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CRISPR/Cas9-mediated gene deletion of the ompA gene in symbiotic Cedecea neteri impairs biofilm formation and reduces gut colonization of Aedes aegypti mosquitoes

  • Shivanand Hegde,

    Roles Conceptualization, Formal analysis, Investigation, Methodology, Validation, Writing – original draft

    Affiliation Departments of Vector Biology and Tropical Disease Biology, Liverpool School of Tropical Medicine, Liverpool, United Kingdom

  • Pornjarim Nilyanimit,

    Roles Investigation, Methodology

    Affiliation Center of Excellence in Clinical Virology, Chulalongkorn University, Bangkok, Thailand

  • Elena Kozlova,

    Roles Investigation, Methodology

    Affiliation Department of Pathology, University of Texas Medical Branch, Galveston, Texas, United States of America

  • Enyia R. Anderson,

    Roles Methodology

    Affiliation Departments of Vector Biology and Tropical Disease Biology, Liverpool School of Tropical Medicine, Liverpool, United Kingdom

  • Hema P. Narra,

    Roles Investigation, Methodology

    Affiliation Department of Pathology, University of Texas Medical Branch, Galveston, Texas, United States of America

  • Sanjeev K. Sahni,

    Roles Supervision

    Affiliation Department of Pathology, University of Texas Medical Branch, Galveston, Texas, United States of America

  • Eva Heinz,

    Roles Formal analysis, Writing – review & editing

    Affiliation Department of Vector Biology and Department of Clinical Sciences, Liverpool School of Tropical Medicine, Liverpool, United Kingdom

  • Grant L. Hughes

    Roles Conceptualization, Formal analysis, Funding acquisition, Investigation, Project administration, Supervision, Writing – review & editing

    Affiliation Departments of Vector Biology and Tropical Disease Biology, Liverpool School of Tropical Medicine, Liverpool, United Kingdom



Symbiotic bacteria are pervasive in mosquitoes and their presence can influence many host phenotypes that affect vectoral capacity. While it is evident that environmental and host genetic factors contribute in shaping the microbiome of mosquitoes, we have a poor understanding regarding how bacterial genetics affects colonization of the mosquito gut. The CRISPR/Cas9 gene editing system is a powerful tool to alter bacterial genomes facilitating investigations into host-microbe interactions but has yet to be applied to insect symbionts.

Methodology/Principal findings

To investigate the role of bacterial genetic factors in mosquito biology and in colonization of mosquitoes we used CRISPR/Cas9 gene editing system to mutate the outer membrane protein A (ompA) gene of a Cedecea neteri symbiont isolated from Aedes mosquitoes. The ompA mutant had an impaired ability to form biofilms and poorly infected Ae. aegypti when reared in a mono-association under gnotobiotic conditions. In adult mosquitoes, the mutant had a significantly reduced infection prevalence compared to the wild type or complement strains, while no differences in prevalence were seen in larvae, suggesting genetic factors are particularly important for adult gut colonization. We also used the CRISPR/Cas9 system to integrate genes (antibiotic resistance and fluorescent markers) into the symbionts genome and demonstrated that these genes were functional in vitro and in vivo.


Our results shed insights into the role of ompA gene in host-microbe interactions in Ae. aegypti and confirm that CRISPR/Cas9 gene editing can be employed for genetic manipulation of non-model gut microbes. The ability to use this technology for site-specific integration of genes into the symbiont will facilitate the development of paratransgenic control strategies to interfere with arboviral pathogens such Chikungunya, dengue, Zika and Yellow fever viruses transmitted by Aedes mosquitoes.

Author summary

Microbiota profoundly affect their host but few studies have investigated the role of bacterial genetics in host-microbe interactions in mosquitoes. Here we applied the CRISPR/Cas9 gene editing system to knockout a membrane protein in Cedecea neteri, which is a dominant member of the mosquito microbiome. The mutant strain had an impaired capacity to form biofilms, infected larvae and adults at lower titers, and had a reduced prevalence in adults. The lower prevalence in adults, but not larvae, likely reflects the difference in the modes of bacterial acquisition from the larval water of these two life stages. Importantly from an applied perspective, we also demonstrated that this editing technology can be harnessed for site-specific integration of genes into the bacterial chromosome. In proof-of-principle studies we integrated either a fluorescent protein or gene conferring antibiotic resistance into the bacterial genome and showed these transgenes were functional in mosquitoes. The specificity, flexibility, and simplicity of this editing approach in non-model bacteria will be useful for developing novel symbiotic control strategies to mitigate the burden of arthropod-borne disease.


Mosquitoes harbor a community of microbes within their guts. In general, the gut-associated microbiome of mosquitoes tends to have low species richness but can differ greatly between individuals and habitats [18]. Importantly, these microbes can modulate many host phenotypes, several of which can influence vectorial capacity [911]. As such, it is imperative that we understand how the microbiome is acquired and maintained within mosquito vectors. While environmental factors unquestionably influence the mosquito microbiome composition and abundance [24, 8], studies are elucidating the role of microbial interactions [5, 7, 12, 13] and host genetic factors [1418] in shaping the microbiome. However, we have a poor understanding of bacterial factors that influence colonization of the mosquito gut and this is likely an underappreciated force influencing host-microbe interactions in mosquitoes.

In other invertebrates, several bacterial genes have been implicated in gut colonization. For example, a genome wide screen exploiting transposon-sequencing found a suite of genes from the bacterium Snodgrasselia alvi involved in colonization of the honey bee gut [19]. These bacterial genes were classified into the broad categories of extracellular interactions, metabolism, and stress response [19]. Knockout of a purine biosynthesis gene in Burkholderia impaired biofilm formation and reduced bacterial colonization rates in a bean bug [20]. Biofilm formation was also shown to play a role in virulence of pathogenic Pseudomonas in artificial infections of Drosophila, with strains that lacked the capacity to form biofilms being more virulence to the host, although a hyperbiofilm strain was less virulent than the wild type (WT) strain [21]. In other blood feeding invertebrates, bacterial genetics also appears critical for host colonization. Knockout of the type II secretion system in Aeromonas veronii reduced infection in Hirudo verbena leeches [22]. In tsetse flies, the outer-membrane protein A (ompA) gene of Sodalis glossinidius is essential for symbiotic interactions [23]. Sodalis mutants lacking the ompA gene poorly colonized the fly gut compared to the WT symbionts [23], likely due to the mutant strains reduced capacity to form biofilms [24]. Heterologous expression of the ompA gene from pathogenic Escherichia coli in Sodalis mutants induced mortality in the fly implicating this gene as a virulence factor in pathogenic bacteria [23]. Taken together, these studies suggest that bacterial genetic factors are critical for host colonization of invertebrates and that biofilm formation facilitates symbiotic associations in insects.

In mosquitoes, few studies have investigated how bacterial genetics affect gut colonization. However, evidence from experimental evolution studies suggests bacterial genetics plays a critical role. In two separate studies, Enterobacter was selected for increased persistence in the gut of Anopheles gambiae mosquitoes, the major malaria vector in sub-Saharan Africa, by repeatedly infecting mosquitoes with strains that persisted in the gut for longer periods of time [25, 26]. Transcriptomics comparisons of effective and ineffective colonizers in liquid media identified 41 genes that were differentially expressed between these two strains [26], further implicating the importance of bacterial genetics in mosquito infection, however the role of these genes in colonization of the mosquito gut has not been resolved. In a separate study, in vitro screening of a transposon mutant library of Enterobacter identified a waaL gene mutant that was insensitive to oxidative stress [27]. The waaL gene encodes an O antigen ligase which is needed for attachment of the O antigen to lipopolysaccharide. The mutant was found to have lower rates of colonization of the midguts of Anopheles mosquitoes [27].

Gene knockouts approaches in bacteria provide compelling evidence of the role of bacterial genes in host-microbe interactions [2224, 2729]. In general, most studies use transposon mutagenesis for gene knockout, which requires screening of the mutant library. A targeted gene knockout approach is highly desirable to investigate the functionality of bacterial genes in host-microbe interactions. In the past few years, the CRISPR/Cas9 gene editing system has been employed to modify bacterial genomes [3032]. While much of the work has been done in model bacterial species [3137], editing approaches have expanded into non-model bacterial systems [3843]. Despite this expansion, the approach has been used less frequently for host-associated microbes [39, 44], and rarely for arthropod symbionts. In the vector biology field, gene knockout approaches can be used to interrogate the role of bacterial genes responsible for host-microbe interactions, whilst the ability to integrate genes into the bacterial symbiont genome has great potential for applied paratransgenic control strategies [10, 4547]. To date, manipulation of non-model symbionts that associate with insect vectors has been accomplished by plasmid transformation [4855] or stable transformation of the genome using transposons or integrative plasmids [5663], but the use of CRISPR/Cas9 gene editing in insect gut symbionts has yet to be accomplished. For paratransgenic strategies, stable site-specific integration of transgenes into the symbiont genome is critical. Therefore, the application of CRISPR/Cas9 gene editing technology to non-model bacteria that associate with insect vectors will stimulate research in this field.

We therefore undertook studies to develop CRISPR/Cas9 genome editing approaches in Cedecea neteri isolated from Aedes mosquitoes. We used the Scarless Cas9 Assisted Recombineering (no-SCAR) method to disrupt the ompA gene of the non-model C. neteri [35]. After characterization of the mutant in vitro, we examined the role of the ompA gene in host-microbe interactions by re-infecting bacteria into mosquitoes in a mono-association. To demonstrate that the CRISPR/Cas9 gene-editing system could be useful for applied symbiotic control approaches we inserted genes conferring antibiotic resistance or a fluorescent protein into the bacterial genome and re-infected the altered strains back into mosquitoes. Our result sheds insights into the role of the ompA gene in host-microbe interactions in Ae. aegypti and confirm that CRISPR/Cas9 gene editing can be a powerful tool for genetic manipulation of native gut-associated microbes of mosquitoes.


C. neteri biofilm formation in Ae. aegypti guts

Over the course of conducting mono-axenic infections in Ae. aegypti mosquitoes with a Cedecea symbiont, we repeatedly observed a conglomeration of bacterial cells in the anterior and posterior midgut (Fig 1, S1 Fig) that had a similar appearance to biofilms observed in the guts of other insects [21, 24]. We also infected mosquitoes with the E. coli BL21(DE3) lab strain as a control, but we did not see any evidence of infection (Fig 1, S1D–S1F Fig) although infection with this bacterium enabled mosquito development [64]. The E. coli BL21(DE3) lab strain does not have the capacity to form biofilms [65], possibly explaining its inability to infect mosquitoes. We therefore set out to examine the role of bacterial genetics in biofilm formation and host colonization of gut-associated bacteria of Aedes mosquitoes. We used multilocus sequence typing (MLST) to confirm the species of our isolate, which indicated the bacterium was C. neteri (S2 Fig). Several genes have been implicated in biofilm formation [21, 24], but we chose to knockout the ompA gene of C. neteri given that this gene has been demonstrated to influence biofilm formation and gut colonization of Sodalis [23, 24], an Enterobacteriaceae symbiont of tsetse flies. We used the CRISRP/Cas9 genome editing system to mutate the symbiont genome to demonstrate this approach could be employed for non-model symbiotic bacteria that associate with mosquitoes.

Fig 1. Midgut infection of C. neteri and E. coli in mono-associations of Aedes mosquitoes.

C. neteri forms a biofilm in the gut of 3–4 day old Ae. aegypti adult mosquitoes (left) while no bacteria were observed in the gut of mosquitoes reared with E. coli under gnotobiotic conditions (right). Bacteria possessed the pRAM-mCherry plasmid, which expresses the mCherry fluorescent protein and conferred resistance to kanamycin. Blue–host nuclei stained by DAPI. Green–host actin cytoskeleton stained with phalloidin. The scale bar is 70 μm.

Genome editing in C. neteri bacteria isolated from mosquitoes

To edit the Cedecea isolate that resides within the gut of Aedes mosquitoes, we employed the no-SCAR gene editing approach that had been developed in E. coli [35]. To optimize the approach in our hands, we performed initial experiments in E. coli to delete a ~1 kb region of the ompA gene (Fig 2A). As the no-SCAR approach exploits the λ-Red recombineering system to repair double stranded breaks, we transformed bacteria with a double stranded DNA template that had regions of homology flanking the gRNA site (250 bp for each arm). Using this approach, we successfully deleted a 1001 bp fragment of the ompA gene. From the colonies we screened, we saw an editing at a frequency of 6.25% (N = 48) (Fig 2A). For C. neteri, we altered our editing procedure to delete a 598 bp fragment from the ompA gene. This was done to enhance the efficiency of obtaining mutants [35] and accommodate the PAM site which was at a different location in the ompA gene in C. neteri. Using a donor template designed for the C. neteri ompA gene that had flanking homology arms of similar length as the previous experiment done in E. coli, we obtained mutant knockouts at a rate of 32% (N = 50) (Fig 2B). For both bacterial species, Sanger sequencing across the integration site indicated the deletion occurred at the expected loci in the bacterial genome (Fig 2C; S1 Appendix).

Fig 2. CRISPR/Cas9 genome editing in bacteria.

A schematic of the editing approach and screening of putative mutants in (A) E. coli and (B) C. neteri. A ~1kb fragment of E. coli BL21(DE3) was deleted using no-SCAR protocol. The 250 bp of the left arm (LA) and right arm (RA) was assembled to generate the 500 bp donor DNA. The transformants were screened via colony PCR with primers binding in regions flanking the deletion. Similar to the strategy employed in E. coli, the knockout of the ompA gene from C. neteri isolated from the mosquito gut was created by deleting the 598 bp fragment. The grey area indicates the PAM site in the ompA gene and arrow shows the cleavage site in the genome. (C) The sequence of the ompA mutation in E. coli and C. neteri was confirmed by Sanger sequencing. The sequence above the gene within the dotted line has been deleted. The chromatogram shows the 10 bp flanking the deletion.

Characterization of the C. neteri ompA mutant

We quantified the growth rates of the ΔompA mutant in comparison to the WT C. neteri and the ΔompA/ompA complement in liquid LB media. We saw minimal differences in the growth between the WT, the ΔompA mutant or the ΔompA/ompA complement (Fig 3A). To examine the stability of the deletion, we subcultured the ΔompA mutant on LB media for 10 generations and performed PCR to amplify across the deletion. At alternative generations, PCR analysis indicated the deletion was present indicating genomic stability at this site (Fig 3B).

Fig 3. In vitro characterization of the ompA mutation.

(A) The C. neteri ΔompA mutant had a similar growth rate compared to both the WT and the ΔompA/ompA complement in liquid LB media. Five technical replicates were used to create growth curves. (B) The stability of mutant was evaluated in vitro by continuous subculturing in LB media. Genomic DNA from alternative subcultures was used as template for PCR using primers that amplified across the deletion. The stability assay was repeated twice. Two separate gel images were merged to create figure 3B (passage 8 was run on a separate gel to passages 0–6). (C) Biofilm formation was assessed using the CV biofilm assay for the WT, ΔompA mutant and the ΔompA/ompA complement. Two biological replicates were completed. (D) Quantification of the relative biofilm formation normalized by the number of bacteria per well (N = 3). Error bars represent standard error. The assay was repeated twice.

Previously, ompA has been shown to be important in biofilm formation as Sodalis deletion mutants were unable to form biofilms [24]. Therefore, we characterized in vitro biofilm formation using the crystal violet (CV) biofilm assay. From visual inspection, it was clear the ΔompA mutant had distinctly less biofilm deposition compared to either the WT or the ΔompA/ompA complement (Fig 3C). After quantification and normalization to account for any difference in growth between the strains, biofilm formation was confirmed to be significantly different between the ΔompA mutant and the WT or complement (Fig 3D; Tukey’s multiple comparisons test, P < 0.0001). There was no significant difference between the WT and the ΔompA/ompA complement (Tukey’s multiple comparisons test P = 0.2).

The role of ompA gene in mosquito infection

To examine the importance of the ompA gene on bacterial colonization of mosquitoes, we infected Ae. aegypti mosquitoes in a mono-association under gnotobiotic conditions [64]. This infection method was used to avoid other gut-associated microbes influencing host colonization rates [7] and it also enabled straightforward quantification of introduced bacteria by measuring colony forming units (CFUs). No significant changes were seen in the prevalence of infection (number of mosquitoes infected) in the larval stage (Fig 4A, Fisher’s exact test; WT compared to ΔompA P = 0.24 and ΔompA compared to ΔompA/ompA P = 0.24) with rates of infection consistently high (WT 100%, ΔompA 96%, and ΔompA/ompA 100%). In adults, the prevalence of infection was significantly different (Fig 4B, Fisher’s exact test; WT compared to ΔompA P < 0.0001 and ΔompA compared to ΔompA/ompA P < 0.0001), with only 45% of adults infected by the ΔompA mutant compared to 95% and 88% by the WT and ΔompA/ompA complement, respectively. In larvae, we saw a significant reduction in bacterial titer in the mutant compared to both the WT (Kruskal-Wallis test with Dunn’s test; P < 0.05) and the ΔompA/ompA complement (Kruskal-Wallis test with Dunn’s test; P < 0.05) (Fig 4C) with median values of 1.5x105, 2.3x104, and 1.5x105 for the WT, ΔompA, and ΔompA/ompA complement respectively. Similarly, in adults, there was a significant reduction in bacterial infection in the ΔompA mutant compared to either the WT or ΔompA/ompA complement (Kruskal-Wallis test with Dunn’s test; P < 0.001) (Fig 4D), with median value of 8.1x102, 0, and 7.5x102 for the WT, ΔompA, and ΔompA/ompA complement respectively. However, when considering only the infected mosquitoes for analysis, we saw no significant difference between the treatments (S3 Fig, Kruskal-Wallis test with Dunn’s test; P > 0.99). For both the larvae and adult density quantifications, the non-parametric test (Kruskal-Wallis test) was used due to non-normal distribution of data (Sharpiro-Wilks test; P<0.001). We also monitored the growth rates of mosquitoes administered with the WT, ΔompA mutant and ΔompA/ompA complement. No significant differences were seen in the time to pupation (Fig 5A) or percentage of first instar larvae that reached adulthood (Fig 5B) between any of the bacterial strains.

Fig 4. The ΔompA mutant poorly infected mosquitoes.

Infection of C. neteri strains (WT, ΔompA mutant and ΔompA/ompA complement; the former two possessed the pRAM-Cherry plasmid while the latter possessed the pRAM-Cherry-Ent-OmpA plasmid) reared in a mono-association using a gnotobiotic rearing approach for larvae (A and C) and adults (B and D). L4 larvae and 3–4 days post emergence adults were screened for bacterial load by plating on selective LB media with kanamycin to quantify the bacteria. The prevalence of infection (number of mosquitoes infected) between the treatments was calculated comparing the number of infected to uninfected larvae (A) or adults (B). Density of bacteria (CFU/mosquito) in larvae (C) and adults (D). The assay was repeated twice. Results display pooled data from each independent replicate. Box and whiskers show the median, the 25th and 75th percentiles and the minimum and maximum values.

Fig 5. The ΔompA mutant does not affect growth rates or development of mosquitoes.

The growth rate (time to pupation) (A) and development (percentage of L1 larvae to reach adulthood) (B) was observed in mosquitoes infected with C. neteri strains (WT, ΔompA mutant and ΔompA/ompA complement) reared in a mono-association. The experiment was done twice with a minimum of 15 individuals. Sample size for panel A indicates number of individuals, while for B indicates the number of replicate flasks. Each flask has 20 mosquitoes.

Integration of genes into the C. neteri chromosome

We undertook experiments to demonstrate the CRISPR/Cas9 gene-editing approaches can be used to integrate genes into the chromosome of non-model bacteria that associate with mosquitoes. We created two independent transgenic strains that had either a gene encoding mCherry fluorescence or a gene encoding resistance to the antibiotic gentamicin inserted into the bacterial chromosome. Before undertaking these integration experiments we confirmed that C. neteri was susceptible to gentamicin. These genes were integrated into the genome using the same gRNA that was used for deletional mutagenesis (S1 Table), and as such, these insertions also disrupted the ompA gene. Sequencing across the integration site indicated the insertion of these genes occurred within the ompA gene and thereby disrupted its function (Fig 6A and 6B). Continual subculturing was undertaken for both strains and molecular analysis indicated the stability of these lines for ten generations (Fig 6C and 6D). Expression of mCherry fluorescence and growth of the ΔompA::gentamicin strain on media containing gentamicin demonstrated the integrated genes were functional in vitro (Fig 6E and 6F).

Fig 6. Integration of mCherry and gentamicin into the C. neteri genome.

Sanger sequence across the integration site, stability of the inserted gene and in vitro expression of the inserted gene for the ΔompA::mCherry (A-C) and the ΔompA:: gentamicin (B-D) strains. The chromatogram shows the sequence spanning the inserted sites. Strains were continually subcultured for 10 passages and PCR was done to examine the stability of the insert (C; ΔompA::mCherry plus WT, D; ΔompA::gentamicin passaged with (ab+) or without (ab-) gentamicin plus WT). mCherry fluorescence (E) or ability to grow on selective media containing gentamicin (F) confirmed the expression of the transgene in vitro. Mosquitoes were inoculated with the C. neteri strains to confirm expression of the transgene in vivo. Dissected midgut infected with ΔompA::mCherry (left) or negative control (right; WT bacteria without expression plasmid) (G). Midguts were stained with phalloidin (green) and DAPI (blue). The scale bar is 30 μM. The WT and ΔompA::gentamicin C. neteri strains were fed to adult mosquitoes for 3 days in a sugar meal before gentamicin was administered to mosquitoes in sugar without bacteria (H). Mosquitoes were collected every second day and CFUs assessed. Pairwise comparisons were conducted at each time point using a T test.

To examine the functionally of the integrated genes in the mosquito we administered either WT, ΔompA::mCherry, or ΔompA::gentamicin to conventionally reared 3–4 day old adult female Ae. aegypti in a sugar meal for 3 days or larvae cleared of their microbiota. For gnotobiotic infection we used bacteria expressing mCherry from a plasmid. The dissected gut from 3–4 day old adults showed a higher percentage of WT bacteria compared to either of the integrated mutants. After screening midgut samples from each treatment, we found that mosquitoes infected with WT bacteria had the highest infection prevalence (69%) and that the mCherry and gentamicin knockin mutants were found only in 4% and 33% of the samples, respectively (S4 Fig, S4 Table). In addition, biofilms were seen mainly in mosquitoes infected with WT bacteria (31%) whilst midguts infected with mutants had few or no biofilms (0–2%) (S4 Fig, S4 Table). In sugar fed adult mosquitoes, ΔompA::mCherry bacteria were observed in the gut of mosquitoes with a distinct punctate distribution, whereas no signal was seen in autofluorescence controls (WT C. neteri infected mosquitoes) (Fig 6G). The C. neteri ompA::gentamicin was successfully rescued from mosquitoes reared on gentamicin and stably infected mosquitoes over time at a density of approximately 1x104 CFUs/mosquito. Consistent with our previous result (Fig 4B), WT bacteria initially infected mosquitoes at higher titers compared to the mutant (T test; day 0 P < 0.001). However, after 4 days rearing on antibiotic the total bacterial load in mosquitoes administered WT C. neteri was significantly reduced compared to the ΔompA::gentamicin (T test; day 4 P < 0.05) while the prevalence of mosquitoes with culturable microbiota was reduced to 80%. After 6 days rearing on antibiotic, the ΔompA::gentamicin density was significantly elevated compare to the WT (T test; day 6 P < 0.001) only one mosquito was infected, which had a low density infection (10 CFUs/mosquito) (Fig 6H).


We harnessed the CRISPR/Cas9 gene editing system to create knockout mutants in a C. neteri gut symbiont of Aedes mosquitoes to examine the role of bacterial genetics in biofilm formation and gut colonization. A deletion of the ompA gene of C. neteri decreased bacterial colonization of mosquitoes after infection in a mono-association. Strikingly, we found this effect was most pronounced in adult mosquitoes with more than half of the mosquitoes not possessing any culturable mutants, whereas there was no difference in prevalence of infection between the mutant and WT bacteria in larvae. The reduced prevalence of mutant bacteria in adults likely reflects differences in microbial colonization of each mosquito life stage. Larvae are continually subjected to bacteria in the larval water habitat while adults only have a short time frame to acquire bacteria from the aquatic environment immediately after eclosion. Alternatively, the reduced prevalence in adults could be due an impaired ability of mutant bacteria to be transstadially transmitted. Several bacterial species have been shown to exploit this process to transfer between life stages [6669]. When only analysing adult mosquitoes where bacteria did colonize the host, we saw no differences in the density of the mutant strain compared to the WT or complement, suggesting that ompA is acting at the colonization stage but has minimal effect on post-colonization processes. However, when examining midguts using fluorescent microscopy, in general, we observed reduced loads of the mutant strains. When quantifying bacterial load by CFU we used whole mosquitoes. It may be possible that mutant bacteria were residing in other tissues in the adult but poorly re-infected the midgut. If this occurred, it would indicate involvement of ompA in transstadial transmission. The greater variability seen in the prevalence of adults compared to the larval is consistent with other sequence-based studies that indicate adult stages have greater variability in species composition of their microbiota, whereas the microbiome of immature stages is similar to the microbiota in larval water habitat [25, 8, 70].

Mutant bacteria colonized mosquitoes at higher densities when administered to adults as opposed to larvae. There are several possible explanations for this finding. The first relates to the method of inoculation with adults being administered bacteria in a sugar meal while larvae were exposed to bacteria in their aquatic environment. The different inoculation process itself may influence titer but also when sugar feeding, adults had the opportunity for repeated infections whereas emerging adults only had a narrow window for inoculation as they did not have further access to the larval water habitat after eclosion. The second explanation relates to differences in the microbiome of these mosquitoes. The mosquitoes inoculated as adult were reared conventionally, and as such, had an intact microbiome, while larvae reared in the gnotobiotic system only possessed the individual Cedecea strains that were administered. For the latter group there was no opportunity for the native WT bacteria (either of the same or different species) to rescue the mutant phenotype. In the Sodalis-tsetse system, mutant bacteria were capable of infecting flies that had an intact microbiome but were unable to infect Sodalis-free tsetse flies [23], suggesting WT Sodalis facilitated colonization of the mutant strain. In mono-axenic infections, the C. neteri mutant strain was able to infect Ae. aegypti, indicating that ompA is not essential for infection in the mosquito-Cedecea system.

Our results, in conjunction with studies in the Sodalis-tsetse system [23, 24], suggests that biofilm formation may be a strategy employed by bacteria to colonize the gut of insects. In pathogenic infections in mammals, biofilms enable bacteria to colonize new niches, promote infection, and are associated with virulence [71]. Although less is known regarding the importance of biofilm formation in insects, in an artificial Pseudomonas-Drosophila infection model, biofilm formation was associated with virulence and host survival [21]. In a natural symbiotic association between bean bugs and Burkholderia, disruption of a purine biosynthesis gene in the bacterium reduce biofilm formation and colonization of the insect [20]. In mosquitoes, gut biofilm formation could also have implications for vector competence. Chromobacterium, which was isolated from Aedes mosquitoes, produced molecules that inhibited dengue virus only when grown in vitro as a biofilm but not when grown in a planktonic state [72], however it is unknown if biofilm formation occurred in vivo in the mosquito. Our data provide evidence that biofilms occur within the gut of mosquitoes and facilitate host colonization.

Although we have shown that the ompA gene of C. neteri is important for host colonization, we see no evidence that deletion of this gene alters mosquito development or growth rates. This is in contrast to the Riptortus-Burkholderia symbiosis whereby mutation of the purT gene in Burkholderia resulted in reduced growth rates and reduction in body weight of the host compared to insects that were infected with the WT bacterium [20]. The difference in our study to the findings in the Riptortus-Burkholderia symbiosis could be related to different requirements of the bean bug compared to the mosquito host as well as the different genes mutated in the symbionts. Our findings are consistent with a previous study in Ae. aegypti whereby an ompA mutant of E. coli did not influence growth when reared in a mono-association [73]. Using a similar gnotobiotic system that exploits the ability to sterilize mosquito eggs and rescue development by nutritional supplementation, several recent reports describe approaches to create bacteria-free mosquitoes [73, 74]. Here, we reared mosquitoes in a mono-association where they were only subjected to C. neteri. However, more than half the adult mosquitoes inoculated with the ΔompA mutant were not infected by bacteria, as evidenced by the inability to culture bacteria from these insects. Nevertheless, these mosquitoes had similar development and growth rates compared to mosquito possessing WT bacteria. The use of mutant bacteria that rescue development but have an impaired ability to colonize mosquitoes may provide a simple means to create axenic adult mosquitoes.

CRISPR/Cas9 gene editing has revolutionized genetic approaches in model and non-model bacteria [3143]. However, there has been limited use of this technology in symbiotic microbes of arthropods. Here we demonstrate that editing approaches functional in E. coli can be easily applied with minimal adaptation to phylogenetically related symbiotic bacteria that are found within the guts of mosquitoes. The application of CRISPR/Cas9 genome editing to gut-associated bacteria of mosquitoes has significant applied potential. Paratransgenesis strategies are being evaluated in a range of medical and agricultural systems to mitigate pathogen transmission from insect vectors, however, most approaches engineer symbionts by plasmid transformation [4955, 75] and where genome integration has been accomplished in symbionts [5861], it has often been done with technologies that did not allow for site specific integration. Paratransgenic approaches suitable for use in the field will need to stably integrate genes into the bacterial genome in a manner that does not compromise bacterial fitness. Exploiting the flexibility and specificity of the CRISPR/Cas9 system to integrate genes in intergenic regions of the bacterial chromosome will undoubtedly be beneficial for these applied approaches.

In summary, we have demonstrated that the CRISPR/Cas9 gene editing system can be applied to symbiotic bacteria that associate with eukaryotic hosts to interrogate the role of bacterial genes in host-microbe associations. We created knockout and knockin mutants by deleting and disrupting the ompA gene of C. neteri. The knockout mutant displayed a reduced ability to form biofilms and colonize the gut of Ae. aegypti mosquitoes in a mono-association demonstrating bacterial genetic factors are important determinants that influence colonization of mosquito guts. Aedes mosquitoes are becoming powerful systems to investigate the genetics of host-microbe interactions given the scientific community has simple and efficient approaches to alter both the microbes (this study) and mosquito host genome [76, 77] at their disposal, as well as methods to create mono-associated mosquito lines [7, 64]. Finally, rapid, efficient, and site specific gene editing approaches for gut bacteria that associate with mosquitoes will facilitate the development of novel paratransgenic approaches to control arthropod-borne disease [57].

Material and methods

Bacterial and mosquito strains

E. coli BL21(DE3) (NEB) and Cedecea neteri strain Alb1, previous isolated from a lab-reared colony of Ae. albopictus (Galveston) mosquitoes [7], were used in this study. To further classify the gut-associated bacteria we completed multilocus sequence typing [78]. DNA from the single colony was used as a template in a PCR to amplify genes for MLST analysis (S3 Table). Amplicons were resolved on a 1% agarose gel, extracted and purified, and Sanger sequenced. The atpD, infB, gyrB and rpoB genes were aligned separately, using the species diversity as in [79] with several Cedecea sp. sequences and then concatenated using seaview [79]. The phylogenetic tree was constructed using iqtree [80] under the general time-reversible (GTR) model with 1000 fast bootstrap replicates, which are shown as percentage branch support values (S4 Fig). The sequences of our isolate are available under accessions (MN329096 (atpD), MN329097 (gyrB), MN329098 (infB), MN329099 (rpoB). For gene editing and mosquito infections, cultures were grown in liquid LB media at 37°C with the appropriate antibiotic unless stated otherwise. Mosquitoes were reared in the UTMB insectary under conventional conditions or in a mono-association (described below).

CRISPR gene editing

Designing protospacer sequence and cloning: The E. coli BL21 ompA gene sequence was retrieved from NCBI (accession number LR536431). The C. neteri Alb1 ompA gene was PCR amplified and Sanger sequenced using primers (OmpA-F and OmpA-R, S3 Table), which were designed based on the Enterobacter cloacae ompA (accession number CP017990). Editing the ompA gene of E. coli and C. neteri was complete as described in Reisch and Prather [35]. Protospacer sequences for the ompA gene were designed using CHOPCHOP [81, 82]. To clone the protospacer sequences into pKDsgRNA-ack (S2 Table; Addgene plasmid #62654) we amplified the entire plasmid with primers that contained the protospacer sequence and this amplicon was self-ligated. This PCR was done using 0.5μM of each primer (S1 Table), 1x reaction buffer, 200μM dNTPs, 0.5U of Phire Host Start Taq polymerase (Thermo Scientific) and 200 ng of plasmid DNA as template. The cycling condition consisted of an initial denaturation step 98°C for 2 minutes, followed by 35 cycles of 98°C for 2 seconds, 58°C for 15 seconds, and 72°C for 2 minutes and 30 seconds, and then a final extension at 72°C for 10 minutes before holding at 16°C. The PCR products had a 15–17 bp overlapping sequence which was used to ligate the plasmid. The PCR product was digested with DpnI to remove any template plasmid. PCR products were then ligated by transformation into E. coli harbouring the Red/ET plasmid following the REPLACR mutagenesis protocol [83], thereby creating plasmids pKDsgRNA-Ec-ompA-1, pKDsgRNA-Ec-ompA-2, pKDsgRNA-Ent-ompA-1, and pKDsgRNA-Ent-ompA-2 (S2 Table). Colonies were screened for the protospacer insertion by PCR and confirmed by Sanger sequencing.

Knockout of ompA

The two protospacers were evaluated by transforming plasmids into either E. coli or C. neteri containing the pCas9-CR4 plasmid (S2 Table; Addgene plasmid 62655), which expressed Cas9 nuclease. Transformants were selected at 30°C on LB agar plate containing spectinomycin (50 μg/mL), chloramphenicol (34 μg/mL), and either with or without anhydrotetracycline (aTC; 100ng/mL). The escape rate was quantified by comparing colonies in the plates with or without aTC. The protospacer with a lack of or few escape mutants was used for further experiments. Colonies from the–aTC plate were grown overnight in LB broth with the appropriate antibiotic at 30°C. A 1:100 diluted overnight culture was (grown until 0.4 OD600) supplemented with 1.2% arabinose to induce the expression of λ-Red recombinase for 20 min. Cells were then transformed with 1–1.5 μg of double stranded donor DNA for homologous recombination. Donor DNA was created by PCR amplifying the flanking left arm (LA) and right arm (RA) from E. coli and C. neteri genomic DNA. Each arm had flanking regions of 250 bp homologous to the target DNA. The resulting fragment was assembled using Gibson assembly (NEB). The assembled product was amplified to generate full length dsDNA for transformation. Colonies were screened for mutations by colony PCR with primers flanking the integration site and positive clones were Sanger sequenced (S3 Table). Positive colonies were grown in LB broth and genomic DNA was isolated. For further validation, the flanking regions of deletion or insertions were amplified, and the PCR product Sanger sequenced.

Insertion of mCherry and gentamicin gene into C. neteri genome

The plasmid pKDsgRNA-Ent-ompA was transformed into C. neteri and the gene editing procedure was repeated as described above. To generated the donor sequence for homologous recominbination the mCherry or gentamicin sequence (driven by the AmTr promoter) and each homology arm were amplified and ligated. The assembled product was amplified to generate a full length dsDNA fragment for transformation.

Stability of insertion

The stability of the knockout ΔompA mutant and the knockin ompA::gentamicin and ompA::mCherry strains was assessed in LB media. The ompA::mCherry and knockout ΔompA mutant cultures were grown for 10 passages in LB broth. At each passage 40 μl of culture was transferred into 4ml fresh LB media. The ompA::gentamicin strain was grown with or without gentamicin (50 μg/mL). Genomic DNA was isolated from the 0, 2, 4, 6, 8 and 10th subculture and PCR that amplified across the integration site was performed.

Complementation of ompA mutant

Functional rescue of the ompA mutation was achieved by complementing the mutant with the WT gene. The WT ompA gene was amplified from C. neteri genomic DNA and cloned into the pRAM-mCherry vector [7] in front of the ompA promoter, thereby creating pRAM-mCherry-Ent-OmpA plasmid. The Sanger sequence-verified plasmid was transformed into the ΔompA mutant, thereby generating the ΔompA/ompA complement strain. Colonies that acquired the plasmid were selected on LB plates containing kanamycin (50 μg/mL).

In vitro characterization of C. neteri strains

To assess the impact of the gene deletion on bacterial growth the WT, ΔompA mutant and ΔompA/ompA complement were grown in LB broth and the density of bacteria (OD600) was quantified by spectrophotometer. A 1:100 dilution of an overnight culture was inoculated into a 5 ml LB broth in a 50 ml tube and incubated at 37°C for 24 hrs. At 2, 4, 6, 8, 10, 12 and 24 hours growth was recorded at OD600. The biofilm assay was performed as described previously [84, 85]. Briefly, biofilm formation by C. neteri strains was quantified on polystyrene microtiter plates after 72 h of incubation at 37°C by CV staining. Three independent experiments were performed, and the data were represented as CV OD570 after normalizing by CFUs.

Mosquito infections

Mono-association in Ae. aegypti mosquitoes were done using gnotobiotic infection procedure [7, 64], with slight modifications. Briefly, mosquito eggs were sterilized for 5 min in 70% ethanol, 3 min in 3% bleach+0.01% Coverage Plus NPD (Steris Corp.), 5 min in 70% ethanol then rinsed three times in sterile water. Eggs were vacuumed hatched for 30–45 min and left overnight at room temperature to hatch any remaining eggs. Exactly twenty L1 larvae were transferred to T175 flask containing 60 ml of sterile water and fed on alternative days with 60 μl of fish food (1 μg/μl). Larvae were inoculated with 1x107/ml of either the WT C. neteri, the ΔompA mutant or the ΔompA/ompA complement. The WT and ΔompA strains were transformed with the pRAM-mCherry plasmid [7] that conferred resistance to kanamycin (but did not possess a functional ompA gene). We also performed gnotobiotic infections with WT C. neteri, knockin mutants all expressing mCherry from a plasmid. In order to confirm that eggs were successfully sterilized, a T175 flask containing twenty L1 larvae were reared in identical fashion to mono-associations, albeit without bacterial supplementation. These larvae did not develop beyond the L2 stage, indicating our rearing process was free from contamination. To quantify bacteria, L4 larvae were collected, washed three times with 1X PBS, and then homogenized in 500 μl of 1X PBS and 50 μl of homogenate was plated on LB agar containing 50 μg/mL kanamycin. Similarly, adult mosquitoes were collected 3–4 days post emergence and bacterial infection was quantified in the same manner as larvae. In order to assess the growth of the mosquitoes, time to pupation and growth rate were observed. Time to pupation was determined by quantifying the number of pupae each day post hatching, while survival to adulthood was calculated by quantifying the number of L1 larvae that reached adulthood. The experiment was repeated three times.

Reinfection of knockin mutants to mosquitoes

Knockin mutants were administered to 3–4 days adult Ae. aegypti in a sugar meal. These mosquitoes were reared under normal laboratory condition. Mosquitoes were fed with 1x107 of WT or the ΔompA::gentamicin strain for three days in 10% sucrose solution. After three days, mosquitoes were either administered sugar supplemented with gentamicin (50 μg/mL) or sugar without antibiotic. CFUs were determined at days 0, 2, 4, and 6 dpi by plating homogenized mosquitoes (N = 10) on LB agar. Similarly, the ΔompA::mCherry and WT C. neteri were fed to mosquitoes and midguts were dissected to assess colonization of bacteria in the tissue. For visualization of bacteria, midguts were fixed in 1% paraformaldehyde (PFA) in 1X PBS for 30 minutes and permeabilized with 0.01% Triton X-100 in 1X PBS for 20 min. The tissues were stained with 1:250 diluted Phalloidin (Sigma) for 20 minutes and samples were washed twice with 1X PBS for 10 minutes. Finally, midguts were then stained with 1:500 diluted DAPI (Invitrogen) for 10 min. Samples were transferred to slides and mounted with ProLong Gold Antifade (Invitrogen). The slides were observed using a Revolve FL microscope (ECHOLAB).

Supporting information

S1 Fig. Midgut infection of C. neteri and E. coli in mono-associations of Aedes mosquitoes.

Dissected gut tissue showing the conglomeration of bacterial cells when infected in mono-association in Aedes mosquitoes with C. neteri (A-C). However, E. coli (D-F) and ΔompA (G-I) could not be seen in the midgut. Images were captured from the dissected midguts of different mosquitoes. Bacteria possessed the pRAM-mCherry plasmid which expressed the mCherry fluorescent protein.


S2 Fig. Phylogenetic analysis of WT and mutants.

Multilocus sequence analysis according to [78] indicates isolates to be members of the C. neteri species; the MLST genes were amplified in the wild type isolate and two mutants as shown in the tree (red).


S3 Fig. Average CFUs of mosquito infection.

Average CFU recovered in adult mosquitoes infected with C. neteri strains (WT, ΔompA mutant and ΔompA/ompA complement) reared in a mono-association using a gnotobiotic rearing approach. The uninfected mosquitoes were removed from the analysis. Box and whiskers show the 25th and 75th percentiles and the minimum and maximum values, respectively.


S4 Fig. Mono-association infection and biofilm assessment.

L1 axenic larvae were infected with WT C. neteri (A), ΔompA::mCherry (B) and ΔompA::gentamicin (C) and adults gut were analysed for presence of bacterial conglomerations (biofilm formation). For each treatment, 15–18 midguts were screened. Scale bar 130 μm.


S1 Table. gRNA sequence used in this study.


S2 Table. Plasmids and bacterial strains used in this study.


S4 Table. Infection and biofilm prevalence of mosquito midguts when reared in a mono-association.


S1 Appendix. Multiple sequence alignment of WT and mutant ompA sequences of C. neteri and E. coli.



We would like to thank the UTMB insectary core for providing mosquitoes. We also wish to thank Dr Ulrike Munderloh (University of Minnesota) for the kind gift of the pRAM plasmid.


  1. 1. Osei-Poku J, Mbogo CM, Palmer WJ, Jiggins FM. Deep sequencing reveals extensive variation in the gut microbiota of wild mosquitoes from Kenya. Molecular Ecology. 2012;21(20):5138–50. pmid:22988916
  2. 2. Boissière A, Tchioffo MT, Bachar D, Abate L, Marie A, Nsango SE, et al. Midgut microbiota of the malaria mosquito vector Anopheles gambiae and interactions with Plasmodium falciparum infection. PLoS Pathogens. 2012;8(5):e1002742. pmid:22693451
  3. 3. David MR, Santos LMBD, Vicente ACP, Maciel-de-Freitas R. Effects of environment, dietary regime and ageing on the dengue vector microbiota: evidence of a core microbiota throughout Aedes aegypti lifespan. Memórias do Instituto Oswaldo Cruz. 2016;111(9):577–87. pmid:27580348
  4. 4. Muturi EJ, Kim C-H, Bara J, Bach EM, Siddappaji MH. Culex pipiens and Culex restuans mosquitoes harbor distinct microbiota dominated by few bacterial taxa. Parasites & Vectors. 2016;9(1):18. pmid:26762514
  5. 5. Hughes GL, Dodson BL, Johnson RM, Murdock CC, Tsujimoto H, Suzuki Y, et al. Native microbiome impedes vertical transmission of Wolbachia in Anopheles mosquitoes. Proceedings of the National Academy of Sciences of the United States of America. 2014;111(34):12498–503. pmid:25114252
  6. 6. Coon KL, Brown MR, Strand MR. Mosquitoes host communities of bacteria that are essential for development but vary greatly between local habitats. Molecular Ecology. 2016;25(22):5806–26. pmid:27718295
  7. 7. Hegde S, Khanipov K, Albayrak L, Golovko G, Pimenova M, Saldaña MA, et al. Microbiome interaction networks and community structure from laboratory-reared and field-collected Aedes aegypti, Aedes albopictus, and Culex quinquefasciatus mosquito vectors. Frontiers in Microbiology. 2018;9:715.
  8. 8. Wang Y, Gilbreath TM, Kukutla P, Yan G, Xu J. Dynamic gut microbiome across life history of the malaria mosquito Anopheles gambiae in Kenya. PLoS ONE. 2011;6(9):e24767. pmid:21957459
  9. 9. Weiss BL, Aksoy S. Microbiome influences on insect host vector competence. Trends in Parasitology. 2011;27(11):514–22. pmid:21697014
  10. 10. Saldaña MA, Hegde S, Hughes GL. Microbial control of arthropod-borne disease. Memórias do Instituto Oswaldo Cruz. 2017;112(2):81–93. pmid:28177042
  11. 11. Hegde S, Rasgon JL, Hughes GL. The microbiome modulates arbovirus transmissionin mosquitoes. Current Opinion in Virology. 2015;15:97–102. pmid:26363996
  12. 12. Audsley MD, Ye YH, McGraw EA. The microbiome composition of Aedes aegypti is not critical for Wolbachia-mediated inhibition of dengue virus. PLoS Neglected Tropical Diseases. 2017;11(3):e0005426. pmid:28267749
  13. 13. Muturi EJ, Bara JJ, Rooney AP, Hansen AK. Midgut fungal and bacterial microbiota of Aedes triseriatus and Aedes japonicus shift in response to La~Crosse virus infection. Molecular Ecology. 2016;25(16):4075–90. pmid:27357374
  14. 14. Kumar S, Molina-Cruz A, Gupta L, Rodrigues J, Barillas-Mury C. A peroxidase/dual oxidase system modulates midgut epithelial immunity in Anopheles gambiae. Science. 2010;327(5973):1644–8. pmid:20223948
  15. 15. Pang X, Xiao X, Liu Y, Zhang R, Liu J, Liu Q, et al. Mosquito C-type lectins maintain gut microbiome homeostasis. Nature Microbiology. 2016;1(5):16023. pmid:27170846
  16. 16. Short SM, Mongodin EF, MacLeod HJ, Talyuli OAC, Dimopoulos G. Amino acid metabolic signaling influences Aedes aegypti midgut microbiome variability. PLoS Neglected Tropical Diseases. 2017;11(7):e0005677–29. pmid:28753661
  17. 17. Stathopoulos S, Neafsey DE, Lawniczak MKN, Muskavitch MAT, Christophides GK. Genetic dissection of Anopheles gambiae gut epithelial responses to Serratia marcescens. PLOS Pathogens. 2014;10(3):e1003897. pmid:24603764
  18. 18. Xiao X, Yang L, Pang X, Zhang R, Zhu Y, Wang P, et al. A Mesh-Duox pathway regulates homeostasis in the insect gut. Nature Microbiology. 2017;2(5):17020. pmid:28248301
  19. 19. Powell JE, Leonard SP, Kwong WK, Engel P, Moran NA. Genome-wide screen identifies host colonization determinants in a bacterial gut symbiont. Proceedings of the National Academy of Sciences of the United States of America. 2016:201610856. pmid:27849596
  20. 20. Kim JK, Kwon JY, Kim S-K, Han SH, Won YJ, Lee J-H, et al. Purine biosynthesis, biofilm formation, and persistence of an snsect-microbe gut symbiosis. Appl Environ Microbiol. 2014;80(14):4374–82. pmid:24814787
  21. 21. Mulcahy H, Sibley CD, Surette MG, Lewenza S. Drosophila melanogaster as an animal model for the study of Pseudomonas aeruginosa biofilm infections in vivo. PLOS Pathogens. 2011;7(10):e1002299–14. pmid:21998591
  22. 22. Maltz M, Graf J. The type II secretion system is essential for erythrocyte lysis and gut colonization by the leech digestive tract symbiont Aeromonas veronii. Applied and Environmental Microbiology. 2011;77(2):597–603. pmid:21097598
  23. 23. Weiss BL, Wu Y, Schwank JJ, Tolwinski NS, Aksoy S. An insect symbiosis is influenced by bacterium-specific polymorphisms in outer-membrane protein A. Proceedings of the National Academy of Sciences of the United States of America. 2008;105(39):15088–93. pmid:18815366
  24. 24. Maltz MA, Weiss BL, O apos Neill M, Wu Y, Aksoy S. OmpA-mediated biofilm formation is essential for the commensal bacterium Sodalis glossinidius to colonize the tsetse fly gut. Applied and Environmental Microbiology. 2012;78(21):7760–8. pmid:22941073
  25. 25. Riehle MA, Jacobs-Lorena M. Using bacteria to express and display anti-parasite molecules in mosquitoes: current and future strategies. Insect Biochemistry and Molecular Biology. 2005;35(7):699–707. pmid:15894187
  26. 26. Dennison NJ, Saraiva RG, Cirimotich CM, Mlambo G, Mongodin EF, Dimopoulos G. Functional genomic analyses of Enterobacter, Anopheles and Plasmodium reciprocal interactions that impact vector competence. Malaria Journal. 2016;15(1):425. Epub 3. pmid:27549662
  27. 27. Pei D, Jiang J, Yu W, Kukutla P, Uentillie A, Xu J. The waaL gene mutation compromised the inhabitation of Enterobacter sp. Ag1 in the mosquito gut environment. Parasites & Vectors. 2015:1–10. pmid:26306887
  28. 28. Enomoto S, Chari A, Clayton AL, Dale C. Quorum sensing attenuates virulence in Sodalis praecaptivus. Cell Host &Microbe. 2017;21(5):629–36.e5. pmid:28494244
  29. 29. Dale C, Young SA, Haydon DT, Welburn SC. The insect endosymbiont Sodalis glossinidius utilizes a type III secretion system for cell invasion. Proceedings of the National Academy of Sciences. 2001;98(4):1883–8. pmid:11172045
  30. 30. Selle K, Barrangou R. Harnessing CRISPR–Cas systems for bacterial genome editing. Trends in Microbiology. 2015; 23(4):225–32. pmid:25698413
  31. 31. Sander JD, Joung JK. CRISPR-Cas systems for editing, regulating and targeting genomes. Nature Biotechnology. 2014;32(4):347–55. pmid:24584096
  32. 32. Barrangou R, van Pijkeren JP. Exploiting CRISPR-Cas immune systems for genome editing in bacteria. Current Opinion in Biotechnology. 2016;37:61–8. pmid:26629846
  33. 33. Jiang Y, Chen B, Duan C, Sun B, Yang J, Yang S. Multigene editing in the Escherichia coli genome via the CRISPR-Cas9 system. Applied and Environmental Microbiology. 2015;81(7):2506–14. pmid:25636838
  34. 34. Jiang W, Bikard D, Cox D, Zhang F, Marraffini LA. RNA-guided editing of bacterial genomes using CRISPR-Cas systems. Nature Biotechnology. 2013;31(3):233–9. pmid:23360965
  35. 35. Reisch CR, Prather KLJ. The no-SCAR (Scarless Cas9 Assisted Recombineering) system for genome editing in Escherichia coli. Scientific Reports. 2015;5:15096. pmid:26463009
  36. 36. Li Y, Lin Z, Huang C, Zhang Y, Wang Z, Tang Y-j, et al. Metabolic engineering of Escherichia coli using CRISPR–Cas9 meditated genome editing. Metabolic Engineering. 2015;31:13–21. pmid:26141150
  37. 37. Ronda C, Pedersen LE, Sommer MOA, Nielsen AT. CRMAGE: CRISPR Optimized MAGE Recombineering. Scientific Reports. 2016;6(1):1200. pmid:26797514
  38. 38. Tong Y, Robertsen HL, Blin K, Weber T, Lee SY. CRISPR-Cas9 Toolkit for Actinomycete Genome Editing. Methods in molecular biology 2018;1671(1):163–84. pmid:29170959
  39. 39. Oh J-H, van Pijkeren JP. CRISPR-Cas9-assisted recombineering in Lactobacillus reuteri. Nucleic Acids Research. 2014;42(17):e131–e. pmid:25074379
  40. 40. Mougiakos I, Bosma EF, Weenink K, Vossen E, Goijvaerts K, van der Oost J, et al. Efficient genome editing of a facultative thermophile using mesophilic spCas9. ACS Synthetic Biology. 2017;6(5):849–61. pmid:28146359
  41. 41. Li K, Cai D, Wang Z, He Z, Chen S. Development of an efficient genome editing tool in Bacillus licheniformis ssing CRISPR-Cas9 nickase. Applied and Environmental Microbiology. 2018:AEM.02608-17. pmid:29330178
  42. 42. Jiang Y, Qian F, Yang J, Liu Y, Dong F, Xu C, et al. CRISPR-Cpf1 assisted genome editing of Corynebacterium glutamicum. Nat Commun. 2017;8:15179. Epub 2017/05/05. pmid:28469274
  43. 43. Cobb RE, Wang Y, Zhao H. High-efficiency multiplex genome editing of Streptomyces species using an engineered CRISPR/Cas system. ACS Synth Biol. 2015;4(6):723–8. Epub 2014/12/03. pmid:25458909
  44. 44. Waller MC, Bober JR, Nair NU, Beisel CL. Toward a genetic tool development pipeline for host-associated bacteria. Current Opinion in Microbiology. 2017;38:156–64. pmid:28624690
  45. 45. Wilke ABB, Marrelli MT. Paratransgenesis: a promising new strategy for mosquito vector control. Parasites & Vectors. 2015;8(1):391–19. pmid:26104575
  46. 46. Arora AK, Douglas AE. Hype or opportunity? Using microbial symbionts in novel strategies for insect pest control. Journal of insect physiology. 2017;103:10–7. pmid:28974456
  47. 47. Ricci I, Damiani C, Capone A, DeFreece C, Rossi P, Favia G. Mosquito/microbiota interactions: from complex relationships to biotechnological perspectives. Current Opinion in Microbiology. 2012;15(3):278–84. pmid:22465193
  48. 48. Beard CB, Mason PW, Aksoy S, Tesh RB, Richards FF. Transformation of an insect symbiont and expression of a foreign gene in the Chagas’ disease vector Rhodnius prolixus. American Journal of Tropical Medicine and Hygiene. 1992;46(2):195–200. pmid:1539755
  49. 49. Bisi DC, Lampe DJ. Secretion of anti-Plasmodium effector proteins from a natural Pantoea agglomerans isolate by using PelB and HlyA secretion signals. Applied and Environmental Microbiology. 2011;77(13):4669–75. pmid:21602368
  50. 50. Hughes GL, Allsopp PG, Webb RI, Yamada R, Iturbe-Ormaetxe I, Brumbley SM, et al. Identification of yeast associated with the planthopper, Perkinsiella saccharicida: potential applications for Fiji leaf gall control. Current Microbiology. 2011;63(4):392–401. pmid:21850475
  51. 51. Medina F, Li H, Vinson SB, Coates CJ. Genetic transformation of midgut bacteria from the red imported fire ant (Solenopsis invicta). Current Microbiology. 2009;58(5):478–82. pmid:19159973
  52. 52. Bextine B, Lauzon C, Potter S, Lampe D, Miller TA. Delivery of a genetically marked Alcaligenes sp. to the glassy-winged sharpshooter for use in a paratransgenic control strategy. Current Microbiology. 2004;48(5):327–31. pmid:15060727
  53. 53. Hurwitz I, Hillesland H, Fieck A, Das P, Durvasula R. The paratransgenic sand fly: A platform for control of Leishmania transmission. Parasites & Vectors. 2011;4(1):82. pmid:21595907
  54. 54. Durvasula RV, Gumbs A, Panackal A, Kruglov O, Taneja J, Kang AS, et al. Expression of a functional antibody fragment in the gut of Rhodnius prolixus via transgenic bacterial symbiont Rhodococcus rhodnii. Medical and Veterinary Entomology. 1999;13(2):115–9. pmid:10484156
  55. 55. Wang S, Ghosh AK, Bongio N, Stebbings KA, Lampe DJ, Jacobs-Lorena M. Fighting malaria with engineered symbiotic bacteria from vector mosquitoes. Proceedings of the National Academy of Sciences of the United States of America. 2012;109(31):12734–9. pmid:22802646
  56. 56. Wu SC-Y, Maragathavally KJ, Coates CJ, Kaminski JM. Steps toward targeted insertional mutagenesis with class II transposable elements. Methods in molecular biology 2008;435:139–51. pmid:18370073
  57. 57. Tikhe CV, Martin TM, Howells A, Delatte J, Husseneder C. Assessment of genetically engineered Trabulsiella odontotermitis as a ‘Trojan Horse’ for paratransgenesis in termites. BMC Microbiology. 2016;16(1):355. pmid:27595984
  58. 58. Pittman GW, Brumbley SM, Allsopp PG, O apos Neill SL. Assessment of gut bacteria for a paratransgenic approach to control Dermolepida albohirtum larvae. Applied and Environmental Microbiology. 2008;74(13):4036–43. pmid:18456847
  59. 59. Pontes MH, Dale C. Lambda red-mediated genetic modification of the insect endosymbiont Sodalis glossinidius. Applied and Environmental Microbiology. 2011;77(5):1918–20. pmid:21216910
  60. 60. Wang S, Dos-Santos ALA, Huang W, Liu KC, Oshaghi MA, Wei G, et al. Driving mosquito refractoriness to Plasmodium falciparum with engineered symbiotic bacteria. Science (New York, NY). 2017;357(6358):1399–402. pmid:28963255
  61. 61. Dotson EM, Plikaytis B, Shinnick TM, Durvasula RV, Beard CB. Transformation of Rhodococcus rhodnii, a symbiont of the Chagas disease vector Rhodnius prolixus, with integrative elements of the L1 mycobacteriophage. Infection, genetics and evolution. 2003;3(2):103–9. pmid:12809804
  62. 62. Bextine B, Lampe D, Lauzon C, Jackson B, Miller TA. Establishment of a genetically marked insect-derived symbiont in multiple host plants. Curr Microbiol. 2005;50(1):1–7. pmid:15723145
  63. 63. Wu P, Sun P, Nie K, Zhu Y, Shi M, Xiao C, et al. A gut commensal bacterium promotes mosquito permissiveness to arboviruses. Cell Host & Microbe. 2019;25(1):101–12.e5. pmid:30595552
  64. 64. Coon KL, Vogel KJ, Brown MR, Strand MR. Mosquitoes rely on their gut microbiota for development. Molecular Ecology. 2014;23(11):2727–39. pmid:24766707
  65. 65. Zalewska-Piatek B, Wilkanowicz S, Bruzdziak P, Piatek R, Kur J. Biochemical characteristic of biofilm of uropathogenic Escherichia coli Dr(+) strains. Microbiol Res. 2013;168(6):367–78. Epub 2013/02/05. pmid:23375236
  66. 66. Lindh JM, Borg-Karlson AK, Faye I. Transstadial and horizontal transfer of bacteria within a colony of Anopheles gambiae (Diptera: Culicidae) and oviposition response to bacteria-containing water. Acta Tropica. 2008;107(3):242–50. pmid:18671931
  67. 67. Bäckman S, Näslund J, Forsman M, Thelaus J. Transmission of tularemia from a water source by transstadial maintenance in a mosquito vector. Scientific Reports. 2015;5:7793. pmid:25609657
  68. 68. Chavshin AR, Oshaghi MA, Vatandoost H, Yakhchali B, Zarenejad F, Terenius O. Malpighian tubules are important determinants of Pseudomonas transstadial transmission and longtime persistence in Anopheles stephensi. Parasites & Vectors. 2015;8(1):36. pmid:25604581
  69. 69. Chen S, Bagdasarian M, Walker ED. Elizabethkingia anophelis: Molecular Manipulation and Interactions with Mosquito Hosts. Applied and Environmental Microbiology. 2015;81(6):2233. pmid:25595771
  70. 70. Coon KL, Brown MR, Strand MR. Gut bacteria differentially affect egg production in the anautogenous mosquito Aedes aegypti and facultatively autogenous mosquito Aedes atropalpus (Diptera: Culicidae). Parasites & Vectors. 2016;9(1):375. pmid:27363842
  71. 71. Ribet D, Cossart P. How bacterial pathogens colonize their hosts and invade deeper tissues. Microbes and infection. 2015;17(3):173–83. pmid:25637951
  72. 72. Ramirez JL, Short SM, Bahia AC, Saraiva RG, Dong Y, Kang S, et al. Chromobacterium Csp_P reduces malaria and dengue infection in vector mosquitoes and has entomopathogenic and in vitro anti-pathogen activities. PLoS Pathogens. 2014;10(10):e1004398. pmid:25340821
  73. 73. Valzania L, Martinson VG, Harrison RE, Boyd BM, Coon KL, Brown MR, et al. Both living bacteria and eukaryotes in the mosquito gut promote growth of larvae. PLoS Neglected Tropical Diseases. 2018;12(7):e0006638. pmid:29979680
  74. 74. Correa MA, Matusovsky B, Brackney DE, Steven B. Generation of axenic Aedes aegypti demonstrate live bacteria are not required for mosquito development. Nature Communications. 2018;9(1):R37. pmid:30367055
  75. 75. Favia G, Ricci I, Damiani C, Raddadi N, Crotti E, Marzorati M, et al. Bacteria of the genus Asaia stably associate with Anopheles stephensi, an Asian malarial mosquito vector. Proceedings of the National Academy of Sciences. 2007;104(21):9047–51. pmid:17502606
  76. 76. Chaverra-Rodriguez D, Macias VM, Hughes GL, Pujhari S, Suzuki Y, Peterson DR, et al. Targeted delivery of CRISPR-Cas9 ribonucleoprotein into arthropod ovaries for heritable germline gene editing. Nature Communications. 2018;9(1):245.
  77. 77. Li M, Bui M, Yang T, Bowman CS, White BJ, Akbari OS. Germline Cas9 expression yields highly efficient genome engineering in a major worldwide disease vector, Aedes aegypti. Proceedings of the National Academy of Sciences of the United States of America. 2017;114(49):E10540–E9. pmid:29138316
  78. 78. Brady C, Cleenwerck I, Venter S, Coutinho T, De Vos P. Taxonomic evaluation of the genus Enterobacter based on multilocus sequence analysis (MLSA): Proposal to reclassify E. nimipressuralis and E. amnigenus into Lelliottia gen. nov. as Lelliottia nimipressuralis comb. nov. and Lelliottia amnigena comb. nov., respectively, E. gergoviae and E. pyrinus into Pluralibacter gen. nov. as Pluralibacter gergoviae comb. nov. and Pluralibacter pyrinus comb. nov., respectively, E. cowanii, E. radicincitans, E. oryzae and E. arachidis into Kosakonia gen. nov. as Kosakonia cowanii comb. nov., Kosakonia radicincitans comb. nov., Kosakonia oryzae comb. nov. and Kosakonia arachidis comb. nov., respectively, and E. turicensis, E. helveticus and E. pulveris into Cronobacter as Cronobacter zurichensis nom. nov., Cronobacter helveticus comb. nov. and Cronobacter pulveris comb. nov., respectively, and emended description of the genera Enterobacter and Cronobacter. Systematic and Applied Microbiology. 2013;36(5):309–19. pmid:23632228
  79. 79. Gouy M, Guindon S, Gascuel O. SeaView version 4: A multiplatform graphical user interface for sequence alignment and phylogenetic tree building. Mol Biol Evol. 2010;27(2):221–4. Epub 2009/10/27. pmid:19854763
  80. 80. Nguyen LT, Schmidt HA, von Haeseler A, Minh BQ. IQ-TREE: a fast and effective stochastic algorithm for estimating maximum-likelihood phylogenies. Mol Biol Evol. 2015;32(1):268–74. Epub 2014/11/06. pmid:25371430
  81. 81. Montague TG, Cruz JM, Gagnon JA, Church GM, Valen E. CHOPCHOP: a CRISPR/Cas9 and TALEN web tool for genome editing. Nucleic Acids Research. 2014;42(W1):W401–W7. pmid:24861617
  82. 82. Labun K, Montague TG, Gagnon JA, Thyme SB, Valen E. CHOPCHOP v2: a web tool for the next generation of CRISPR genome engineering. Nucleic Acids Research. 2016;44(W1):W272–W6. pmid:27185894
  83. 83. Trehan A, Kiełbus M, Czapinski J, Stepulak A, Huhtaniemi I, Rivero-Müller A. REPLACR-mutagenesis, a one- step method for site-directed mutagenesis by recombineering. Scientific Reports. 2015:1–9. pmid:26750263
  84. 84. Kozlova EV, Khajanchi BK, Popov VL, Wen J, Chopra AK. Impact of QseBC system in c-di-GMP-dependent quorum sensing regulatory network in a clinical isolate SSU of Aeromonas hydrophila. Microbial pathogenesis. 2012;53(3–4):115–24. pmid:22664750
  85. 85. O’ Toole GA, Kolter R. Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Molecular Microbiology. 1998;30(2):295–304. pmid:9791175