNA for damage detected in late elongating stages of spermiogenesis. Our results are also consistent with a previous study on elevated oxidative DNA damage in CI spermatocytes (Brennan et al., “Disruption of Redox Homeostasis Leads to Oxidative DNA Damage in Spermatocytes of Wolbachia-infQcted Drosophila simulans ” Insect Mol Biol doi: 21 (5): 510-20 (2012), which is hereby incorporated by reference in its entirety). In sum, CI sperm suffers increased DNA damage beyond levels essential for a normal H-P transition and chromatin architecture.Example 21 - CI embryos Succumb to DNA Damage

[0292] Because CifAB-expressing CI males suffer from depleted IncRNA and enhanced DNA damage resulting in sperm development with altered H-P abundance, it was next investigated whether fertilization by the compromised CI sperm causes adverse effects in embryos during nuclear divisions. Previous studies showed that Drosophila embryonic DNA damage is only evident in late stages of development ~6 hours after egg deposition (AED) towards gastrulation (Arama & Steller, “Detection of Apoptosis by Terminal Deoxynucleotidyl Transferase-mediated dUTP Nick-end Labeling and Acridine Orange in Drosophila Embryos and Adult Male Gonads,” Nat. Protoc. 1 : 1725-1731 (2006), Abrams et al., “Programmed Cell Death During Drosophila Embryogenesis,” Development 117:29-43 (1993), and Harumoto et al., “Common and Unique Strategies of Male Killing Evolved in Two Distinct Drosophila Symbionts,” Proceedings of the Royal Society B: Biological Sciences 285(1875):20172167 (2018), each of which is hereby incorporated by reference in its entirety) as the cells at this stage become hyperprolifertive, hence more susceptible to damage (Bartek, J., “DNA Damage Response, Genetic Instability and Cancer: From Mechanistic Insights to Personalized Treatment,” Mol. Oncol. 5:303-307 (2011), Baonza et al., “Regulation and Coordination of the Different DNA Damage Responses in Drosophila,” Front. Cell. Dev. Biol. 10:993257 (2022), and Zhang et al., “Erratum: An Intergenic Regulatory Region Mediates Drosophila Myc-induced Apoptosis and Blocks Tissue Hyperplasia,” Oncogene 34:2412-2412 (2015), each of which is hereby incorporated by reference in its entirety). While CI is often associated with a first mitotic division defect within 1 hour of egg fertilization, additional developmental defects also occur in late CI embryos developing through the pre-blastoderm divisions, syncytial, and cellularized blastoderms (LePage et al., “Prophage WO Genes Recapitulate and Enhance Wolbachia- induced Cytoplasmic Incompatibility,” Na ture 543:243-247 (2017), Lassy & Karr, “Cytological Analysis of Fertilization and Early Embryonic Development in Incompatible Crosses of Drosophila simulansf Meeh. Dev. 57:47-58 (1996), Callaini et al., “Mitotic Defects Associated with Cytoplasmic Incompatibility in Drosophila simulansf J. Invertebr. Pathol. 67:55-64 (1996) and Warecki et al., “Wolbachia Action in the Sperm Produces Developmentally Deferred Chromosome Segregation Defects During the Drosophila Midblastula Transition,” Elife 11 :e81292 (2022), each of which is hereby incorporated by reference in its entirety).- Il l -

[0293] Embryos aged 0-1 hour, 1-2 hours, and 2-3 hours AED representing CI, non- CI, and rescue crosses were collected. The pH2Av marker, a variant of histone H2Av that is phospohorylated in response to DNA damage was used (Harumoto et al., “Male-killing Symbiont Damages Host’s Dosage-compensated Sex Chromosome to Induce Embryonic Apoptosis,” Nat. Commun. 7: 12781 (2016), and Perlmutter et al., “The Phage Gene wmk is a Candidate for Male Killing by a Bacterial Endosymbiont,” PLoS Pathog. 15(9): e 1007936 (2019), each of which is hereby incorporated by reference in its entirety). As expected with the literature, pH2Av signals indicative of DNA damage were not detected in early 0-1 hour old CI embryos (nos;cifAB x wMel-) arrested after first mitotic division as well as 0-1 hour and 1-2 hours old rescue embryos (nos;cifAB x wMel+) undergoing rounds of nuclear divisions (FIG. 26) (Abrams et al., “Programmed Cell Death During Drosophila Embryogenesis,” Development 117:29-43 (1993), which is hereby incorporated by reference in its entirety). However, DNA damage signals started to be detected in 2-3 hours AED CI embryos during cellular-blastoderm formation (nuclear cycles 12-14) (Kotadia et al., “Blastoderm Formation and Cellularisation in Drosophila melanogaster ” in ELS (Wiley, 2010), which is hereby incorporated by reference in its entirety), when dividing nuclei migrate to the cortex periphery of the embryo (Abrams et al., “Programmed Cell Death During Drosophila Embryogenesis,” Development 117:29-43 (1993) and Kotadia et al., “Blastoderm Formation and Cellularisation in Drosophila melanogaster ” in ELS (Wiley, 2010), each of which is hereby incorporated by reference in its entirety). Intense pH2Av signals were observed in dense chromatin nuclei with cellular shrinkage in 32% of tota...

Claims

CLAIMSWHAT IS CLAIMED IS:

1. A method of population suppression of arthropods, the method comprising: contacting a population of male arthropods with a histone acetyltransferase (HAT) inhibitor, wherein said contacting:(i) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females lacking a bacterial endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont or in comparison to uncontacted males; or(ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to uncontacted males; and releasing the population of male arthropods into a target population of arthropods lacking a rescuing bacterial endosymbiont under conditions effective to reduce the target population.

2. A method of population replacement of target arthropods, the method comprising: contacting a population of male arthropods with a histone acetyltransferase (HAT) inhibitor, wherein said contacting:(i) increases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a rescuing bacterial endosymbiont in comparison to arthropod females lacking a rescuing bacterial endosymbiont, or(ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to uncontacted males; and either introducing the male arthropods into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, wherein said introducing replaces the target population lacking the rescuing endosymbiont orintroducing a population of female arthropods having a rescuing bacterial endosymbiont and the contacted male arthropods into a target population of arthropods, wherein said introducing replaces the target population of arthropods.

3. The method of any one of the preceding claims, wherein the male arthropods lack a bacterial endosymbiont.

4. The method of any one of the preceding claims, wherein the male arthropods have a bacterial endosymbiont.

5. The method of any one of the preceding claims, wherein the HAT inhibitor is selected from the group consisting of: anacardic acid, garcinol, a garcinol analog, curcumin, y-butyrolactone, MB-3, isothiazolones, quinoline derivatives such as MCI 626, C646, 1-CBP112, MG149, NU9056, EP300 / CBP inhibitor-2, and TIP60 inhibitor-1.

6. The method of any one of the preceding claims, wherein said contacting is carried out at a larval stage of the male arthropod.

7. The method of any one of the preceding claims, wherein the contacted male arthropod comprises reduced histone H3 and / or H4 acetylation levels during sperm development in comparison to a non-contacted male arthropod.

8. The method of claim 7, wherein the histone H3 and / or H4 acetylation levels during sperm development in the contacted male arthropod are reduced by at least 20%.

9. The method of claim 1, wherein histone H2B acetylation levels during sperm development are not reduced.

10. The method of any one of the preceding claims, wherein viable offspring are reduced by at least 20%.

11. The method of any one of the preceding claims, wherein the arthropod is an insect.

12. The method of any one of the preceding claims, wherein the insect is selected from the group consisting of a mosquito, planthopper, rootworm, beetle, cricket, leafhopper, aphid, fly, ant, wasp, cockroach, termite, looper, caterpillar, and moth.

13. The method of any one of the preceding claims, wherein the bacterial endosymbiont is Wolbachia.

14. An arthropod control formulation comprising: a histone acetyltransferase (HAT) inhibitor and a carrier comprising the HAT inhibitor, wherein the carrier is suitable for delivering the HAT inhibitor to a male arthropod to cause a reduction in an ability of the male arthropod to produce viable offspring with a female arthropod (i) lacking a bacterial endosymbiont or (ii) having a non-rescuing endosymbiont compared to a female arthropod having a rescuing bacterial endosymbiont.

15. The arthropod control formulation of claim 14, wherein the carrier is a food or a spray.

16. The arthropod control formulation of claim 14 or claim 15, wherein the HAT inhibitor is anacardic acid.

17. A genetically engineered arthropod comprising: a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase gene; or a promoter operably linked to a histone deacetylase, wherein expression of the inhibitory RNA molecule targeting a histone acetyltransferase gene or overexpression of the histone deacetylase during sperm development in a male arthropod causes a reduction in fertility and / or the ability7of the male arthropod to produce viable offspring with arthropod females lacking a bacterial endosymbiont or having a non-rescuing endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont.

18. The genetically engineered arthropod of claim 17, wherein the male arthropod comprises reduced histone H3 and / or H4 acetylation levels in sperm in comparison to a wild-type male arthropod.

19. The genetically engineered arthropod of claim 17 or claim 18, wherein viable offspring are reduced by at least 20%.

20. The genetically engineered arthropod of any one of claims 17-19, wherein the arthropod is an insect.

21. The genetically engineered arthropod of any one of claims 17-20, wherein the insect is selected from the group consisting of a mosquito, planthopper, rootworm, beetle, cricket, leafhopper, aphid, fly, ant, wasp, cockroach, termite, looper, caterpillar, and moth.

22. The genetically engineered arthropod of any one of claims 17-21, wherein the arthropod male is not infected with a bacterial endosymbiont.

23. The genetically engineered arthropod of any one of claims 17-22, wherein the arthropod male is infected with a bacterial endosymbiont.

24. The genetically engineered arthropod of any one of claims 17-23, wherein the bacterial endosymbiont is Wolbcichia.

25. The genetically engineered arthropod of any one of claims 17-24, wherein the histone acetyl transferase gene is HAT1.

26. The genetically engineered arthropod of claim 22, wherein the HAT1 gene encodes any one of SEQ ID NOs:4, 8, or 10.

27. The genetically engineered arthropod of any one of claims 17-24, wherein the histone deacetylase gene is HDAC1.

28. The genetically engineered arthropod of claim 27, wherein the HDAC1 gene encodes SEQ ID NO: 6.

29. A method of population suppression of arthropods, the method comprising: providing a genetically engineered arthropod, said arthropod comprising: a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase, wherein expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod:(i) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females lacking a bacterial endosymbiont in comparison to arthropod females having a rescuing bacterial endosymbiont or wild-type males; or(ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to wild-type males; and releasing the genetically engineered arthropod into a target population of arthropods under conditions effective to reduce the target population.

30. A method of population replacement of target arthropods, the method comprising: providing a genetically engineered arthropod, said arthropod comprising: a promoter operably linked to an inhibitory RNA molecule targeting a histone acetyltransferase; or a promoter operably linked to a histone deacetylase, wherein expression of the inhibitory RNA molecule or overexpression of the histone deacetylase during sperm production in a male arthropod:(i) increases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a rescuing bacterial endosymbiont in comparison to arthropod females lacking a rescuing bacterial endosymbiont, or(ii) decreases the ability of the male arthropods to produce viable offspring upon mating with arthropod females having a non-rescuing bacterial endosymbiont in comparison to wild-type males; and either introducing the male arthropods into a target population of arthropods in which a portion of the target population has a rescuing bacterial endosymbiont, wherein said introducing replaces the target population lacking the rescuing endosymbiont; or introducing a population of female arthropods having a rescuing bacterial endosymbiont and the male arthropods into a target population of arthropods, wherein said introducing replaces the target population of arthropods.

31. The method of claim 29 or claim 30, wherein the male arthropods lack a bacterial endosymbiont.

32. The method of any one of claims 29-31, wherein the male arthropods have a bacterial endosymbiont.

33. The method of any one of claims 29-32, wherein the male arthropod comprises reduced histone H3 and / or H4 acetylation levels in sperm in comparison to a wildtype male arthropod.

34. The method of any one of claims 29-33, wherein viable offspring are reduced by at least 20%.

35. The method of any one of claims 29-34, wherein the arthropod is an insect.

36. The method of any one of the claims 29-35, wherein the insect is selected from the group consisting of a mosquito, planthopper, rootworm, beetle, cricket, leafhopper, aphid, fly, ant, wasp, cockroach, termite, looper, caterpillar, and moth.

37. The method of any one of claims 29-36, wherein the bacterial endosymbiont is Wolbachia.

38. A chemically treated male arthropod, wherein the male arthropod has at least 10% less viable offspring when mated with a female arthropod having no endosymbiont or a non-rescuing bacterial endosymbiont in comparison to mating with a female arthropod having a rescuing bacterial endosymbiont.

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