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Additional file 7: Fig. S 5. Body size (estimated by forewing length) of adult males (left) and females (right) emerging from experimental plates containing gnotobiotic larvae recolonized by their native microbiota. Adults emerging from trays containing larvae reared conventionally in our standard laboratory colony served as the positive control. A minimum of 600 larvae were assayed per treatment. Box-and-whisker plots show high, low, and median values, with lower and upper edges of each box denoting first and third quartiles, respectively. Plots of the same color represent results from replicate plates (or trays) using larvae derived from independent cohorts of eggs. No significant differences between replicates were detected for adult females (NS). Size likewise did not differ between treatments for adult females (NS; Mann���Whitney U test, p > 0.05) after pooling replicates, while gnotobiotic adult males were marginally smaller than conventional males even after accounting for variation between replicates (*; Mann���Whitney U test, p

Additional file 6: Fig. S 4. PCR analysis of W. smithii larvae under axenic, gnotobiotic, or conventional conditions. Axenic first instars from surface-sterilized eggs were hatched in closed containers containing sterile water and sterilized diet. Gnotobiotic larvae recolonized by their native microbiota were produced by feeding axenic larvae sterilized diet plus material from a glycerol stock containing the mixed community of bacteria present under conventional rearing conditions. Conventional first instars were hatched in open containers containing distilled water and standard diet. For each treatment, DNA was isolated from a pooled sample of at least 10 larvae after surface sterilization as described herein (see ���Methods���). DNA samples were then used as template with universal bacterial 16S rRNA gene or fungal ITS primers. The agarose gel shows ethidium bromide-stained PCR products. Lane 1, molecular mass markers labeled in base pairs (bp); Lanes 2���4, universal 16S rRNA gene primers plus DNA from axenic larvae; Lanes 5���7, universal ITS primers plus DNA from axenic larvae; Lanes 8���10, universal 16S rRNA gene primers plus DNA from gnotobiotic larvae; Lanes 11���13, universal 16S rRNA gene primers plus DNA from conventional larvae.

Additional file 4: Supplementary Figure 2. (Left) Relative abundance of bacterial genera in Ae. aegypti donor pools and recipient samples. Biological replicates were pooled for the bar graphs presented. Low abundance genera (

Additional file 3: Supplementary Figure 1. Significantly negative correlation between relative abundance of ASVs belonging to the bacterial family Enterobacteriaceae and ASVs belonging to the genus Serratia across all Ae. aegypti and Cx. quinquefasciatus donor individuals (Spearman’s rank test, P < 0.05).

Additional file 4: Fig. S 2. Overlap between ASVs in W. smithii larvae and water collected from naturally occurring pitchers in the field and our standard laboratory colony. Values in parentheses indicate the percentage of total ASVs represented in a given sample type or combination of sample types.

Additional file 3: Fig. S 1. Rarefaction data from Illumina sequences of 16S rRNA gene amplicon libraries prepared from W. smithii larvae and water collected from naturally occurring pitchers in the field (left) or our standard laboratory colony (right).

Additional file 5: Fig. S 3. Rarefaction data from Illumina sequences of 16S rRNA gene amplicon libraries prepared from W. smithii egg, water, larval, and adult samples collected from our standard laboratory colony.

Additional file 6: Supplementary Figure 4. Average Bray-Curtis dissimilarity between (b/w) donor pools versus between a given donor pool and recipient samples. Mean values ± standard errors are shown. Asterisks (***) indicate comparisons for which the average dissimilarity between a given donor pool and group of recipient samples was significantly higher than that expected as a result of the transplantation procedure itself (i.e., between donor pools) (Dunn’s test with Bonferroni correction, P < 0.0125).

Additional file 5: Supplementary Figure 3. (Left) Relative abundance of bacterial genera in Cx. quinquefasciatus donor pools and recipient samples. Biological replicates were pooled for the bar graphs presented. Low abundance genera (

Additional file 7: Supplementary Figure 5. Significantly negative correlation between relative abundance of ASVs belonging to the bacterial family Enterobacteriaceae and ASVs belonging to the genus Serratia across all recipient Ae. aegypti adults (Spearman’s rank test, P < 0.05).

Most species of mosquitoes are detritivores that feed on decaying plant and animal materials in their aquatic environment. Studies of several detritivorous mosquito species indicate that they host relatively low diversity communities of microbes that are acquired from the environment while feeding. Our recent results also indicate that detritivorous species require a living gut microbiota in order to grow beyond the first instar. Less well known is that some mosquitoes, including those belonging to the genus Toxorhynchites, are predators that feed on other species of mosquitoes and other nektonic prey. In this study, we asked whether predaceous Toxorhynchites amboinensis larvae still require living microbes in their gut in order to develop. Using the detritivorous mosquito Aedes aegypti as prey, we found that T. amboinensis larvae harbour bacterial communities that are highly similar to that of their prey. Functional assays showed that T. amboinensis first instars provided axenic (i.e. bacteria-free) prey failed to develop, while two bacterial species present in gnotobiotic (i.e. colonized by one or more known bacterial species) prey successfully colonized the T. amboinensis gut and rescued development. Axenic T. amboinensis larvae also displayed defects in growth consistent with previously identified roles for microbe-mediated gut hypoxia in nutrient acquisition and assimilation in A. aegypti. Collectively, these results support a conserved role for gut microbes in regulating the development of mosquitoes with different feeding strategies.

The mosquito microbiome is critical for host development and plays a major role in many aspects of mosquito biology. While the microbiome is commonly dominated by a small number of genera, there is considerable variation in composition among mosquito species, life stages, and geography. How the host controls and is affected by this variation is unclear. Using microbiome transplant experiments, we asked whether there were differences in transcriptional responses when mosquitoes of different species were used as microbiome donors. We used microbiomes from four different donor species spanning the phylogenetic breadth of the Culicidae, collected either from the laboratory or field. We found that when recipients received a microbiome from a donor reared in the laboratory, the response was remarkably similar regardless of donor species. However, when the donor had been collected from the field, far more genes were differentially expressed. We also found that while the transplant procedure did have some effect on the host transcriptome, this is likely to have had a limited effect on mosquito fitness. Overall, our results highlight the possibility that variation in mosquito microbiome communities are associated with variability in host-microbiome interactions and further demonstrate the utility of the microbiome transplantation technique.