Selected gut bacteria promote mosquito larval growth through the regulation of ionic microenvironment and protease expression

Jhuma Samanta; Sibnarayan Datta

doi:10.1016/j.isci.2025.113756

. 2025 Oct 13;28(11):113756. doi: 10.1016/j.isci.2025.113756

Selected gut bacteria promote mosquito larval growth through the regulation of ionic microenvironment and protease expression

Jhuma Samanta ¹, Sibnarayan Datta ^1,^2,^∗

PMCID: PMC12616018 PMID: 41244566

Summary

The anterior midgut of mosquito larvae harbors microbiota. Besides, the hyperalkaline (pH∼10–11) anterior midgut also provides an optimal ionic microenvironment for protein digestion. Although gut microbiota is known to promote larval growth and development, the underlying host-microbe mechanisms remain less understood. Using the Aedes albopictus axenic-gnotobiotic model, we show that anterior midgut alkalinization necessitates colonization by “live” bacteria belonging to selected genera. Results of our bacteriophage-mediated gut bacteria decolonization studies suggest that the colonizing bacteria also mediate the expression of crucial larval proteases. Additionally, the colonizing bacteria contribute a significant proportion of proteases (including serine proteases, aminopeptidases, etc.) to the larval gut, which are crucial for the digestion of dietary proteins. Collectively, these findings reveal that specific members of the gut microbiota promote larval growth and development by facilitating larval protein metabolism through modulation of the midgut microenvironment. These findings may have important implications for the development of targeted vector-control strategies.

Subject areas: Entomology, Biochemistry, Microbiology

Graphical abstract

Highlights

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Colonization with selected bacteria is needed for Aedes larval midgut hyperalkalinization
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These colonizing bacteria modulate expression of key larval proteases in the gut
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These bacteria also contribute proteases essential for digestion of dietary proteins

Entomology; Biochemistry; Microbiology

Introduction

Insects are among the most diverse and adaptable organisms on Earth. Many species have evolved strategies to maximize exploitation of available resources by shifting ecological niches and feeding behaviors throughout their life cycle.¹^,² The natural ability of insects to feed upon and digest a wide variety of substrates has been linked to the unique physiology and microbiology of their gut, and therefore, the insect gut has been a subject of intense research for a long time.³^,⁴ According to their feeding habits, gut microenvironment in herbivorous insects is more alkaline as compared to carnivorous insects.⁵ Surprisingly, among holometabolous insect larvae from the orders Diptera, Lepidoptera, Coleoptera, and Trichoptera, some species exhibit exceptionally high gut pH levels (pH 9.0–12.0), whereas some others display extremely low pH levels (pH 2.0–3.0).²^,³^,⁴^,⁵^,⁶^,⁷ The fact that insect species-specific gut pH is among the critical internal factors that significantly influence digestion has been known for a long time.⁸

Among holometabolous insects, mosquitoes have remained at the center stage of vector research, owing to their significance in public health. Mosquitoes are most vulnerable during their larval stages. Therefore, a great deal of research effort to develop vector-control strategies is focused on these stages. Earlier researchers have conducted extensive studies on the larval stages, revealing several intriguing anatomical and physiological characteristics, especially in the gut.⁹^,¹⁰^,¹¹ The larval gut is a simple structure that lacks morphological barriers, yet it displays a unique pH gradient profile in the midgut.¹⁰^,¹² The midgut includes a neutral to alkaline (pH∼7–8) gastric cecum followed by a hyperalkaline (pH∼10–11) anterior midgut and a near-neutral posterior midgut (Figures 1A and 1B). This gastric arrangement is remarkably contradictory to other metazoans, where optimal digestion initiates in an acidic to highly acidic environment.

The gut pH profile in mosquito larvae

(A) Gut pH profile in monoculture (*Bacillus cereus*) associated with *Aedes albopictus* gnotobiotic larva.

(B) Gut pH profile in a conventionally reared *Ae. albopictus* larva (harboring complex microbiota). Larvae were allowed to ingest a pH indicator dye (0.01% Phenol Red) for 15 min and subsequently visualized under a stereomicroscope. Gastric caeca (GC, corresponding to the thorax segment), anterior midgut (AMG, corresponding to the first 3–4 abdominal segments), and posterior midgut (PMG, corresponding to abdominal segments following AMG) show distinct colors—red (∼pH 8), magenta (∼pH 10–11), and orange/yellow (∼pH 7–7.5), respectively—indicating the prevailing pH conditions in the corresponding anatomical compartments. Scale bars on the micrographs represent 200 μm in length.

The hyperalkaline midgut microenvironment in mosquito larvae is believed to play a crucial role in dissociation, coagulation, digestion of ingested protein complexes, detritus, neutralization of toxic materials, and control of pathogens, all of which contribute toward larval survival, growth, and development.²^,⁸^,¹⁰^,¹³ The hyperalkaline midgut is believed to be a distinctive and evolutionarily conserved taxonomical feature that has probably evolved from the adaptation of the ancestral insect to a specific diet.⁸ A group of researchers has recently proposed the use of triazabutadienes as target-specific mosquito larvicides that become active when exposed to the highly alkaline environment of the larval midgut.¹⁴ Development and successful application of such highly specific larvicidal strategies require a detailed understanding of the factors that modulate this unique ionic microenvironment in the mosquito larval gut.

It is also well known that mosquito larvae harbor diverse microbiota within their gut, which essentially modulates a wide variety of phenotypes associated with the evolutionary success of the host.⁴^,¹⁵^,¹⁶ Extensive review of existing scientific literature suggests that gut microbiota performs at least two core functions in the larvae, namely nutrition and defense.³^,¹⁷ Although an imperative and positive role of gut bacteria in larval survival, growth, and development is principally accepted, the exclusive requirement of “live” bacteria for the implementation of such functions is seriously debated.⁴^,¹⁸^,¹⁹^,²⁰^,²¹^,²²^,²³ While some researchers argue that “live” bacteria are essential for larval growth and development, others challenge this proposition.²⁰^,²²^,²⁴^,²⁵^,²⁶^,²⁷

Initially, bacterial respiration-mediated hypoxia in the larval gut was suggested as the crucial molecular mechanism that triggers larval development.²² This mechanism was immediately challenged by a subsequent study, in which researchers reared axenic larvae (bacteria-free) on a high-concentration diet and proposed that the association between a larva and its microbiota is principally nutritional over symbiotic.²⁰ Despite larval growth and development, a significant delay in the development of axenic larvae was evident in this study.²⁰ Concurrently, another study found that axenic larvae could not survive beyond the first instar stage, even when fed with a nutritionally complete diet.²⁵ These researchers also found that expression of genes associated with protein digestion, amino acid transport, hormone signaling, metabolism, etc., are significantly disrupted in the absence of gut bacteria.²⁵ Later, Romoli and colleagues demonstrated that absence of gut microbiota significantly alters the expression of key genes involved in protein and lipid metabolism, amino acid storage and protease inhibitors.²⁴

Further, based on upregulation of genes associated with the folate (vitamin B9) pathway in decolonized larvae, researchers proposed an important role of gut microbiota in the contribution of folate.²⁴ Contemporaneously, Wang and colleagues showed that gut microbiota triggers larval development through contribution of riboflavin (vitamin B2),²⁸ rather than by microbiota-associated hypoxia as proposed earlier.²² However, a more recent study demonstrated that dietary supplementation with a cocktail of vitamin B (including B1, B2, B3, B6, B7, B9, and choline) alone does not significantly improve the development of bacteria-free larvae.²⁹ These findings suggest that larval development not only depends on bacterial vitamin B but also on other essential contributions that are exclusive to live bacteria. Recently, Raquin and colleagues showed that larval survival is principally a trade-off between diet concentration and bacterial inoculum size and is probably driven by specific taxa of bacteria.³⁰ Nevertheless, the imperative role of microbiota in various aspects of the mosquito life cycle is quite evident in relevant literature.³^,³¹

It is now acknowledged that larval digestion, growth, and development are tightly regulated through an evolutionarily conserved mechanism involving specific members of the gut microbiota.⁴^,²¹^,³⁰^,³² Further, growing evidence suggests that the effects of gut microbiota on larval stages are also carried over to the adult stages, having consequences on reproduction, fecundity, vector competence, etc.¹⁵^,¹⁶^,³³^,³⁴^,³⁵^,³⁶ Therefore, a better understanding of host-microbe interaction mechanisms would allow exploration of controlled gut microbiota manipulation as a potential strategy for vector control.

Incidentally, while studying the effects of bacteriophage-mediated manipulation of gut bacteria (Bacillus cereus) in Aedes albopictus gnotobiotic larvae, we observed that the characteristic gut pH profile was lost in a significantly higher percentage of larvae (p < 0.0001) after treatment with a lytic bacteriophage infecting B. cereus (Figures 2A–2C). Further, the growth rate of the phage-treated larvae was also significantly reduced (p < 0.001) as compared to the mock-treated gnotobiotic larvae (Figure 2D). These observations led us to hypothesize that the colonizing bacteria influence gut physiology by modulating the ionic microenvironment, which in turn affects larval growth. Subsequently, we evaluated this hypothesis using the Ae. albopictus axenic-gnotobiotic larval model and pursued the possible mechanism involved. Our results show that midgut alkalinization and optimal protease activity in larval gut are modulated by the colonizing bacteria in a taxa-specific manner. In this manuscript, we report our findings and discuss their significance in light of available literature.

Gut pH profile in gnotobiotic larvae is abolished after treatment with a lytic bacteriophage

(A) Gut pH profile in mock (no phage) treated gnotobiotic larvae (*Bacillus cereus*).

(B) Disrupted gut pH profile in lytic bacteriophage (Bacillus phage BCE1)-treated gnotobiotic larvae (*Bacillus cereus*). The photomicrograph shows loss of alkalinity in the gastric caeca and anterior regions of the midgut.

(C) The bar graph represents a significant reduction in the percentage of larvae displaying the typical pH profile in the bacteriophage-treated experimental group (pale lime green bar) as compared to the untreated gnotobiotic group (light magenta bar). Colored bars show the mean (of data obtained from three independent experiments) for the corresponding experimental group, and error bars represent the S.D. ∗∗∗∗p < 0.0001 by unpaired two-tailed t test.

(D) Violin plot showing significant differences in the length of bacteriophage-treated larvae (pale lime green violin) as compared to the untreated gnotobiotic larvae (light magenta violin). Length data were collected 2 days post-treatment. Violin plots show the distribution of data points with median and quartiles. ∗∗∗p < 0.001 by unpaired nonparametric two-tailed Mann-Whitney U test. Scale bars on the micrographs represent 200 μm in length.

Results

Concurrent presence of bacteria and carbonic anhydrase activity is required for the establishment of a distinctive pH profile in the larval gut

Compared to the presence of distinctive gut pH profiles in the gnotobiotic larval group, pH profiles were not visible in any of the axenic larvae (Figures 3A and 3B). Additionally, the percentage of larvae showing pH profile was significantly lower in antibiotic (gentamicin)-treated (p < 0.0001) and carbonic anhydrase inhibitor (acetazolamide)-treated (p < 0.0001) gnotobiotic larvae (Figures 3C–3E). The loss of gut pH profile in groups of gnotobiotic larvae separately treated with an antibiotic and a carbonic anhydrase inhibitor suggested that gut-associated bacteria, along with carbonic anhydrase activity, are required for midgut alkalinization.

The continual presence of live bacteria is required for the maintenance of a distinctive pH profile in the larval gut

When gnotobiotic larvae displaying a well-established gut pH profile were subjected to antibiotic treatment, the distinctive pH profile was abolished in a significantly greater percentage of larvae (p < 0.001) (Figures 4A–4C). This observation suggested that the establishment and maintenance of a gut pH profile by gut bacteria is not a typical “hit and run” phenomenon; rather, it requires the continual presence of bacteria.

The continual presence of live bacteria is required for the maintenance of a distinctive pH profile

(A) Gut pH profiles in gnotobiotic larvae before antibiotic treatment.

(B) Loss of gut pH profile after treatment with antibiotic (gentamicin, 20 μg mL⁻¹ concentration).

(C) The bar graph represents a significant reduction in the percentage of larvae displaying the typical pH profile after treatment with antibiotic (gentamicin, light orange bar), as compared to the gnotobiotic larval group before antibiotic treatment (light magenta bar). Colored bars show the mean (of data obtained from three independent experiments) for the corresponding experimental group, and error bars represent the S.D. ∗∗∗p < 0.001 by unpaired two-tailed t test. Scale bars on the micrographs represent 200 μm in length.

Only live bacteria can establish and maintain larval gut pH, and is associated with larval growth

Our experiments on axenic larvae reared with live bacteria, cell-free bacterial culture supernatant, and bacterial cell lysates demonstrated that the distinctive pH profile is established only when reared with live bacteria (Figures S1A–S1D). A gut pH profile was observed neither in larvae reared with bacterial culture supernatant nor in larvae cultured with bacterial cell lysate (Figures S1C and S1D). Further, the growth of larvae reared with live bacteria having a distinctive pH profile was significantly greater (p < 0.0001) than the growth of larvae reared under axenic conditions or with bacterial cell lysate or bacterial culture supernatant (Figure S1E).

Only selected members of the gut microbiota can establish a distinctive gut pH profile, modulate proteolytic activity, and promote larval growth

We observed that among the members of the gut microbiota included in the present study, bacteria from the families Bacillaceae (B. cereus and Lysinibacillus fusiformis) and Aeromonadaceae (Aeromonas hydrophila) could establish the gut pH profile and promote larval growth, while bacteria from the family Microbacteriaceae (Microbacterium paraoxydans and Leucobacter coleopterorum) could not (Figures 5A–5F). The lengths of gnotobiotic larvae reared with B. cereus, L. fusiformis, or A. hydrophila were significantly greater (p < 0.0001) as compared to axenic larvae (Figure 5G). On the other hand, no significant difference was observed in the lengths of gnotobiotic larvae reared with either M. paraoxydans (p = 0.3775) or L. coleopterorum (p = 0.8482) as compared with the axenic larvae (Figure 5G).

Detection of proteolytic activities in larval lysates from diverse groups of gnotobiotic larvae corroborate with the presence of a distinctive gut pH profile and larval growth (Figure 5H). Markedly, gnotobiotic larvae colonized with B. cereus and L. fusiformis showed higher levels of proteolytic activity and growth as compared with larvae colonized with A. hydrophila (Figures 5G and 5H). No proteolytic activity was observed in lysates from axenic larvae or gnotobiotic larvae colonized with M. paraoxydans or L. coleopterorum (Figure 5H).

Furthermore, when each of these five bacteria was grown individually on casein agar plates, B. cereus, L. fusiformis, and A. hydrophila showed zones of hydrolysis, representing extracellular protease activity (Figure 5I). No extracellular proteolytic activity was observed on casein plates inoculated with M. paraoxydans (Figure 5I). On the other hand, L. coleopterorum did not even grow on the casein plates (plate not shown).

Live bacteria colonize the larval gut rapidly and are localized within the anterior midgut and gastric caeca regions

We conducted a live bacteria localization experiment (using live bacteria labeling fluorescent dye) to explore the causal link between bacterial colonization and the hyperalkaline microenvironment in the larval anterior midgut. For the bacterial localization study, we selected B. cereus as the representative growth-promoting bacteria. Remarkably, fluorescence microscope images showed intense signals originating from the labeled bacteria that could be detected almost immediately (within 5 min of feeding labeled bacteria to the axenic larvae). The fluorescent signal was observed in the regions corresponding to the gastric caeca in the thorax segment and the anterior midgut in the first 3–4 abdominal segments (Figure 6A). Even though the fluorescent signal in the gastric caeca was initially high, it declined noticeably over the period of observation. On the other hand, fluorescent signals remained intense and strictly localized to the anterior midgut throughout the observation period (60 min).

On the other hand, in a localization experiment conducted with a representative non-growth-promoting bacteria colonized (M. paraoxydans) gnotobiotic larvae, we could localize fluorescent signals in the anterior midgut region after 60 min, similar to the localization observed in B. cereus gnotobiotic larvae (Figures 6B and 6C). However, the intensity of the fluorescent signal was markedly different among the Bacillus and Microbacterium gnotobiotic larvae (Figure S2). No fluorescent signal was detected in the mock-labelled axenic larvae (Figure 6D).

Bacteria colonizing the larval anterior midgut contribute different proteases and induce the larva to secrete host proteases

We conducted experiments to determine the nature and source (bacterial or larval) of proteases present in the gnotobiotic larvae, adopting a subtractive approach (a schematic diagram is presented as Figure 7A). In this approach, proteases present in the untreated gnotobiotic larvae (considered to be a combination of larval- and bacterial-origin proteases) were compared with proteases present in the bacteriophage-treated larvae (considering the removal of colonizing bacteria and thus its proteases, leaving only proteases of larval origin). The difference between the protease profiles in these two experimental groups was considered to represent the proteases contributed by the colonizing bacteria. In this experiment, axenic and gnotobiotic larvae (B. cereus) were generated as described in the previous section. A subset of the gnotobiotic larvae was then separately treated with a highly lytic bacteriophage, isolated against the B. cereus strain used in the present study. In parallel, a subset of gnotobiotic larvae treated with a standard antibiotic (gentamicin) was included as control to compare the decolonizing potential of the bacteriophage. Results of bacteriophage-mediated decolonization of gnotobiotic larvae were verified by plate counting and real-time PCR detection of bacterial DNA in larval lysates, which demonstrated the effectiveness of bacteriophage in decolonizing bacteria, comparable to the standard antibiotic treatment (Figures 7B and 7C).

Results of the total protease activity assay revealed practically no detectable activity in the axenic larvae, while a significantly high level of protease activity was present in the gnotobiotic larvae (p < 0.0001) (Figure 7D). In contrast, a significant decrease (p < 0.01) in protease activity (nearly half as compared to gnotobiotic larvae) was evident in both bacteriophage-treated and antibiotic-treated larvae (Figure 7D). Subsequent results of the protease inhibition assay demonstrated the predominance of serine protease activity followed by cysteine protease and aminopeptidase activities in the untreated gnotobiotic larvae (Figure 7E). On the other hand, in phage-treated gnotobiotic larvae (representing larval-origin proteases), moderate levels of cysteine protease, serine protease, and aminopeptidase activities were observed. A significant difference (p < 0.0001) was observed in serine protease and aminopeptidase activities between gnotobiotic and phage-treated larval groups, indicating the bacterial origin of these proteases. Further, almost all the metalloprotease activity and the bulk of the Ca²⁺-dependent protease activities detected in gnotobiotic larvae were also mapped to be of probable bacterial origin (p < 0.0001).

Subsequently, the nature of proteases secreted by the colonizing bacterial species (B. cereus) in casein media (pH 8.0) was evaluated. Results of these experiments showed a predominance of aminopeptidase and serine protease activities in the bacterial supernatant, while minor amounts of cysteine and aspartyl protease activities were also detected (Figure 7F). Of the total protease activity, ∼86% of the activity was found to be associated with metal ion-activated proteases (EDTA inhibitable), and an increase (∼16%) in total protease activity in the presence of CaCl₂ suggested the presence of Ca²⁺-dependent protease activity (Figure 7F).

Additionally, we cultured the colonizing bacterial species (B. cereus) in casein media (pH 10.0) to imitate the pH observed in the larval anterior midgut. Apart from a slight increase in activity of some of the proteases at pH 10, we could not see any significant changes in other proteases secreted by the bacteria (Figure S3).

Supplementary diet alone has no significant effect on larval anterior midgut alkalinization, protease activity, and growth

We conducted experiments to examine if a proteinaceous supplementary diet has any role in larval anterior midgut alkalinization, protease activity, and growth. In both the axenic larval groups (with and without supplementary diet), midgut alkalinization and protease activity was not observed (Figures 8A–8E). Growth was also significantly lower in both the axenic larval groups as compared to the gnotobiotic larval groups (Figure 8F). On the other hand, both the gnotobiotic larval groups (with and without supplementary diet) showed comparable growth, protease activity, and anterior midgut alkalinization (Figures 8E–8G). These results indicate that supplementary diet itself has no significant effects on the tested parameters that influence larval growth.

Discussion

Host-microbe relationships in mosquitoes have been investigated from various perspectives. However, microbiological aspects associated with midgut alkalinization and its impact on larval physiology have remained understudied. To the best of our knowledge, only one study demonstrated the effects of a commensal bacterium (Asaia bogorensis) associated with midgut alkalinization on pathogen proliferation in adult Anopheles stephensi mosquitoes.³⁷ We could not find any other study that focused on exploring the role of gut microbiota in larval physiology, especially midgut alkalinization and digestive activity. In this context, results of our present study reveal interesting facets of the larval host-microbe association.

Results of our studies on gut bacteria localization show that although both B. cereus and M. paraoxydans can colonize the larval midgut, only B. cereus induces anterior midgut alkalinization. Interestingly, a difference in fluorescent signal intensity was observed between the Bacillus and Microbacterium gnotobiotic larvae. We assume that this difference might be attributable to the relatively higher packing density of Microbacterium due to its smaller size, as compared with Bacillus. Further, we observed that bacteria rapidly colonize the larval midgut, localize, and remain restricted to the anterior midgut in high abundance and in low abundance in the gastric caeca. This relative abundance of colonizing bacteria apparently coincides with the levels of alkalinity detected in these two anatomical sites. Further, highly abundant localization of the colonizing bacteria in the anterior midgut precisely matches with a previous study conducted on Ae. aegypti larvae colonized with E. coli (live GFP⁺), suggesting that bacterial colonization sites are conserved among these mosquito species.²²^,²³ These findings also signify the presence of broad-spectrum bacterial attachment mechanisms in the larval midgut, to which bacteria from various genera can attach. In our experiments, we could isolate live bacteria from the gnotobiotic larvae even days after colonization. This contrasts with the earlier study, which showed that viability of colonizing GFP⁺ E. coli (not a natural member of the larval gut microbiota) markedly declined within 240 min.²² The researchers attributed this observation to the inability of E. coli to withstand highly alkaline midgut conditions. To us, these observations probably indicate a larval mechanism to selectively retain bacterial species that are natural members of the gut microbiota. Frequent detection of bacteria belonging to certain genera, irrespective of the geographical location or microbes present in the respective breeding environment, seems to support such an assumption.

Our experiments on gnotobiotic larval groups colonized with species belonging to the bacterial genera Bacillus, Lysinibacillus, or Aeromonas revealed co-occurrence of larval midgut alkalinization, protease activity, and growth, suggesting a close association between these events. In contrast, none of these events were evident in gnotobiotic larvae colonized with members of bacterial genera Microbacterium or Leucobacter. These findings agree with an earlier study on lepidopteran larvae, showing that among the diverse bacteria present in the gut, selected extracellular protease-producing gut bacteria contribute proteolytic activity in the larval gut.³⁸ Our observations exactly match with previous studies showing larval growth and development in gnotobiotic larvae colonized with Aeromonas, but not with Microbacterium.²³^,³² Overall, these observations provide a fresh insight into the bacterial species-specific contribution to larval digestion and growth.

On the other hand, the present results also reconcile with earlier electrophysiological and biochemical studies, reiterating the significance of carbonic anhydrase-mediated ionic homeostatic mechanisms in midgut alkalinization and, in turn, survival and growth of larvae.³⁹^,⁴⁰^,⁴¹ Conventionally, colonization or decolonization of microbiota is believed to be a function of larval gut pH.⁴^,²²^,⁴² Such a notion would then require exclusive involvement of “larval” carbonic anhydrase activity in midgut alkalinization. However, previous studies have shown that the larval carbonic anhydrase expression patterns (remarkably high expression in gastric caeca/posterior midgut and exceptionally low expression in anterior midgut) do not corroborate with the gut pH profile.⁴⁰^,⁴³^,⁴⁴^,⁴⁵^,⁴⁶ Subsequently, a “soluble extracellular carbonic anhydrase” located in the ectoperitrophic space was proposed as the principal factor involved in anterior midgut alkalinization.⁴⁰ During the literature review, we found that the earlier observations of high luminal pH in live, intact, and conventionally reared larvae and the dramatic loss of gut pH in dissected, in vitro-bathed, and buffer-perfused gut preparations seem to suggest a bacterial origin of the proposed “soluble extracellular carbonic anhydrase.”⁴⁰^,⁴³^,⁴⁷ Detection of α and β classes of carbonic anhydrases (also encoded by certain prokaryotes) in the ectoperitrophic fluid and gut lumen of larvae and sturdy expression of carbonic anhydrase-like proteins by certain members of the larval gut microbiota (e.g., Pseudomonas, Serratia, and Proteus) appear to implicate bacterial carbonic anhydrases in midgut alkalinization.³⁶^,⁴⁴^,⁴⁵^,⁴⁶^,⁴⁸^,⁴⁹^,⁵⁰^,⁵¹^,⁵²

Dadd earlier demonstrated that mosquito larvae display similar gut pH profiles irrespective of the mosquito genera, instar stage, or diet status and that the hyperalkaline microenvironment of anterior midgut is crucial for optimal digestion of dietary proteins.¹⁰ Among the other proteases, dominance of alkaline serine proteases in insect larval gut is well documented.⁸^,¹⁰^,⁵³^,⁵⁴^,⁵⁵ Though the possibility of bacterial origin of insect gut proteases was suggested long back, most of the evidence available in literature is indirect.⁸^,⁵³ Strong interference in protein degradation capabilities, survival, growth, and development after treatment with protease inhibitors, bactericidal agents, or antibiotics has been demonstrated in larval and adult stages of several insects, including mosquitoes.³⁸^,⁵⁶^,⁵⁷^,⁵⁸^,⁵⁹^,⁶⁰^,⁶¹^,⁶²^,⁶³ Furthermore, a drastic decline in proteolytic activity during or after the larval-pupal ecdysis phase, exactly coinciding with major turnover or complete removal of the gut microbiota in holometabolous insects, designates the role of gut bacteria in larval gut protease activities.⁵⁵^,⁶¹^,⁶³^,⁶⁴

In the present study, utilizing the phage-mediated gut bacteria decolonization approach, we quantitatively and qualitatively analyzed various proteases of bacterial origin in the larvae. Recently, Tikhe and Dimopoulos demonstrated successful phage therapy in mosquito larvae (Anopheles gambiae) as a biocontrol approach.⁶⁵ We preferred bacteriophage over antibiotic treatment to avoid potential confounding effects (e.g., modulation of eukaryotic gene expression) associated with the antibiotics.⁶⁶^,⁶⁷ Our detection of diverse proteases in larval gut is in accordance with previous studies on several insect species, including those having hyperalkaline midgut (e.g., Lepidopteran larvae).⁶^,⁶⁸^,⁶⁹^,⁷⁰^,⁷¹^,⁷² In addition, our results show that the colonizing bacteria contributes a significant proportion of important proteases to the larvae, which corroborates with the secretory protease profile of the colonizing bacteria.⁷³^,⁷⁴^,⁷⁵^,⁷⁶

In addition, significant differences in protease activity between axenic larvae (never colonized) and decolonized larvae indicate that exposure to gut bacteria is crucial for expression of larval proteases. These observations corroborate with two earlier transcriptome studies (conducted on E. coli-colonized Ae. aegypti larvae) showing significant modulation in expression of genes encoding proteins that are crucial for protein metabolism.²⁴^,²⁵ In one of these studies, significant downregulation of the serine protease gene along with upregulation of the serine protease inhibitor gene (serpin) was observed in decolonized larvae, signifying specific suppression of serine protease activity through multiple mechanisms in the absence of gut bacteria.²⁴^,⁷⁷

Interestingly, in certain insects (belonging to the order Lepidoptera), among the different gut proteases, only serine protease activity correlated with digestive competence in larvae.⁸ Studies have also shown that depletion in gut proteases by various means causes serious disturbances in digestion, nutrient absorption, and amino acid bioavailability, which adversely affect survival, growth, and development, particularly during larval stages.⁷⁸^,⁷⁹^,⁸⁰^,⁸¹^,⁸² Even though insects are known to upregulate protease expression in response to loss of gut protease activity, such adaptive mechanisms are inadequate to counter substantial losses.⁷⁹ Apparently, a similar phenomenon was observed in decolonized Ae. aegypti larvae, where loss of essential endopeptidase activity (e.g., serine protease) overlapped with upregulation of exopeptidase activity (e.g., carboxypeptidase), yet survival, growth, or development could not be rescued.²⁴

In the present study, we observed that among the different proteases, serine protease and aminopeptidase activities are significantly reduced after removal of the colonizing bacteria. These two protease classes represent basic proteases, omnipresent across the animal kingdom and engage in various functions, including the digestion of food.⁸³ Serine proteases are endopeptidases that digest complex dietary proteins into simpler peptides. On the other hand, aminopeptidases are exopeptidases that cleave proteins and peptides at their amino ends and release amino acids. In living systems, different classes of proteases act on the ingested complex dietary proteins and release amino acids, which are subsequently absorbed and used for various cellular and physiological functions.⁸³ In this context, our observation of delayed growth in decolonized larvae suggests that host-associated proteases alone are not sufficient to support normal larval growth and implies the conserved nature of gut bacteria-assisted protein digestion mechanism.

Although we cannot conclude from the present data, our observations along with results from previous studies tempt us to assume gut bacteria-mediated midgut alkalinization is the chemical signal that modulates expression of certain host genes, especially those related to digestion and nutrient assimilation. This proposal draws substantial support from the results of the previous study, where expression of high pH optima enzymes, especially serine proteases and alkaline phosphatases, is significantly modulated depending upon the presence or absence of the gut bacteria.²⁴ In different insect larvae (having a high gut pH), pH optima of various proteases (serine proteases, aminopeptidase, and metalloproteases) have been found to be high, as compared to other enzymes. In addition, signaling through conserved bacterial products or modulation of host microRNAs may be involved, as proposed earlier.²³^,⁸⁴

It is worth mentioning here that the results discussed in the foregoing text were based on experiments performed without adding any supplementary diet. Since the present study was focused exclusively on bacterial colonization-mediated changes in larval physiology, exclusion of diet helped to ensure that the effects were attributed solely to the colonizing bacteria under investigation. Previous researchers have shown that different microorganisms serve as food source, from which larvae can acquire nutrients essential for survival, growth, and development.³⁰^,⁸⁵ Even though the results so far indicated a dominant role of colonizing bacteria, we also examined whether diet supplementation alone has any effect on larval physiology. Our results indicated that supplementary diet (or proteins present therein) has neither any significant effect on midgut alkalinization nor in protease expression. Interestingly, a recent study demonstrated that survival, growth, and development of mosquito larvae (Anopheles gambiae) depend solely on the species of the colonizing bacteria, irrespective of the diet.⁶⁵ While supporting these earlier findings, our results provide a mechanistic explanation for the observed bacterial species-specific differences in larval growth and development, regardless of diet supplementation.

Altogether, our findings propose an interesting model of host-microbe interaction, where gut bacteria influence larval growth and development by modulating larval gut physiology. In this model, specific members of the gut microbiota facilitate larval digestion and nutrition through establishing a unique ionic microenvironment in the midgut, which is imperative for optimal activity of the digestive enzymes. These bacteria also contribute various proteases, in addition to inducing the expression of larval proteases, thereby ensuring maximal protein digestion in the larval gut. Apart from assimilating previous observations of bacterial taxa-specific influences on larval physiology, this model also explains the wide-ranging carryover phenotypic effects seen in adults emerging from larvae reared with sub-optimal microbiota. Additionally, it also accommodates the necessity of bacterial B vitamins (e.g., folate and riboflavin) as essential cofactors involved in various metabolic pathways (including amino acid metabolism). Nevertheless, we believe that the gut microbiota also controls other important metabolic pathways in the larvae, which remain to be explored.

Conclusion

A better understanding of the host-microbe interaction in mosquitoes is believed to reveal newer targets for controlling mosquito-borne diseases. Currently, gut microbiota modulation is actively being pursued as a potential eco-friendly strategy to combat mosquito-borne diseases, as compared to the conventional chemical-based control methods. In this direction, our study reveals an important functional mechanism of the host-microbe association, which can be targeted through various approaches, including the use of lytic bacteriophages against growth-promoting gut bacteria, the use of natural larval gut pH modulators, protease inhibitors, and alkaline pH-activated insecticides. We believe that our findings will help in better understanding the role of gut microbiota in vector biology and in developing efficient and eco-friendly vector-control strategies.

Limitations of the study

While our results signify that certain bacterial species are associated with anterior midgut hyperalkalinization and protease activity, we could not explore the underlying mechanisms in the present study. Future studies involving colonization of larvae with mutant bacterial strains that are unable to express carbonic anhydrase/proteases may provide insights into the mechanism of this host-microbe association. Additionally, studies focused on the expression of proteases specific to different bacterial genera/species could reveal more interesting facts.

Resource availability

Lead contact

Further information on resources, reagents, protocols, etc., should be directed to and will be fulfilled by the lead contact, Sibnarayan Datta (sndatta.drl@gov.in).

Materials availability

This study did not generate new unique reagents.

Data and code availability

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All the data generated in this study are presented within the main/supplementary figures and tables. Bacillus phage BCE1 genome and bacterial 16S rRNA gene sequences have been deposited in GenBank. GenBank accession numbers are provided in the key resources table against the corresponding item.
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No custom code was generated or used in this study.
•
Any supplementary information essential to reanalyze the data presented in this paper is available from the lead contact upon request.

Acknowledgments

This work was funded by intramural grants and a research fellowship (JS) from the Defence Research & Development Organization (DRDO). We thank Md. Kawish, Dr. Bipul Rabha, Dr. Diganta Goswami, and Dr. Rajan Pilakandy for help with larval culture maintenance and handling. We thank Dr. Ratan Baruah (SAIC, Tezpur University) for help with TEM imaging. We thank Dr. Ottavia Romoli of Institut Pasteur (Paris, France) for useful suggestions and help with the transcriptomic data (VectorBase). We thank Dr. Vanlalhmuaka and Dr. Dev Vrat Kamboj for helpful discussions. We thank Ms. Dolly Samanta for the original hand-drawn art (graphical summary), which was converted into a digital version using the BioRender tool. We express our sincere gratitude to the reviewers for their constructive comments and guidance toward improving the work.

This manuscript is published under permission from the Director, Defence Research Laboratory, Tezpur (manuscript reference number DRL/EBM/01/2024).

Author contributions

J.S. and S.D. conceptualized the study; J.S. performed experiments; J.S. and S.D. conducted formal analysis; and J.S. and S.D. wrote the original draft and reviewed and edited the manuscript.

Declaration of interests

The authors declare no competing interests.

STAR★Methods

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Bacterial and virus strains

Aeromonas hydrophila	This study	N/A
Bacillus cereus	This study	N/A
Leucobacter coleopterorum	This study	N/A
Lysinibacillus fusiformis	This study	N/A
Microbacterium paraoxydans	This study	N/A
Bacillus phage BCE1 (bacteriophage)	This study	N/A

Chemicals, peptides, and recombinant proteins

Acetazolamide	Sigma-Aldrich	A6011
AEBSF	Sigma-Aldrich	A8556
Azocasein	Sigma-Aldrich	A2765
Bestatin	Sigma-Aldrich	B8385
Calcium chloride	Merck	10238005001730
Casein	HiMedia	GRM087
Cell lysis/extraction reagent	Sigma-Aldrich	C 2360
Ethanol	Merck	100983
EDTA	Sigma-Aldrich	E5134
E64	Sigma-Aldrich	E3132
Gentamicin	Amresco	E737
Glycine	Sigma-Aldrich	G8898
Leupeptin	Sigma-Aldrich	L9783
Luria-Bertani agar	Himedia	G1151
Luria-Bertani broth	Himedia	M1245
Pepstatin A	Sigma-Aldrich	P5318
Phenol Red	Sigma-Aldrich	P3532
Phosphate-buffered saline	Sigma-Aldrich	P3813
Protease inhibitor cocktail (General use)	Amresco	M221
Protease inhibitor cocktail (for use with bacterial cell extracts)	Sigma	P8465
Sodium hydroxide	Merck	61843805001730
Sodium hypochlorite solution	SRL	25366
Tetramin Baby micro fish flakes	Tetra	NA
Trichloroacetic Acid	BioString	T2034

Critical commercial assays

Bradford assay kit	Sigma-Aldrich	B6916
BactoView™ Live Green Fluorescent Bacterial Stain	Biotium	40102
PCR Mastermix	Roche	11636103001
Qiagen tissue DNA isolation kit	Qiagen	51404
qPCR mastermix (SYBR Green)	Sigma-Aldrich	S4438

Deposited data

Aeromonas hydrophila	NCBI GenBank	GenBank: OR513036
Bacillus cereus	NCBI GenBank	GenBank: OR513030
Leucobacter coleopterorum	NCBI GenBank	GenBank: OR513031
Lysinibacillus fusiformis	NCBI GenBank	GenBank: OR513032
Microbacterium paraoxydans	NCBI GenBank	GenBank: OR513035
Bacillus phage BCE1	NCBI GenBank	GenBank: PQ655445

Experimental models: Organisms/strains

Aedes albopictus	This study	N/A

Oligonucleotides

Act-2F (5′-atggtcggyatgggncagaaggactc-3′)	Eurofins Genomics	1427455
Act-8R (5′-gattccatacccaggaaggadgg-3′)	Eurofins Genomics	1427456
Bac-Rev (5′-acatctcacgacacgagctgacgac-3′)	Eurofins Genomics	2094624
BsF2 (5′-tagggaagaacaagtgctagttgaataa-3′)	Eurofins Genomics	2094616
fD1 (5′-agagtttgatcctggctcag-3′)	Xcelris Labs	152189-1/97
rP2 (5′-acggctaccttgttacgactt-3′)	Xcelris Labs	152189-2/97
usff (5′-attggagggcaagtctggtg-3′)	Bioserve Biotech	137212
usfr (5′-ccgatccctagtcggcatag-3′)	Bioserve Biotech	137213

Software and algorithms

CellSens v2.1	Olympus	CS-ST-V2 & CS-D1-V3
CFX Maestro 1.0 v4.0	Bio-Rad	N/A
Spreadsheet (Excel)	Microsoft	version 2501
ImageJ v1.54g	ImageJ	https://imagej.net/ij/
GraphPad Prism v8.4.2	GraphPad	www.graphpad.com

Other

Biosafety cabinet (Class II A2)	Thermo Fisher Scientific	1386
Bacteriological Incubator	Thermo Fisher Scientific	SHKE6000-8CE
Thermal cycler	Bio-Rad	C1000
Environmental chamber	Creative Lab World	CLW144
Fluorescent microscope	Olympus	BX43
Gel documentation system	UVP	BioSpectrum415
High-speed Centrifuge	Hermle	Z36HK
qPCR	Bio-Rad	CFX96
Spectrophotometer (UV-Vis)	Thermo Fisher Scientific	Evolution Pro
Stereomicroscope	Olympus	SZX16
Syringe filter (0.2 μm)	Pall Corporation	4612
Tissue culture plate (6-well)	Tarsons	980010
Tissue homogenizer	Genetix	Ez-Lyzer

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Experimental model and study participant details

Experimental model

Larvae emerging from eggs laid by laboratory-reared Aedes albopictus were used as an experimental model in this study. Experimental larval groups were maintained inside an environmental chamber, set at 27±2°C, 80±10% relative humidity, and a 12 h light/dark cycle.

Bacterial strains and growth conditions

All the bacterial strains used in experiments described in this manuscript (A. hydrophila, B. cereus, L. coleopterorum, L. fusiformis, and M. paraoxydans) were isolated from laboratory-reared Ae. albopictus larvae during this study. Bacterial isolates were cultured in/on Luria-Bertani (LB) broth/agar plates and incubated at 37°C. All microbiological manipulations were performed inside BSL2 using sterile plasticware. Standard microbiological laboratory procedures and precautions were followed strictly. The study protocol was approved by the Institutional Biosafety Committee (DRL/IBSC/PROJ/02).

Virus strain

A lytic bacteriophage (BCE1) infecting B. cereus was isolated from a soil sample during this study. This bacteriophage was used in decolonization experiments conducted with B. cereus-associated gnotobiotic larvae.

Method details

Generation of axenic larvae

Axenic and gnotobiotic larvae were generated from surface-sterilized Ae. albopictus eggs following a previously published protocol,⁸⁶ with minor modifications. For the generation of axenic larvae, eggs were at first immersed in 70% ethanol for 5 min, followed by washing with sterile water. Subsequently, eggs were submerged in 1% bleach solution for 5 min, followed by washing with sterile water and then retreated with 70% ethanol for 5 min. Finally, sterilized eggs were rinsed thrice with sterile water. Sterilized eggs were then added to sterile LB broth and allowed to hatch inside an environmental chamber, set at 27±2°C, 80±10% relative humidity, and a 12 h light/dark cycle. Sterilization was conducted in petri plates, and the plates were gently swirled during the incubation and washing steps to allow maximum exposure of the eggs to each liquid. Wide-bore P1000 pipette tips were used for handling and transfer of eggs into different liquids. The complete sterilization process was performed inside a biosafety cabinet (BSL2), using sterile plasticware.

Generation of gnotobiotic larvae

For the generation of gnotobiotic larvae, a subset of axenic larvae was collected from the hatching media and washed thoroughly with sterile water. Ten larvae were added to each well of 6-well culture plates holding 5 mL sterile water. For initial experiments, B. cereus was used for larval gut colonization. This bacterial species is reported to stably and harmlessly colonize the mosquito larval gut for longer periods without interfering with their growth and development.⁸⁷ Bacterial colonization was conducted by adding culture to a final dose of ∼10⁶ CFU mL^-1 in each of the plate wells containing axenic larvae and kept in an environmental chamber. After 3 days larvae were collected, washed thoroughly with sterile water, and transferred to a fresh plate of wells holding sterile water.

Microbiological assessment of axenic and gnotobiotic larvae

To assess the success of axenic/gnotobiotic larvae generation procedures, approximately 40-50 washed larvae from both groups were sampled separately into sterile 1.5 mL microcentrifuge tubes, and excess water was removed by pipetting. Larvae were euthanized by chilling the tubes on ice for 1 h. Larval lysate was then prepared by adding 250 μL of 1x PBS buffer solution and 2.0 mm zirconia beads to the tubes, followed by homogenization (Ez-lyzer) at 25 Hz for 3 min. The lysate was clarified by centrifugation at 1,800 g for 5 min at 4°C, and 100 μL of the resulting supernatant was cultured on LA (Luria-Bertani agar) plates for up to 48 h.

Additionally, larval lysates were assessed by universal bacterial PCR (using primer pair fD1/rP2 targeting the bacterial 16S rRNA gene; amplicon size ∼1.5 kb) and fungal PCR (using primer pair usff/usfr targeting the fungal 18S rRNA gene; amplicon size size ∼485 bp) assays. In brief, total DNA was prepared from larval lysates using a commercially available kit (Qiagen), following the manufacturer’s recommended protocol. After quality and quantity assessment of the DNA preparations, PCR reactions were conducted in 25 μL reactions containing 1x PCR Mastermix, 0.4 μM each of the assay-specific primer pair, and 25 ng of the test DNA. PCR was performed in an automated thermal cycler (BioRad), programmed for sequential incubation of the reaction tubes at 94°C for 2 min, followed by 35 cycles of incubation at 95°C for 15 s, 56°C for 15 s, 72°C for 1 min, and final incubation at 72°C for 7 min. PCR products were run on agarose gels, stained with ethidium bromide, and imaged. Appropriate controls were included in each PCR experiment.

Representative bacterial and fungal PCR results are shown in Figure S4.

Bacteriophage-mediated decolonization of gut bacteria in gnotobiotic larvae

Wherever applicable, monoculture gnotobiotic larvae (B. cereus) were treated by a highly lytic Bacillus phage BCE1 (Siphovirus; latent period ∼20 min; burst size ∼60 infective particles). The phage has a dsDNA genome of 42.1 kb. For phage treatment, 10 gnotobiotic larvae were transferred to each well of a fresh culture plate with 5 mL of sterile water. Subsequently, purified BCE1 phage suspension was added to the wells at a dose of approximately 10⁷ PFU mL^-1. Two days post-treatment, larval specimen was thoroughly washed with sterile water and transferred to fresh plate wells for subsequent examination. Bacteriophage-mediated manipulations were performed inside a BSL2 facility, following microbiological precautions.

Visualization of gut pH profile in larvae by pH indicator dye method

The larval gut pH profile was visualized through the pH indicator dye method, following a previously published protocol,¹² with minor modifications. In brief, larvae from different control and test groups were collected from experimental plate wells and washed with sterile water. Subsequently, 5 larvae were placed in each well of fresh plates holding 5 mL sterile water. Fifty microliters of 1% Phenol Red solution were added to the holding water in each well (0.01% final concentration of the dye). Larvae were allowed to ingest the dye for at least 15 min at room temperature. Before visualization, plates were kept on ice for at least 5 min to reduce larval movement. Larvae were then mounted on clean glass slides (chilled) with a drop of chilled water and imaged using a stereomicroscope and the accompanying software (CellSens). Wherever relevant, larval length was measured using the size measurement tool available in the software and compared to the reference larval lengths specified previously.⁸⁸ Ten to fifteen larvae from each experimental group were examined and data were collected. No modifications in the color or size of the larvae were made during acquisition or analysis of the images.

Effects of antibiotic and carbonic anhydrase inhibitor treatment on gut pH profile

For experiments to examine the effects of removal of bacteria and inhibition of carbonic anhydrase activity on larval gut pH profile, B. cereus gnotobiotic larvae were generated as described in the previous section. Subsets of gnotobiotic larvae were then washed thoroughly and placed separately in well plates holding 5 mL of sterile water. Antibiotic (gentamicin) was added to the designated wells (20 μg mL^-1 final concentration) and incubated for 4 h. Simultaneously, a standard carbonic anhydrase inhibitor (acetazolamide) was added to the designated wells (200 μM final concentration) and treated for 2 h. Untreated axenic and gnotobiotic larvae were included as controls. Post-treatment, gut pH profile was examined as mentioned previously.

Gut pH profile in larvae reared with live bacteria, bacteria cell-free culture supernatant and bacterial cell lysate

Experiments were conducted to verify if cell-free component(s) from bacterial cultures could substitute the requirement of ‘live’ bacteria for the establishment of a gut pH profile. Bacterial cell-free culture supernatant and cell lysate were prepared from log-phase B. cereus cultures. Bacterial cell-free culture supernatant was prepared by centrifuging 2 mL of bacterial culture at 8,000 g for 10 min at 4°C, followed by passing the supernatant through a 0.2 μm syringe filter, and the filtrate was stored at 4°C until use. On the other hand, bacterial cell lysate was prepared by lysing bacterial cells that were pelleted from 2 mL of culture broth. One hundred microliters of 1x PBS buffer solution and zirconia beads (∼0.3-0.4 mm diameter) were added to the pellet, followed by lysis in a homogenizer for 3 min at 25 Hz. Sample tubes and adapters were precooled to minimize damage to heat-labile components. The volume of the lysate was made up to 2 mL (with PBS buffer ) and clarified by centrifugation and filtration, as mentioned previously for bacterial cell-free culture supernatant preparation. Aliquots from both the preparations were cultured on LA plates to rule out the presence of live bacterial cells.

Subsets of axenic larvae were reared separately in 6-well plates in the presence of bacterial cell-free culture supernatant, bacterial cell lysate, live bacteria (B. cereus), and no additive (control) for 3 days. The larval gut pH profile was visualized, and larval growth (length) was recorded under a stereomicroscope as described previously. The length of at least fifteen larvae from each of the experimental groups was examined and data were collected.

Gut pH profile in gnotobiotic larvae colonized with different bacteria

To examine bacterial species-specific effects in the larval gut pH profile, different groups of monoculture gnotobiotic larvae were generated by colonizing axenic larval subsets with various bacterial species (B. cereus, A. hydrophila, M. paraoxydans, L. coleopterorum, L. fusiformis), following the gnotobiotic larvae generation procedure mentioned in the preceding section. Axenic larvae were included as a control group. Gut pH profile was visualized, and larval length was recorded 3 days post colonization to compare the pH profile and variation in growth rates among different groups of gnotobiotic larvae.

Qualitative assessment of proteolytic activity (casein plate method)

The presence of proteolytic activity in different monoculture gnotobiotic larvae was qualitatively evaluated by incubating larval homogenates on standard casein agar plates. In brief, approximately 50-60 larvae from each of the gnotobiotic larval groups (colonized with different bacteria, as mentioned in the preceding section) were collected, rinsed, and homogenized using ∼2.0 mm diameter zirconia beads in a tissue homogenizer, operated at 25 Hz for 3 min. The resulting lysate was clarified by centrifugation at 8,000 g for 10 mins at 4°C. The supernatant was then collected and used for casein hydrolysis assays. A volume of 100 μL of each of the clarified lysates was added separately to designated casein agar plate (supplemented with gentamicin 20 μg mL^-1) wells and incubated overnight at 37°C. Subsequently, zones of hydrolysis were visualized after treating the plates with 2% trichloroacetic acid. The area under zones of hydrolysis was calculated with the help of an open-source computer program, ImageJ v1.54g.⁸⁹

To verify the protease secretion capabilities of the five bacterial species included in the colonization studies, inoculum from log-phase cultures of individual bacteria was streaked separately on casein agar plates. Plates were incubated overnight at 37°C, followed by visualization of the zones of hydrolysis.

Localization of bacteria in larval gut

Experiments were conducted to visualize bacterial localization in the larval gut, using a live bacterial cell stain (BactoView™ Live Green fluorescent dye). Live bacterial cells (B. cereus) were stained as per the manufacturer’s protocol. In brief, 1 mL (∼10⁸ CFU mL^-1) of fresh bacterial culture was centrifuged, and the cell pellet was resuspended in PBS (pH 7.4) buffer. Subsequently, the dye was added to the bacterial suspension to a final concentration of 1x and incubated at room temperature for 30 mins in the dark, followed by centrifugation and resuspension of the labelled cells in fresh PBS buffer. This suspension of labelled bacterial cells was then added (at a final dose of 10⁶ CFU mL^-1) to the 3-day-old axenic larvae held in sterile water in well plates, and the bacterial cells were allowed to colonize the larval gut. Specimen larvae were collected at different time points starting from 5 min to 30 min and visualized under an upright fluorescent microscope (BX43, Olympus, Tokyo, Japan), set to detect fluorescent signal through the FITC channel (absorbance at 500 nm, emission at 520 nm). Images were acquired, processed, and analyzed using the CellSens v2.1 imaging software (Olympus). No modifications in fluorescent intensity were made during acquisition or analysis of the images.

Additionally, live M. paraoxydans was also fluorescently labeled and allowed to colonize another subset of axenic larvae, following the abovementioned protocol. The localization pattern of labelled M. paraoxydans was compared with that of B. cereus 60 min post-colonization. A mock (no-bacteria) labelling reaction was also processed identically and added separately to another subset of axenic larvae to serve as a dye control.

Real-time PCR-based quantification of B. cereus

For qPCR, total DNA was extracted from larval homogenates as described in preceding sections. qPCR was conducted using a commercially available qPCR mastermix to target and amplify B. cereus DNA (using primer pair BsF2 (target-specific)/Bac-Rev; amplicon size ∼645 bp) and a part of the host housekeeping gene (using primer pair Act-2F/Act-8R targeting mosquito actin gene) as reference.⁹⁰ The primer BsF2 was selected among other in-house designed primers based on its specificity and sensitivity towards Bacillus specific template DNA. The efficiency of the primer pair BsF2/Bac-Rev was tested against a dilution series of target template standards, and the results showed good linearity over the range of tested standards with 91% efficiency, R² 0.995, and a slope of -3.557 (Figure S5).

qPCR reactions were conducted in 25 μL reactions containing 1x qPCR Mastermix, 0.2 μM of each assay-specific primer pair, and 25 ng of the test DNA. qPCR was performed by sequential incubation of the reaction tubes at 95°C for 5 min, followed by 40 cycles of incubation at 95°C for 15 s and 65°C for 30 s, and the plate was read during annealing/extension steps. qPCR reactions were run in triplicate on a CFX96 Real Time PCR detection system (Bio-Rad); data were collected and analyzed using an accompanying software, CFX Maestro^TM 1.0 v4.0 (Bio-Rad). The relative quantity of B. cereus in the samples was calculated by the ΔΔCq method using the software.

Total protein, total protease and protease inhibition assays

For protein assays, approximately 50-60 larval specimens from each experimental group were pooled and homogenized (as mentioned in preceding sections) in an optimized cell lysis and protein solubilization reagent. Homogenates were then centrifuged at 8,000 g for 10 mins, followed by collection of supernatants in a fresh tube for downstream protein assays. In addition to larvae, protein preparations were made from bacterial cultures. In brief, overnight bacterial cultures (LB supplemented with casein) were centrifuged at 8,000 g for 5 mins, followed by retention of the supernatant for analysis. The amount of total protein in the preparations was estimated using Bradford assay reagent following the BSA standard curve method, as recommended by the manufacturer.

Total protease activity in the preparations was estimated following a previously described azocasein digestion-based method,³⁸ with minor modifications. In brief, 50 μL of analytes (containing an equivalent amount of total protein) were mixed with 500 μL of 1% azocasein solution in 0.1 M glycine-NaOH buffer (pH 8.0). The mixture was then incubated at room temperature for 60 mins in the dark, followed by treatment with 2% trichloroacetic acid for 10 min to stop the enzymatic digestion reaction. Subsequently, the mix was centrifuged at 10,000 g for 10 mins at room temperature. The supernatant was then mixed with an equal volume of 1 N NaOH solution, and the absorbance was measured at 450 nm using a double-beam UV-Vis spectrophotometer.

The specific nature of protease activity present in a specimen analyte was determined by measuring the residual protease activity after incubation with specific inhibitors/cocktails and comparing it with protease activity in the absence of inhibitor/cocktail. In the present study, a general-purpose protease inhibitor cocktail, a prokaryotic protease inhibitor cocktail, and specific protease inhibitors were used in recommended concentrations, which include serine protease inhibitors (1.0 mM AEBSF, 0.3 μM aprotinin, 10.0 μM leupeptin), an aminopeptidase inhibitor (10.0 μM bestatin), a cysteine protease inhibitor (10.0 μM E64, 10.0 μM leupeptin), an aspartic acid protease inhibitor (110.0 μM pepstatin A), and a metalloprotease inhibitor (5.0 mM EDTA). Additionally, CaCl₂ (10 mM) was also included in the protease assay to verify the presence of Ca²⁺-dependent protease activity. In brief, 50 μL of analyte (containing an equivalent amount of total protein) were separately incubated with and without protease inhibitor/cocktail/CaCl₂ for 60 minutes at RT. Subsequently, residual protease activity was measured following the azocasein method mentioned previously.

Evaluation of the effects of diet on larval anterior midgut alkalinization, protease activity and growth

To evaluate the effects of supplementary diet on larval anterior midgut alkalinization, protease activity and growth, subsets of axenic larvae and gnotobiotic larvae (colonized with Bacillus) were separately reared in absence and presence 5% fish-meal diet, as recommended in Romoli & Gendrin.⁸⁴ Gut pH profile was visualized, larval length was recorded, and protease activity was estimated 3 days post colonization, following the methods described in the previous sections.

Quantification and statistical analysis

Data was initially compiled in a spreadsheet application. For presentation and statistical analysis of the data, appropriate tests were performed using GraphPad Prism. Statistical analysis included one-way ANOVA followed by either Tukey’s or Dunnett’s multiple comparison test, two-way ANOVA followed by Sidak’s test, and unpaired t-tests, wherever appropriate. Data showing differences at various levels of significance, namely p < 0.01, p < 0.001, and p < 0.0001, were expressed by ∗∗, ∗∗∗, and ∗∗∗∗, respectively, wherever applicable in the figures.

Published: October 13, 2025

Footnotes

Supplemental information can be found online at https://doi.org/10.1016/j.isci.2025.113756.

Supplemental information

Document S1. Figures S1–S5

mmc1.pdf^{(406.2KB, pdf)}

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Document S1. Figures S1–S5

mmc1.pdf^{(406.2KB, pdf)}

Data Availability Statement

•
All the data generated in this study are presented within the main/supplementary figures and tables. Bacillus phage BCE1 genome and bacterial 16S rRNA gene sequences have been deposited in GenBank. GenBank accession numbers are provided in the key resources table against the corresponding item.
•
No custom code was generated or used in this study.
•
Any supplementary information essential to reanalyze the data presented in this paper is available from the lead contact upon request.

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PERMALINK

Selected gut bacteria promote mosquito larval growth through the regulation of ionic microenvironment and protease expression

Jhuma Samanta

Sibnarayan Datta

Summary

Graphical abstract

Highlights

Introduction

Figure 1.

Figure 2.

Results

Concurrent presence of bacteria and carbonic anhydrase activity is required for the establishment of a distinctive pH profile in the larval gut

Figure 3.

The continual presence of live bacteria is required for the maintenance of a distinctive pH profile in the larval gut

Figure 4.

Only live bacteria can establish and maintain larval gut pH, and is associated with larval growth

Only selected members of the gut microbiota can establish a distinctive gut pH profile, modulate proteolytic activity, and promote larval growth

Figure 5.

Live bacteria colonize the larval gut rapidly and are localized within the anterior midgut and gastric caeca regions

Figure 6.

Bacteria colonizing the larval anterior midgut contribute different proteases and induce the larva to secrete host proteases

Figure 7.

Supplementary diet alone has no significant effect on larval anterior midgut alkalinization, protease activity, and growth

Figure 8.

Discussion

Conclusion

Limitations of the study

Resource availability

Lead contact

Materials availability

Data and code availability

Acknowledgments

Author contributions

Declaration of interests

STAR★Methods

Key resources table

Experimental model and study participant details

Experimental model

Bacterial strains and growth conditions

Virus strain

Method details

Generation of axenic larvae

Generation of gnotobiotic larvae

Microbiological assessment of axenic and gnotobiotic larvae

Bacteriophage-mediated decolonization of gut bacteria in gnotobiotic larvae

Visualization of gut pH profile in larvae by pH indicator dye method

Effects of antibiotic and carbonic anhydrase inhibitor treatment on gut pH profile

Gut pH profile in larvae reared with live bacteria, bacteria cell-free culture supernatant and bacterial cell lysate

Gut pH profile in gnotobiotic larvae colonized with different bacteria

Qualitative assessment of proteolytic activity (casein plate method)

Localization of bacteria in larval gut

Real-time PCR-based quantification of B. cereus

Total protein, total protease and protease inhibition assays

Evaluation of the effects of diet on larval anterior midgut alkalinization, protease activity and growth

Quantification and statistical analysis

Footnotes

Supplemental information

References

Associated Data

Supplementary Materials

Data Availability Statement

ACTIONS

PERMALINK

RESOURCES

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