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. 2025 Aug 21;20(8):e0330703. doi: 10.1371/journal.pone.0330703

Asaia spp. exposure for improving mosquito mass-rearing, and the effects on Culex pipiens pipiens vector competence for West Nile virus

Alessandro Roman 1,2,#, Charlotte Linthout 2,#, Ben Raymond 1, Constantianus J M Koenraadt 2,*
Editor: Guido Favia3
PMCID: PMC12370026  PMID: 40839618

Abstract

Various vector control strategies are in place to reduce the spread of arthropod-borne viruses. Some of these, such as application of insecticides, are encountering operational challenges and a reduced overall effectiveness due to evolution of resistance. Alternative approaches for mosquito population control, such as the sterile insect technique, depend on efficient mass-rearing of healthy mosquitoes prior to mass-release in the field. Therefore, improving efficiency and quality of mass-rearing techniques is crucial to obtain fit mosquitoes. Previous studies have shown that Acetic Acid Bacteria of the genus Asaia can have a mutualistic effect on larval development in different mosquito species and can thus contribute to improved rearing output. However, whether improved performance in the larval stages may have knock-on effects in the adult stage, for example by increasing their capability to transmit arbovirus, remains unclear. Such effects may jeopardize future control efforts. We tested the effects of two Asaia species, Asaia krungthepensis and Asaia bogorensis, on development time and adult size under two rearing conditions: individual rearing and group rearing of Culex pipiens larvae. Besides investigating development and size, we also investigated whether Asaia spp. exposure during the larval stage can influence the vector competence of Culex pipiens pipiens for West Nile virus (WNV). Our work shows the potential of improving mass-rearing efficiency by employing Asaia krungthepensis as a mutualist for Culex pipiens pipiens. Importantly, this study reveals no significant increase in dissemination and transmission rate of WNV by Culex pipiens pipiens when inoculated with Asaia spp., although an increase in viral titer in the legs and the saliva was observed when the mosquitoes were inoculated with the two Asaia species. Interestingly, we confirmed that Asaia spp. bacteria did not establish as a permanent member of the microbiota of Culex pipiens pipiens. As Asaia spp. did not establish in adult mosquitoes, the observed change in WNV titers can be a result of indirect interactions of Asaia with the native Culex pipiens pipiens microbiome. Our results stress the importance of carefully evaluating host-symbiont interactions to avoid the potential of releasing mosquitoes with enhanced vector competence.

Introduction

Effective mosquito control programs are needed to minimize outbreaks of mosquito-borne diseases such as malaria, chikungunya, Zika, yellow fever, dengue and Rift Valley fever [1]. The control of mosquitoes is key in interfering with the transmission cycle of these viruses, as it limits the frequency of contact between mosquitoes and humans. Various control approaches are in place such as the introduction of natural enemies (e.g., copepods) in water bodies [2,3], habitat elimination, and the application of chemical insecticides [4]. However, frequent and inappropriate use of chemical insecticides has adverse effects on populations of non-target organisms and may result in insecticide resistance in mosquito populations [5].

Novel approaches to mosquito control have been studied intensively over the past years, including the Sterile Insect Technique (SIT) and genetic modification of mosquitoes [6–8]. Both techniques are based on the mass-rearing and release of sterile males or of genetically modified males that must compete with wild-type males once they are released. However, irradiation needed for SIT increases larval and pupal mortalities and decreases adult emergence from pupae [9]. Furthermore, genetic modification of mass-produced mosquitoes may reduce mosquito fitness and sexual competitiveness in different mosquito species [10–12]. There is, therefore, a need for healthier and fitter mass-reared males. Recent studies have shown that the use of symbionts as probiotics can help to improve the fitness of mass-reared mosquitoes [13–15]. In addition, bacterial symbionts such as the obligate intracellular bacterium Wolbachia, can protect mosquitoes from harmful pathogens in mosquitoes and in other insects [16,17].

Possible candidates as beneficial symbionts in mosquito mass-rearing systems are bacteria of the genus Asaia [18–20]. Asaia bacteria have been identified in plants originating from tropical regions [21–23] and reported as spoilage microorganisms of fruit-flavoured bottled water [24–26]. These bacteria are of increasing interest to the scientific community because of their complex interactions with mosquito hosts. For example, Mitraka and collaborators [19] have shown an increase in the development rate of Anopheles gambiae mosquitoes when inoculated with Asaia bacteria. Asaia bacteria are known to inhabit various tissues within mosquito bodies, including the midgut and salivary glands [20], suggesting their potential involvement in vector competence. Therefore, Acetic Acid Bacteria of the genus Asaia have also been studied as potential vehicles for the expression of heterologous effector proteins that, for example, inhibit oocyst formation of Plasmodium berghei or activate immune responses in Aedes aegypti against Dirofilaria immitis [27,28]. In Culicoides nubeculosus a reduction in the abundance of Asaia bacteria in gut bacterial communities due to antibiotic treatment, was associated with an increased infection with Schmallenberg virus [16]. Similar changes in Asaia abundance in Ae. aegypti however, did not lead to any changes in susceptibility to Zika or chikungunya virus [16]. Asaia can thus have a mutualistic effect on larval development in different mosquito species and contribute to improved rearing output. However, whether addition of Asaia as probiotic may have knock-on effects in the adult stage, for example by increasing their capability to transmit arbovirus, remains unclear. As such effects may jeopardize mosquito control efforts, we therefore investigated the effects of Asaia on development of Culex pipiens and their capacity to transmit West Nile virus (WNV).

The first part of our work investigated the hypothesis that various bacterial species within the genus Asaia act as mutualists for Culex pipiens pipiens development when exposed to these bacteria in the rearing water, similar to our earlier observations with Ae. aegypti [29]. We investigated two scenarios: an individual rearing scenario, for which we studied the effects of Asaia spp. treatment on individually reared mosquitoes, avoiding additional effects of competition amongst larvae, and a group rearing scenario to simulate mass-rearing conditions. It is particularly important to demonstrate the potential beneficial effects of Asaia bacteria when provided to larvae with a complex microbiome, since mass-rearing facilities do not employ axenic production techniques. Furthermore, we investigated the effect of the Asaia treatments in the larval stage on the vector competence of adult Cx. p. pipiens females for WNV, as it is inevitable that also a small number of females would be mechanically selected for mass-release, along with the sterile males [30,31].

Results

Development rate of individually reared larvae

We hypothesized that the two species of Asaia act as mutualists for Cx. p. pipiens and reduce the time needed to reach the pupal stage. However, contrary to our hypothesis, exposure to Asaia spp. did not show significant effects on the development rate (Parametric model with Log-logistic error: χ2 = 0.62, d.f. = 2, p = 0.734). At day 15 of the experiment, A. krungthepensis inoculated larvae reached a pupation rate of 89.5%, while A. bogorensis inoculated larvae and the control reached rates of 77.1% and 72.9%, respectively, but these did not differ statistically (Fig 1).

Fig 1. Pupation success over time for Culex pipiens pipiens in the individual rearing after inoculation with two different Asaia species.

Fig 1

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Treatments consisted of 48 biological replicates each. Differences in letters indicate significant differences between the treatments.

Wing length of individually reared larvae

No significant differences in wing length were observed among the treatments, in both males and females (Fig 2, one-way ANOVA: F2, 48 = 1.56, p = 0.227; F2, 50 = 0.58, p = 0.563; respectively).

Fig 2. Wing lengths among Asaia spp. treatments and control in Culex pipiens pipiens in an individual rearing experiment.

Fig 2

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A) Cx. p. pipiens males and B) Cx. p. pipiens females wing length reported in millimeters.

Development rate in group-reared larvae

When the entire experimental period is considered, a significant effect of the Asaia treatment in development rate was observed (Parametric model with Log-logistic error: χ2 = 13.237, d.f. = 2, p = 0.001). It was predominantly A. krungthepensis that accelerated development relative to the control larvae (time ratio = 0.966, z = −2.11, p = 0.035) (Fig 3A). In a mass-rearing facility, achieving a higher daily production of pupae can have significant economic value. In our experiment, the control treatment in the group-reared larvae exhibited similar results as the counterpart in the individually reared larvae, with 50% pupation (P50) reached around day 11 and 75% pupation by day 15 (Fig 1 vs. Fig 3A). Analysis of deviance indicated a significant effect of bacterial treatments at the timepoint in which the control group reached 50% of pupation (χ2 = 46.910, d.f. = 2, p < 0.0001), after controlling for batch-to-batch variability (χ2 = 44.231, d.f. = 1, p < 0.0001) (Fig 3B). More specifically, larvae inoculated with A. krungthepensis reached 64% pupation in comparison with the uninoculated larvae that reached 54% pupation, decreasing the pupation time (mean time changes = −0.291, z = −4.39, p < 0.0001) at day 11. Larvae exposed to A. bogorensis reached 44% pupation at day 11, increasing the pupation time compared to the uninoculated larvae (mean time changes = 0.164, z = 2.38, p = 0.018).

Fig 3. Pupation success over time for group-reared Culex pipiens pipiens larvae inoculated with Asaia bogorensis, Asaia krungthepensis or a control group.

Fig 3

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A) Full time period experiment. B) Graph until 50% pupation was reached by the control group. Different letters (a, b, c) indicate significant differences between the treatments. Each treatment consists of 690 larvae divided into three experimental replicates: 190 larvae in the first experimental replicate and 250 larvae in both the second and third experimental replicate. The horizontal dotted line corresponds to 50% pupation success. Vertical dotted lines correspond to the day in which the treatments reached or overcame 50% pupation.

Wing length in group-reared larvae

The effect of Asaia exposure on female wing length was marginally non-significant (F2, 409 = 2.64, p = 0.073) (Fig 4). However, in males, Asaia exposure significantly affected wing length of the adults (F2, 180 = 18.39, p < 0.001). Specifically, both exposure to A. krungthepensis and to A. bogorensis increased male wing length compared to the control (t180 = 5.95, p < 0.0001; t180 = 4.01, p < 0.001). No differences were observed between A. krungthepensis and A. bogorensis. (t180 = 1.95, p = 0.127).

Fig 4. Wing lengths among Asaia spp. treatments and control in Culex pipiens pipiens males (A) and females (B) reared in a group-rearing setting.

Fig 4

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Wing length reported in millimeters. Asterisks (*) indicate significant differences (*** = p < 0.001).

WNV vector competence

We infected eight to twelve days old Cx. p. pipiens mosquitoes with WNV via an infectious blood meal. These mosquitoes emerged from the experiment with group-reared larvae and had thus been exposed to A. bogorensis or A. krungthepensis during larval development. As a control for the vector competence studies, we used Cx. p. pipiens mosquitoes that had not been inoculated with Asaia bacteria during their larval development. An average of 74% of the Cx. p. pipiens adult females took up a WNV-spiked blood meal with a titer ranging between 2 x 106 and 1.12 x 107 TCID50/ml for the three replicates. Due to time restrictions, not all mosquitoes that were still alive after the incubation period of 14 days, were used in the forced salivation assay (see Table 1).

Table 1. Overview of the feeding rate, survival rate, number of females used, and infection, dissemination and transmission rates per treatment.

Treatment Feeding rate (%) Survival rate (%) Number of females used for salivation assay/Number available Infection rate (%) Dissemination rate (%) Transmission rate (%)
Control 167/214 (78%) 160/167 (96%) 132/160 97/132 (73%) 52/132 (39%) 28/132 (21%)
Asaia krungthepensis 131/178 (69%) 120/131 (92%) 120/120 94/120 (78%) 50/120 (42%) 30/120 (25%)
Asaia bogorensis 173/218 (74%) 168/173 (97%) 134/168 92/134 (69%) 50/134 (37%) 30/134 (22%)
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Our GLMM model with wing length and titer of the blood meal as random effects included, showed that the Asaia spp. treatments did not have a significant effect on the infection, dissemination, and transmission rate of Cx. p. pipiens mosquitoes with WNV (GLMM, LTR χ2 = 4.414, d.f. = 2, p = 0.110; LRT χ2 = 0.271, d.f. = 2, p = 0.873; LRT χ2 = 0.955, d.f. = 2, p = 0.620; respectively) (Fig 5).

Fig 5. Infection (A), dissemination (B), and transmission (C) rates of WNV infected Culex pipiens pipiens mosquitoes.

Fig 5

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These mosquitoes were inoculated during larval development with either Asaia bogorensis, Asaia krungthepensis or neither of these Asaia species (control). The dots represent the rates of each individual replicate within a treatment.

Although no effects of treatment were found concerning infection, dissemination, and transmission rates, we also investigated the effect of Asaia treatment on virus titer in the bodies, legs and saliva of the infected mosquitoes (Fig 6). No effects of Asaia spp. treatment on the viral titers in the bodies were detected (LRT: p= 0.751). The bacterial treatments did, however, influence the titer of WNV in mosquito legs (LRT: p < 0.001) after controlling for the blood meal titer effect (LRT: p < 0.001). Inoculation with A. krungthepensis significantly increased WNV titer in legs of Cx. p. pipiens adult females compared to the control (censReg: Estimate = 0.66, t-value = 4.07, p < 0.001) (Fig 6B). After controlling for experimental replicate (LRT: p = 0.69), bacterial treatment also significantly affected WNV titers in saliva (LRT: p = 0.048). Specifically, adult females derived from larvae exposed to A. bogorensis showed an increased titer compared to the control treatment (censReg: Estimate = 0.65, t-value = 2.48, p = 0.013) (Fig 6C).

Fig 6. West Nile Virus (WNV) titers in body (A), legs (B) and saliva (C) samples of Culex pipiens pipiens adult females.

Fig 6

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These females were inoculated with Asaia krungthepensis or Asaia bogorensis bacteria during larval development. The virus titers were compared to a control treatment, which was not inoculated with Asaia bacteria during larval development. Asterisks (*) indicate significant differences (GLMM, * = p < 0.05), ns = not significant.

Detection of Asaia spp. in uninoculated larvae, treated and untreated female adults

We hypothesize that an inoculum of Asaia spp. provided during larval development can establish as a microbiota component in mosquito larvae, and potentially be transstadially transmitted to the adult generation. Cx. p. pipiens larvae, hatched from mosquito egg rafts collected from the mosquito rearing showed no amplification for the genus Asaia. This data confirmed the absence of Asaia bacteria in our laboratory population of Cx. p. pipiens (Figure 7A). Similarly, adult females from the control did not amplify the Asaia DNA fragment, nor did the females that emerged from Asaia spp. exposed larvae (Fig 7B).

Fig 7. PCR detection of Genus Asaia in larvae and adult females. A) PCR indicates that no bacteria of the genus Asaia were detected in newly hatched larvae (pools of five larvae in sample 1–7). B) PCR for the detection of Asaiaspp. in adult females. Culex specific primers were used as positive control of DNA extraction and PCR procedure. The symbols + and – correspond to the positive Asaia and the negative (water) control, respectively.

Fig 7

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Discussion

Fit adult mosquitoes are needed for mass-release programs to reduce mosquito populations worldwide [9,10]. Optimizing the larval diet may be one option to produce these fit mosquito populations [13–15]. However, an increase in vector competence that could be associated with more fit mosquitoes, is not favoured. In this study, we therefore investigated how inoculating Cx. p. pipiens first instar larvae with two different species of Acetic Acid Bacteria, belonging to the genus Asaia, affect fitness parameters (development rate, wing size) and vector competence for WNV.

Interestingly, the two Asaia spp. did not have an equal effect on development of the group-reared larvae. Larvae inoculated with A. krungthepensis reached 64% pupation around day 11, compared to uninoculated larvae that only reached 54% pupation and A. bogorensis inoculated larvae that reached only 44% pupation on the same day. Inoculation with A. krungthepensis therefore leads to a faster pupation. This accelerated development for A. krungthepensis may be advantageous for mass-rearing, as it will result in faster daily production of pupae [32,33], consequently reducing production costs [34–36]. Area-wide inundative mass-release vector control methods, such as the Sterile Insect Technique (SIT), often require releasing male mosquitoes at a ratio of 5–10 males per wild male. Recent advancements in larval rearing systems have enabled the production of approximately 100,000 male pupae per week in just 2 m² of rearing space [36]. With the increasing frequency of vector-borne disease outbreaks, the demand for sterile male mosquitoes in vector control programs is expected to rise. Therefore, reducing the development time from newly hatched larvae to pupae is critical for enhancing the weekly production capacity of mass-rearing facilities. Our findings, which show differences in pupation speed, suggest that optimizing these traits could improve production efficiency and make mass-rearing systems more cost-effective, helping meet the growing demand for sterile mosquitoes in vector control. However, its scalability and consistency under operational conditions require further consideration. While our laboratory experiments demonstrate that one Asaia spp. accelerates larval development, factors such as environment, rearing conditions, mosquito species, microbial competition in rearing water, and the stability of Asaia spp. cultures over time may influence its performance at scale. Implementing the usage of Asaia spp. in mass rearing facilities would necessitate pilot-scale trials and long-term assessments to validate its effectiveness under real-world conditions and to optimize protocols for routine application in mass-rearing programs.

Similar to these results, Asaia increases the development rate of An. gambiae, possibly by modulating the transcription of genes involved in the structural components of the cuticle [19]. However, in contrast with our results for Cx. pipiens, bacteria of the genus Asaia were stably associated in An. gambiae larvae, as they became part of the midgut microflora and were able to be transmitted transstadially [19]. Furthermore, another study demonstrated that Anopheles stephensi larvae deprived of Asaia bacteria, showed asynchrony in moulting and therefore delayed development [18]. A notable difference from our study lies in the characterization of Asaia within An. stephensi’s conventional microbiota component. Our results suggest that in Cx. p. pipiens, the successful colonization and transmission of A. krungthepensis or A. bogorensis may be compromised by microbial competitors present within the native microbiota, as previously shown for Ae. aegypti mosquitoes [29]. Although Asaia spp. was not detected in adult females, we acknowledge that the limited sample size precludes a definitive conclusion about persistence. In addition to microbial competition, the absence of detectable Asaia spp. may also reflect microbial turnover associated with metamorphosis, during which larval microbiota are frequently lost or reshaped. Additional replicates and alternative experimental approaches may help clarify the conditions required for persistence. Concerning the adult size (wing length), no differences were found neither in males nor in females within the two Asaia spp. treatment groups and the control when the larvae were reared individually. Similar results were obtained in Ae. aegypti when inoculated with Escherichia coli or Asaia sp. [14]. In the group-rearing condition, no differences were observed regarding Cx. p. pipiens female size, but inoculation of both Asaia spp. during larval development resulted in larger males compared to the uninoculated control larvae. In summary, our study suggests that particularly A. krungthepensis exhibits potential as larval mutualist within Cx. p. pipiens mosquitoes, based on both accelerated development time and larger adult males derived from both the Asaia spp. inoculated treatments. This species can thus serve as a candidate for vector control techniques based on mass-release of males, such as SIT. Several studies have shown that larger male body size is advantageous for survival and sperm capacity [37,38]. Male dispersal in the wild and mating success are determined by their fitness, with body size serving as a quality indicator. These aspects can directly affect longevity, the number of times a male can mate, and sperm capacity, leading to an increase or decrease of the overall efficacy of the vector control programs [38]. Our vector competence study demonstrates that, although there was no effect of treatment on the dissemination and transmission rate, we did observe increased WNV dissemination and transmission titers in Cx. p. pipiens adults derived from Asaia-exposed larvae. Interestingly, the two Asaia spp. influenced dissemination and transmission titers differently. Specifically, the exposure to A. krungthepensis significantly enhanced disseminated viral titers of mosquitoes compared to the control group, whereas A. bogorensis enhanced the WNV transmission titers in saliva. It is known that midgut and salivary gland barriers can be divided into the infection and escape barrier [39], and that viral infections can be limited by these barriers, mechanically as well as through antiviral immune responses. In addition, mosquito symbiotic bacteria in the gut can also have an important immunological role [40–42]. In our experiment, the transient infection with A. krungthepensis bacteria during larval development may alter the efficacy of the midgut barriers, but not the protective nature of the salivary gland barriers of Cx. p. pipiens adult females. Conversely, exposure to A. bogorensis bacteria resulted in Cx. p. pipiens females exhibiting greater WNV transmission potential compared to the non-exposed control group. In support of our findings, several studies have shown the role of mosquito gut microbiota in pathogen development and competence [43–45]. Similar to our results, the effect of A. bogorensis can be seen in Anopheles stephensi in which it can promote infection and gametogenesis of Plasmodium berghei [46]. Promotion of infection has been attributed to alkalization of the midgut due to proliferation of A. bogorensis in glucose-feeding adults [46]. Conversely, Asaia can stimulate the basal immunity and therefore reduce the development of malaria parasite oocysts in An. stephensi [47]. On the other hand, alterations in the relative abundance of Asaia induced by antibiotic treatment in Aedes aegypti did not lead to changes in susceptibility to Zika nor chikungunya virus [16]. In the Culex pipiens complex, the microbiota could be an important and explanatory factor for the variation in vector competence for WNV [48]. Furthermore, the reduction in abundance of the protective microbiota (Wolbachia) mediated by climate warming [49] can lead to increased WNV viral replication in the mosquito host [48]. Similarly, recent work provided evidence that the exposure to different bacteria during larval development can influence adult traits in Aedes aegypti as well as its susceptibility to systemic dissemination of dengue virus and diversity in the innate immune response [50]. In short, these studies show that it is likely that presence of or exposure to Asaia has effects on arbovirus replication, although reports have been conflicting regarding the directionality of this effect.

Conclusion

Inoculation of Cx. p. pipiens first instar larvae with A. krungthepensis led to an initial acceleration of development time from newly hatched larvae to pupae. It also increased the body size of the emerging males from our mass-rearing set-up. Both the shorter development time and increased body size of male mosquitoes may improve mass-rearing conditions for Cx. p. pipiens mosquitoes considering mass-release based vector control applications. Vector competence studies with female Cx. p. pipiens mosquitoes showed that Asaia spp. inoculation of Cx. p. pipiens mosquitoes had no significant effect on infection, dissemination, and transmission rates of WNV. However, it did influence the WNV titer in the legs and the saliva of these mosquitoes, even though Asaia spp. bacteria were not detected in adults following larval exposure, suggesting that these bacteria did not persist or stably establish in the mosquitoes. Indirect effects of A. bogorensis and A. krungthepensis on the microbiota of the mosquitoes could be the underlying cause of these altered virus titers.

Materials and methods

Culex pipiens pipiens mosquitoes

The Cx. p. pipiens rearing was established in the summer of 2020 by collecting egg rafts in oviposition traps in the proximity of chicken coops across Wageningen, the Netherlands. After identifying a pool of 10 larvae per egg raft for their biotype by real-time PCR [51], the remaining larvae from the same egg raft and with biotype pipiens were grouped in trays to start a new rearing colony. The mosquitoes were maintained at 23°C, 60% relative humidity and a photocycle of 16:8 light:dark [52]. F19 offspring of this population was used in the experiments.

Asaia bacteria cultures

Two Asaia isolates were used for all the experiments, Asaia krungthepensis isolate 1A and Asaia bogorensis isolate 3.10. A. krungthepensis isolate 1A was kindly provided by Dr. Cynara Rodovalho (FIOCRUZ, Rio de Janeiro, Brazil) and A. bogorensis isolate 3.10 was provided by Marc Hendriks from Wageningen University & Research. Asaia spp. were kept as glycerol stocks at −80°C for long-term storage. Cultures of Asaia spp. were grown in GLY liquid media (Glycerol 25 g/l, Yeast Extract 10 g/l, pH 5) at 30°C and 200 rpm shaking. Prior to the experiments, a single colony from each Asaia spp. was pre-inoculated in 3 ml GLY liquid media using 15 ml conical polypropylene centrifuge tubes (Fisherbrand™) for 8 hours at 30°C and 200 rpm, reaching an OD600 of 0.1. Next, 5 ml of sterile GLY liquid media was inoculated with 50 μl of the pre-inoculation culture and incubated for 16 hours at the same incubation conditions as mentioned previously, to reach an OD600 ⋍ 1. Asaia spp. growth was quantified in CFU/ml by serial dilution and plating on GLY agar plates. The targeted inoculation level at time 0 of our experiments was set at approximately 2000 CFU of Asaia spp. per ml of rearing water, using a 100-fold dilution of overnight culture, washed three times in phosphate buffer saline (PBS) and resuspended in sterile tap water.

West Nile virus

A Dutch isolate of a passage 3 WNV lineage 2 (EMC/WNV/20TV2584/NL) was used for mosquito oral inoculation. This virus was isolated during the WNV outbreak in 2020 in the Netherlands and was kindly provided by Erasmus Medical Center (Rotterdam, the Netherlands). WNV was grown at 37°C on a monolayer of Green monkey kidney Vero E6 cells, cultured with Hepes-buffered Dulbecco’s Modified Eagle Medium (DMEM; Gibco, Carlsbad, CA, USA) and supplemented with 10% fetal bovine serum (FBS; Gibco), Gentamicin (50 μg/ml, Life technologies), Fungizone (2.5 μg/ml, Life technologies) and a combination of penicillin (100 U/ml; Sigma-Aldrich, Saint Louis, MO, USA) and streptomycin (100 μg/ml; Sigma-Aldrich, Zwijndrecht, the Netherlands) (P/S). Titrations of the WNV stock, blood meals and virus-positive samples were done on the same cell line with the same supplemented medium to determine the viral loads. This medium is hereafter referred to as DMEM Hepes complete.

Development after exposing individually reared larvae to Asaia spp

Climatic conditions for experiments were as described above for the Cx. p. pipiens rearing. An experiment consisting of three treatment levels (A. krungthepensis, A. bogorensis and Control) was performed. Cx. p. pipiens eggs were hatched in sterile tap water, after which the larvae were transferred individually to sterile 12-well plates with lids that enable gas exchange (Cellstar™). Each well was provided with 3 ml autoclaved tap water. Each treatment consisted of 48 biological replicates, with each replicate being a single larva. Single neonate larvae were inoculated depending on the designed treatment with approximately 2000 CFU/ml of either A. krungthepensis or A. bogorensis. We provided 2.85 mg of sterilized TetraMin® Baby fish food (Tetra Werke Company, Melle, Germany) per larva, over 10 days: 0.15, 0.3, 0.6, 0.6, 0.6 and 0.6 mg on days 0, 2, 4, 6, 8 and 10, respectively. The food was prepared as a stock solution of 100 µg/µl in purified water (Milli-Q® Reference A+ System, Merck Millipore) and autoclaved. Survival and pupation were recorded daily until the end of the experiment. Additionally, adult wing length measurements were taken as an estimate of body size [53,54]. For this purpose, wings were removed and mounted on sticky tape on a glass microscope slide. Wing length was measured as the distance between the alula and the wing tip, excluding the fringe scales using a Dino-Lite Edge 5 MP digital microscope and DinoCapture 2.0 software (Dino-Lite Europe, Almere, NL). The set-up was calibrated with a slide graticule of 0.01 mm.

Development after exposing group-reared larvae to Asaia spp

To simulate mass-rearing conditions, whole eggs rafts of Cx. p. pipiens were placed on autoclaved tap water to achieve synchronous hatching. For each treatment, one plastic 195 x 195 mm tray was used. This experiment consisted of three replicates. The first replicate consisted of ~ 190 larvae/treatment tray. The tray was filled with 570 ml sterile tap water resulting in 3 ml per larva. For the second and third replicates, we aimed for ~ 250 larvae/tray filled with 750 ml sterile tap water. In the trays designed for A. krungthepensis and A. bogorensis treatments, approximately 2000 CFU/ml following the same steps as in the individual rearing study, were added at day 0 immediately before the newly hatched larvae were transferred to the trays. Sterile TetraMin® Baby fish food was provided to obtain 0.15 mg/larva on the initial experimental day. In total, approximately 2.5 to 2.8 mg food per larva was provided during development. Survival and pupation were recorded daily until the end of the experiment. Pupae were collected in 30 ml plastic cups and separated by treatment in 30 cm cubic BugDorm-1 insect rearing cages (MegaView Science Co., Ltd. Taiwan). Emerging adults were fed ad libitum with 6% glucose solution and were allowed to mate for 8–12 days. For all three treatments (A. krungthepensis, A. bogorensis and Control), 8–12 days old Cx. p. pipiens females (n = 198 + /- 20) were used in vector competence studies. Additionally, adult wing length measurements were taken as an estimate of body size [53,54].

Vector competence of adults exposed to Asaia from the group-rearing experiment

In the biosafety level 3 (BSL-3) facilities of Wageningen University, the mosquitoes were allowed to feed on a WNV spiked chicken blood meal via a Hemotek PS5 feeder system. The final virus titer of the blood meals was measured and calculated to range between 2.0*106 and 1.12*107 TCID50/ml (verified by End Point Dilution Assay, EPDA). After one hour of blood feeding, mosquitoes were anesthetized with CO2 and the fully engorged females were selected. These mosquitoes were placed at 23°C in plastic buckets (top diameter 12.5 cm x height 11.5 cm x bottom diameter 10.5 cm) with a 16:8 light:dark cycle for a period of 14 days. As a food source, 6% glucose solution was provided ad libitum via cotton wool on top of the bucket for the entire incubation period.

Forced salivation assay

After incubation for 14 days, all females were immobilized using CO2. The right wing of each mosquito was kept to measure wing length. The left wing and all legs of each female were removed and collected in 1.5 ml SafeSeal Eppendorf tubes, containing 5 mm zirconium oxide beads (Next Advance, AverillPark, NY, USA), without any medium. Saliva samples were collected by inserting the proboscis into a 200 μl pipet tip, containing 5 μl of a solution of 50% FBS and 25% sugar in tap water. After 45 minutes, the body was removed and stored in a 1.5 ml SafeSeal Eppendorf tube containing 5 mm zirconium oxide beads, without any medium. The individual mosquito saliva samples were then resuspended in 55 μl DMEM Hepes complete and stored in 1.5 ml Eppendorf tubes. All individual body, leg and saliva samples were stored at −80°C until further use in the infectivity assay.

Infectivity assay and virus titration

Frozen body and leg samples were initially processed without medium using the Bullet Blender Storm (Next Advance, USA) and the Eppendorf Minispin Plus (Eppendorf, Hamburg, Germany), following the protocol of Vogels et al. [55]. Next, 100 μl of fresh DMEM Hepes complete medium was added to the body samples, and 60 μl to the legs and wing samples. These were then reprocessed in the Bullet blender Storm and the Eppendorf Minispin Plus, as described by Vogels et al. [55]. Subsequently 30 µl of either mosquito body, legs or saliva samples were added to each well of a 96 well plate with a 90% confluent monolayer of Vero E6 cells in 60 µl DMEM Hepes complete. After incubation for two hours at 37°C, 100 μl of fresh DMEM Hepes complete medium was added to the wells. The plates were then incubated at 37°C for six days. Following this incubation period, the wells were examined for the presence or absence of the virus, based on the presence or absence of cytopathic effect (CPE). The infection, dissemination, and transmission rates were determined as the percentage of WNV-positive body, leg, or saliva samples, respectively, of the total number of mosquitoes analysed. For a subset of the body, leg and saliva samples that were scored as virus-positive, the viral titers were determined via EPDA on Vero E6 cells.

Detection of Asaia spp. in uninoculated larvae, treated and untreated female adults

Polymerase Chain Reaction (PCR) was employed to confirm the presence or absence of bacteria of the genus Asaia. Newly hatched Cx. p. pipiens larvae from the mosquito rearing and emerged female adults from the three individual rearing experimental treatments were collected, and stored at −20°C. Prior to DNA extraction, all samples were washed three times in sterile PBS solution and homogenized in 180 µl cold sterile PBS using a TissueLyser II instrument (Qiagen). DNA from homogenized samples was extracted with MagBind ® HDQ blood and tissue DNA 96 Kit (Omega Bio-tek, Inc., Norcross, Georgia, USA), according to the manufacturer’s instructions. Extracted DNA was tested with specific Asaia primers. The primer sequences used for Asaia were AsaiaPrimer f (5’ – GGCGCGTAGGCGGTTTACAC) and AsaiaPrimer r (5’ – TGCGCGTTGCTTCGAATTAAACCA) [56]. Amplification of Asaia specific DNA results in a 400 bp PCR product. PCR settings consist of an initial denaturation step of 95°C for 5 minutes, 35 cycles of 94°C for 30 sec, 55°C for 45 sec, 72°C for 30 sec and a final extension at 72°C for 7 minutes.

To validate the DNA extraction and the PCR, also the CQ11 microsatellite-specific primers for Culex pipiens were used: CQ11F2 (5’ – GATCCTAGCAAGCGAGAAC) and pipCQ11 r (5’ – CATGTTGAGCTTCGGTGAA) [57]. For CQ11 the PCR conditions were set as follows: 95°C for 5 min initial denaturation step, 40 cycles of 94°C for 30 sec, 54°C for 30 sec, 72°C for 40 sec, and a final extension at 72°C for 5 minutes. We tested seven pools of five newly hatched larvae per pool for presence/absence of Asaia at the beginning of the experiment, prior to inoculating the larvae with the bacteria. Furthermore, we evaluated the establishment of Asaia in adults by testing five individual females per treatment. DNA extracted from overnight cultures of A. krungthepensis and A. bogorensis was used as positive control for Asaia-specific primers. PCR products were run on a 2% agarose gel for 60 min at 80 V.

Statistical analysis

All experimental data were analysed in RStudio version 2022.07.2 [58].

Individual rearing experiment.

Data on time to pupation were analysed using parametric models with Log-logistic errors in the Survival package [59] with bacterial treatment (factor with three levels, i.e., Control and the two Asaia spp.; A. krungthepensis and A. bogorensis) as an explanatory variable. Graphical check of the linearity of the predictors, and model fit with Cox-Snell residual analysis were performed. We also used the Akaike information criterion (AIC) to compare Log-logistic models to other error structures or analysis approaches (e.g., non-parametric models). Histograms, Shapiro-Wilk tests and Fligner tests were used to explore wing length of both male and female mosquitoes. Differences in wing lengths between the treatments for the individual rearing experiment were analysed by one-way ANOVA aov function [58].

Vector competence experiment.

Time-to-pupation was analysed using parametric models with Log-logistic errors in the Survival package [59] with bacterial treatment (factor with three levels, i.e., Control, A. krungthepensis and A. bogorensis) and experimental replicate as explanatory variable (experimental replicate was included first in the model to control for variability across batches, followed by treatment to evaluate its effect independently of batch differences). Additionally, the pupation time differences among the bacterial treatments were examined once more at the timepoint when the uninoculated control treatment had achieved 50% pupation, using parametric models with Gaussian errors and the same explanatory variables. Mosquito mass-production benefits from a faster P50 (pupation 50%) time [34]. Graphical check of the linearity of the predictors, and residual analysis were performed. The Akaike information criterion (AIC) and Bayesian information criterion (BIC) were used to compare Log-logistic models to other error structures or analysis approaches (e.g., non-parametric models). Male and female wing length measurements for the vector competence experiment were analysed with a linear mixed-effects model fitted by maximum likelihood (ML), with bacterial treatment as explanatory variable and experimental replicate number as random effect [60]. Outlier values were checked and eventually removed by using the Grubb test included in the package outliers [61]. Model assumptions were checked with graphical analyses of error distribution. When a significant effect of bacterial treatment was found, estimated marginal means were calculated for pairwise comparison, employing the Emmeans package version 1.8.8 [62].

To assess the effects of treatment (A. krungthepensis, A. bogorensis and control) during the larval stage, while controlling for the wing length of the adult females, and the titer of the infectious blood meal on WNV infection, dissemination and transmission rates, Generalized Linear Mixed-Effects Models (GLMM) with binomial distribution were performed, using the lme4 package [60]. Infection, dissemination, and transmission rates (dependent variables) were calculated, respectively, by dividing the number of female mosquitoes with infected bodies, infected legs or infected saliva by the total number of female mosquitoes in the respective treatment.

The effect of treatment on viral titers in body, leg and saliva, were analysed with Censored Regression (Tobit) Models in RStudio using the censReg package version 0.5–36 [63]. By using this package, we can deal with WNV-positive samples that have titer values below the detection limit of 2000 TCID50/ml by setting them at 1000 in the left censored model formula. Bacterial treatment and blood-meal titer were employed as explanatory variables. All the analyses were carried out on the log10 values of the titers.

Supporting information

S1 Data. Raw image.

(PDF)

pone.0330703.s001.pdf (323.7KB, pdf)

Acknowledgments

Thanks to Dr. Cynara Rodovalho and Marc Hendriks for providing Asaia isolates. Thanks to Pieter Rouweler and the other members of the insect rearing group from the Laboratory of Entomology from Wageningen University & Research for providing mosquito eggs for the experiments. For the vector competence analysis, Dr. Gerrit Gort is thanked for the input and help. We also thank Marcel Dicke and Gorben Pijlman for feedback on an earlier version of this chapter.

Data Availability

The data that support the findings of this study are publicly available from DANS Data Station Life Sciences with identifier https://doi.org/10.17026/LS/NWTCLU.

Funding Statement

AR was funded by the ‘Insect Doctors’ program, which has received funding from the European Union’s Horizon 2020 research and innovation program under the Marie Skłodowska-Curie grant agreement No.859850. CL was partly funded by the Dutch Research Council (NWO) as part of the project ‘Preparing for Vector-Borne Virus Outbreaks in a Changing World: a One Health Approach’ (NWA.1160.18.210). The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

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Associated Data

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

Supplementary Materials

S1 Data. Raw image.

(PDF)

pone.0330703.s001.pdf (323.7KB, pdf)

Data Availability Statement

The data that support the findings of this study are publicly available from DANS Data Station Life Sciences with identifier https://doi.org/10.17026/LS/NWTCLU.

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