Wolbachia Density and Cytoplasmic Incompatibility in Aedes albopictus: Concerns with Using Artificial Wolbachia Infection as a Vector Suppression Tool

Maurizio Calvitti; Francesca Marini; Angiola Desiderio; Arianna Puggioli; Riccardo Moretti

doi:10.1371/journal.pone.0121813

Abstract

The mosquito Aedes albopictusi is a competent vector of harmful human pathogens, including viruses causing dengue and chikungunya. Cytoplasmic incompatibility (CI) induced by endosymbiotic Wolbachia can be used to produce functionally sterile males that can be released in the field as a suppression tool against this mosquito. Because the available sexing methods are not efficient enough to avoid unintentional release of a few transinfected females, we assessed the CI pattern in crosses between wPip Wolbachia-transinfected (ARwP) females and wild-type males of Ae. albopictus in this study. Quantitative polymerase chain reaction was used to monitor the titer of the Wolbachia strains that naturally infect Ae. albopictus, that is, wAlbA and wAlbB, in age-controlled males and females. Data were coupled with incompatibility level detected when the above-mentioned males were crossed with ARwP females. Wolbachia infection titer was also monitored in samples of wild caught males. Incompatibility level was positively correlated only with wAlbA density. Crosses between wild-type males having very low wAlbA density (<0.001 wAlbA/actin copy numbers) and ARwP females were partially fertile (CI_corr = 68.06 ± 6.20). Individuals with low wAlbA titer were frequently found among sampled wild males (30%–50% depending on the site and period). ARwP males can be as considered as a very promising tool for suppressing Ae. albopictus. However, crosses between wild males having low wAlbA density and ARwP females may be partially fertile. In the case of local establishment of the transinfected mosquito line, this occurrence may favor the replacement of the wild-type mosquitoes with the ARwP line, thus reducing the long-term efficacy of incompatible insect technique. Various alternative strategies have been discussed to prevent this risk and to exploit Wolbachia as a tool to control Ae. albopictus.

Citation: Calvitti M, Marini F, Desiderio A, Puggioli A, Moretti R (2015) Wolbachia Density and Cytoplasmic Incompatibility in Aedes albopictus: Concerns with Using Artificial Wolbachia Infection as a Vector Suppression Tool. PLoS ONE 10(3): e0121813. https://doi.org/10.1371/journal.pone.0121813

Academic Editor: Kostas Bourtzis, International Atomic Energy Agency, AUSTRIA

Received: October 9, 2014; Accepted: February 4, 2015; Published: March 26, 2015

Copyright: © 2015 Calvitti et al. This is an open access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited

Data Availability: All relevant data are within the paper and its Supporting Information files.

Funding: These authors have no support or funding to report.

Competing interests: The authors have declared that no competing interests exist.

Introduction

Aedes (Stegomyia) albopictus (Diptera: Culicidae), commonly known as Asian tiger mosquito, is one of the most invasive insect species worldwide [1,2]. It shows high vector competence for several arboviruses, including viruses causing dengue (DENV) and chikungunya (CHIKV) [3]. Although Ae. albopictus is not considered as the primary vector of DENV, it is becoming a major cause of viral outbreaks because of rapid changes in its overall distribution. In the last decade, Ae. albopictus caused new DENV epidemics in different countries such as Hawaii [4], Mauritius [5], and China [6]. With respect to CHIKV, a recent mutation in genes encoding envelope glycoproteins of CHIKV of an African lineage enhanced the adaptability of this virus to Ae. albopictus [7], leading to outbreaks in Indian Ocean islands in 2005–2006 [8] and in temperate regions such as Italy [9] and France [10]. The CHIKV mosquito-human transmission cycle recently has established in the Caribbean [11]. The associated epidemic is currently out of control, with more than 500,000 cases in a few months, and models are being developed to predict the possible spread of the virus in the Americas [12].

In recent years, interest in mosquito control strategies based on the principles of autocidal control, such as release of radio-sterilized males (Sterile Insect Technique, SIT) in field [13], has increased, thus providing new approaches by using innovative biotechnology (such as insect transgenesis and endosymbiont manipulation) [14–17]. The potential to combine these recent approaches with the classical SIT may offer viable solutions to overcome intrinsic limitations of the current conventional vector control strategies that mostly rely on insecticides and community participation. In addition to exerting negative effects on non-target insects and having toxicological impact on humans and environment, insecticides result in the establishment of resistant strains, as shown in some recent reports [18,19]. Therefore, alternative strategies to control mosquitoes are being sought worldwide.

Research on the endosymbiont Wolbachia pipientis (Alphaproteobacteria: Rickettsiales) has increased steadily in the last two decades, driven by the possibility of exploiting its biological properties as tools for insect pest and vector control [16,20]. This maternally inherited bacterium manipulates host reproduction and is carried by approximately 40% arthropod insect species as well as some crustaceans, mites, and filarial nematodes [21]. Cytoplasmic incompatibility (CI) is the most common reproductive phenotype observed in arthropod species infected with specific Wolbachia strains [,22–27]. First described in mosquito Culex pipiens [28–30], CI is a conditional embryonic lethality that occurs when males infected with CI-inducing Wolbachia strains are crossed with uninfected females (unidirectional CI [Uni-CI]) or with females carrying other incompatible Wolbachia strains (bidirectional CI [Bi-CI]).

In 2009, an incompatible Ae. albopictus strain (ARwP) was generated to support a SIT project against Ae. albopictus in Italy [31]. ARwP was obtained by replacing natural Wolbachia double infection with a single heterologous strain of Wolbachia (wPip) taken from Culex pipiens molestus [32]. The transinfected wPip strain was attributed to the wPip-IV incompatibility group (Mylene Weill, pers. com.) according to a classification by Atyame et al. [33]. Infection parameters (maternal inheritance and fitness costs), male mating competitiveness performances, and CI features (induction of complete and not age-dependent CI when wPip-transinfected males are crossed with wild-type females) were investigated both at laboratory level and on a semi-field scale [34,35]. The results of these studies were consistent with the traits needed to use the ARwP strain as the provider of ready-made sterility-inducer males for incompatible insect technique (IIT), which is an alternative autocidal approach based on the release of biologically incompatible males rather than irradiated males [15,36,37].

However, some major drawbacks have to be overcome, particularly against mosquito vector species, before IIT can be practiced in field. One of the major constraints is the inevitable co-release of females infected by a non-native Wolbachia strain that results in the release of incompatible males. This is because of the absence of an efficient sexing technology for the perfect separation of male and female pupae. In fact, at least 1% female contamination [38] is expected during the release of males. Model simulations based on laboratory experiments clearly predict that in cases where IIT strategy relies on a strong Uni-CI pattern (i.e., when the target population is uninfected), accidental co-release of Wolbachia-infected males and females may lead to an unwanted replacement of uninfected targeted population with the new infected population [39]. This would make IIT progressively ineffective. However, prediction becomes more complex when the target population harbors an incompatible Wolbachia infection type (Bi-CI), (as an example, if using the ARwP strain against the naturally infected Ae. albopictus). In this case, local coexistence of two different Wolbachia infection types may result in an unstable equilibrium that will evolve over time, resulting in the fixation of either one of the two infection types. According to Dobson et al. [39], this outcome would theoretically depend on two main factors: (i) pattern of CI (Uni-CI or Bi-CI) between the two populations and (ii) their competition both at larval and adult stages.

In this study, we provide new evidence on the first factor, i.e., CI pattern between naturally infected Ae. albopictus males (coinfected with wAlbA and wAlbB Wolbachia strains) and wPip-transinfected ARwP females, that we recently proved to be partially bidirectional [34]. Crosses between ARwP females and wild-type males were partially fertile when wild-type males were aged more than two weeks. On one hand, this weakness in the reproductive barrier between wild-type and transinfected mosquitoes may be convenient because it allows us to easily outcross the ARwP line with wild-type populations, thus restoring genetic variability periodically. However, the possible effects of this in field should be carefully considered.

First, we considered fundamental to provide additional data on wAlbA and wAlbB density in wild type Ae. albopictus from two sites for comparison with previous reports [40,41]. Next, we coupled the molecular determination of wAlbA and wAlbB bacterial titers in age-controlled superinfected males with the CI level observed in their crosses with ARwP females.

At last, comparison of laboratory results with the outcomes of Wolbachia density monitoring performed on captured wild-type males has been discussed as a necessary step for developing a bio-ecologically safe, long-term, and area-wide suppression strategy against Ae. albopictus based on the exploitation of Wolbachia-induced CI.

Materials and Methods

Ethics Statement

Research performed on invertebrates such as mosquitoes does not require a specific permit according to the directive 2010/63/EU of the European Parliament and of the Council on the protection of animals used for scientific purposes. Ae. albopictus is not an endangered or protected species, and no specific permissions are needed for collecting its eggs or adults in Italy. Samples were not collected from private or protected areas (see below for locations). Two of the authors (MC and RM) voluntarily used their arms for blood feeding during the experiments. According to the ethics committee of ENEA, this practice is not considered human experimentation.

Mosquito lines and rearing

This study included three Ae. albopictus populations: a wPip-transinfected population (ARwP) [32] and two naturally superinfected populations (S_CRE and S_ANG) obtained from eggs collected in North and Central Italy, respectively. The first collection site is an urban area in Crevalcore in Bologna province (CRE: 44°43′12.10″N, 11° 8′54.94″E) while the second collection site is in Anguillara Sabazia, a suburban area 25 km north of Rome (ANG: 42°5′29.32″N, 12°16′17.57″E). The three mosquito populations were reared as described below.

Larvae were brought to adulthood in 1.5-l larval trays, at a density of 5 larvae/ml. Larval food was provided as described in Sinkins et al. [42]. Adult mosquitoes were kept in 40 × 40 × 40-cm cages placed in a climatic chamber (T = 27 ± 2C°, RH = 70 ± 10%, L:D = 14:10 hours) and were fed with sucrose solution (10%) soaked on cotton.

Age- and sex-specific Wolbachia density in S_CRE and S_ANG mosquito lines

Adult males and females from the two natural Wolbachia-infected lines were isolated at the emergence and were pooled to be aged in the following six age groups (1–3, 4–6, 7–9, 10–15, 16–20, and >20 days). Males and females from each mosquito line and age group were analyzed by quantitative polymerase chain reaction (qPCR; see below) to evaluate the variation in the mean density of each Wolbachia strain with aging.

Crosses for CI strength assessment

Naturally infected males were crossed with 1-week-old ARwP females to determine eventual correlations between wAlbA and wAlbB densities and induced CI level. This study was planned regardless of the geographical origin; for this reason, only S_CRE males were used.

In all, 50 virgin Ae. albopictus males belonging to 3 different age groups (3 ± 1, 11 ± 1, and 19 ± 1 days) were singly placed in 20 × 10 × 5-cm mating-oviposition cages with a single 1-week old ARwP virgin female (ARwP ♀ × S_CRE ♂).

Because nuclear genes may be involved in generating incompatibility between populations [43], the ARwP strain was outcrossed for 5 generations with Wolbachia-cured S_CRE males before setting up the CI experiments.

After copulation, the males from incompatible crosses were stored in ethanol and frozen for subsequent qPCR to determine the titer of Wolbachia strains of naturally infected Ae. albopictus. Once mated, females were fed with blood on human arms and isolated. Eggs laid from each single female were collected on oviposition devices made of wet strips of crepe paper and were stored in an incubator (temperature, 27°C; RH, 90%) for 5 days. The percentage of hatched eggs was used to compare CI levels with those obtained using fertile control crosses. Females whose eggs did not hatch were dissected to determine whether their spermathecae were filled or not with spermatozoa and in the latter case were excluded from the analysis.

Concurrently, ARwP compatible crosses (ARwP ♀ × ARwP ♂) were set up using 1-week-old virgin females and males belonging to the three age groups mentioned above. Ten crosses were performed for males belonging to each age group to determine the background embryonic mortality unrelated to CI and to compute CI_corr index (see below). In addition, 20 1-week-old virgin females were mated with males aged 3 ± 1 days and were then fed with blood. After egg laying, qPCR analysis was performed to quantify wPip Wolbachia titer and to ascertain whether wPip density and fertility in ARwP females were correlated in the above laboratory conditions.

Wolbachia density in wild caught males

Wolbachia titer of randomly captured wild males was surveyed at collection sites of the two studied populations (Crevalcore and Anguillara Sabazia in September 2013 and July 2014, respectively). Samples were collected using manual aspirators and by catching males flying around human operators. These Wolbachia density data were used to evaluate the risk of CI failure in crosses between wild-type males and ARwP females that were accidentally released or locally established after hypothetical IIT-based suppression programs.

Wolbachia genotyping and qPCR

DNA purification and qPCR.

Total DNA was extracted from the whole body of a single Ae. albopictus mosquito by using ZR Tissue & Insect DNA Kit MicroPrep (Zymo) according to manufacturer's instructions. Strain-specific primers were used to amplify wsp. The wAlbA-wsp, wAlbB-wsp and wPip-wsp loci were amplified using previously described oligonucleotide primers pairs 328F/QArev2, 183F/QBrev2, wPF/wPR respectively, [34,40,44] to obtained 200-, 112- and 271-bp fragments, respectively.

Actin gene of Ae. albopictus was used as a nuclear reference and was amplified using primer pair actAlbqPCRsense (CCCACACAGTCCCCATCTAC) and actAlbqPCRantisense (CGAGTAGCCACGTTCAGTCA) to obtain a 119-bp amplification product.

Amplification reactions were performed using 20 μl of FluoCycle II SYBR Master Mix (Euroclone). Total DNA (2 μl) from each mosquito was used as a template for PCR and each reaction was performed in triplicate. PCR was performed in ABI Prism 7100 (Applied Biosystems) thermal cycler using the following amplification program: initial activation at 95°C for 5 min, followed by 40 cycles at 95°C for 15 s and 60°C for 1 min. Presence of specific amplification products was verified using dissociation curves.

Construction of plasmids for obtaining qPCR standard curves.

Specific DNA sequences encoding wAlbA-wsp, wAlbB-wsp, wPip-wsp and actin (quantitative reference) for qPCR amplification were cloned from the total DNA extracts. DNA fragments of wAlbA-wsp (382 bp) and wAlbB-wsp (501 bp) were amplified from the total DNA extracted from field-caught Ae. albopictus by using primer pairs 328F/691R and 183F/691R, respectively [44]. DNA extracted from Cx. pipiens was used as a template to amplify a wPip-wsp gene fragment (404 bp) with primers 183F and wPR. All the amplicons were cloned in pCR 2.1 vector plasmid (TA Cloning Kit, Invitrogen).

The amplified sequences were assembled to obtain the plasmids pBS-A-B-act (containing wAlbA-wsp, wAlbB-wsp and actin gene fragments) and pBS-Pip-act (containing wPip-wsp and actin gene fragments) by using the following procedure. Actin gene fragment was transferred from pCR 2.1 into BamHI-NotI sites of pBluescript II SK (+) vector to produce pBS-act plasmid. Next, wAlbB-wsp and wPip-wsp fragments were cloned from pCR 2.1 into NotI-SacI sites of the pBS-act plasmid to produce pBS-B-act and pBS-Pip-act plasmids. Finally, wAlbA-wsp fragment was cloned from pCR 2.1 into KpnI-XhoI sites of the pBS-B-act plasmid to produce pBS-A-B-act plasmid. All the obtained constructs were sequenced to assess the correct assembly and absence of unwanted sequence variations.

Statistical analysis and CI computation.

Direct correlation between Wolbachia density (wAlbA and wAlbB) and mosquito age was investigated in both the sexes by using Spearman correlation test.

To test the correlation between CI level and wAlbA and wAlbB bacterial loads in males, a graph was plotted for Wolbachia density in each male used in each single-pair crossing experiment against observed CI expression.

Values of Wolbachia (wAlbA and wAlbB) density in each male used in incompatible crosses were grouped in Wolbachia density classes. For each cluster of Wolbachia density, we associated mean CI values found in crosses involving the corresponding males.

If the data set met the assumptions of normality (Shapiro–Wilk test, P > 0.05), one-way analysis of variance (ANOVA) was performed, followed by Bonferroni–Dunn multiple comparisons test. If the data set did not meet the assumptions of normality, non-parametric Kruskal–Wallis test was performed to determine whether this clustering highlighted significant differences between Wolbachia density in males and mean CI expressed in crosses. Dunn's test was used for pairwise comparison of mean wAlbA and wAlbB densities in males and CI values.

CI expression was calculated using egg mortality observed in each single-pair incompatible cross (ARwP ♀ × S_CRE ♂) and was compared with the mean hatching rate (mean ± SEM) observed in single-pair compatible crosses (e.g., ARwP ♀ × ARwP ♂) by using CI_corr index [45]. This index does not overestimate the CI level caused by other mortality factors such as mean embryonic mortality observed in compatible crosses and male age effects that generally decrease fertility. The formula used for calculating CI_corr index is as follows: where UnE_CI is the proportion of eggs that did not hatch in crosses between different infection types and UnE_CC is the proportion of undeveloped embryos in compatible crosses. The value attributed to this last parameter was derived from ARwP compatible crosses performed using three classes of age-controlled males as described above.

All statistical analyses were performed using GraphPad Prism Software 6.0.1 version (GraphPad Software Inc., San Diego, CA, USA).

Results

Age- and sex-specific wAlbA and wAlbB densities

Ae. albopictus males showed remarkable individual variability in wAlbA density (Fig. 1A). In fact, wAlbA titers ranged from 0.0001–0.165 per actin copy number (wAlbA/actin). Complete loss of this Wolbachia strain was rarely observed in males (only 1 case out of 174); however, most males infected at low wAlbA densities (<0.001 wAlbA/actin) yielded negative results for standard PCR (S1 Fig.). Despite this high variability, we observed an evident reduction in the variation range with an increase in age. In fact, in young males (age, 1–3 days), wAlbA/actin ratios were widely different, ranging from <0.001 to >0.06 (mean ± SEM, 0.024 ± 0.005; N = 60). However, very low wAlbA levels (<0.001 wAlbA/actin) were detected more frequently in older males (age, >16 days), with a narrow range of variability (0.005 ± 0.002 wAlbA/actin; N = 35).

Download:

Fig 1. Age dependent variation of wAlbA density in Ae. albopictus males (a) and females (b) belonging to two Italian populations (S_ANG from Anguillara Sabazia, Rome: empty squares; S_CRE from Crevalcore, Bologna: full circles).

Data trend is displayed via a polynomial trend-line together with the associated function (dashed line for S_ANG, solid line for S_CRE).

https://doi.org/10.1371/journal.pone.0121813.g001

Despite the high individual variability, a significant negative correlation was observed between age of males and wAlbA density (Spearman r = −0.35; P ≤ 0.0001).

In general, S_CRE males were infected with lower mean wAlbA titer (0.010 ± 0.004 wAlbA/actin) than S_ANG males (0.020 ± 0.005 wAlbA/actin) irrespective of their age. However, this difference was not statically significant (ANOVA, F_(1,94) = 1.28; P > 0.05).

In contrast to the trend observed in males, wAlbA density was positively correlated with age in females (Spearman r = 0.92, P ≤ 0.0001; Fig. 1B). The highest average wAlbA titer was observed in S_ANG females (2.420 ± 1.005 wAlbA/actin) and not in S_CRE females (1.800 ± 0.680 wAlbA/actin); however, data were not sufficiently robust to highlight a statistically significant difference with respect to the geographical origin of the females (ANOVA, F_(1,45) = 0.15; P > 0.05).

We confirmed that wAlbB was more abundant than wAlbA (t-test, P < 0.0001; Fig. 2). On an average, wAlbB was 40–50- and 7–9-fold more concentrated than wAlbA in males and females, respectively. In Ae. albopictus males (Fig. 2A), mean wAlbB titer increased with age (Spearman r = 0.38, P = 0.002). On the other hand, wAlbB density increased in females aged 10–15 days (Spearman r = 0.69; P ≤ 0.0001) and declined in older females (age, 19–21 days; Fig. 2B). Like wAlbA, wAlbB was more abundant on an average in both the sexes of the S_ANG population (0.80 ± 0.10 wAlbB/actin in males; 19.28 ± 3.76 wAlbB/actin in females) than in those of the S_CRE population (0.52 ± 0.16 wAlbB/actin in males; 14.00 ± 2.81 wAlbB/actin in females). However, the differences were not statistically significant (ANOVA, F_(1,24) = 1.09; P > 0.05).

Download:

Fig 2. Age dependent variation of wAlbB density in Ae. albopictus males (a) and females (b) belonging to two Italian populations (S_ANG from Anguillara Sabazia, Rome: empty squares; S_CRE from Crevalcore, Bologna: full circles).

Data trend is displayed via a polynomial trend-line together with the associated function (dashed line for S_ANG, solid line for S_CRE).

https://doi.org/10.1371/journal.pone.0121813.g002

Effect of male wAlbA and wAlbB densities on CI expression

A general trend of significant decrease in the percentage of egg hatching related to male aging was observed in compatible ARwP ♀ × ARwP ♂ crosses by using 1-week-old females. In fact, mean percentages of egg hatching were 74.51 ± 4.89, 65.62 ± 3.22, and 49.10 ± 5.88 when males were 3 ± 1, 11 ± 1, and 19 ± 1 days old, respectively. Particularly, the oldest males were significantly less fertile than younger males (ANOVA, F_(3,36) = 9,419; P < 0.0001).

Results of qPCR showed that fertility in compatible crosses was not correlated to wPip density in ARwP females (Spearman r = 0.012; P = 0.958; Fig. 3). Findings allowed us to exclude wPip titer as a factor determining egg mortality under the tested experimental conditions.

Download:

Fig 3. Correlation between percentage of egg hatching in ARwP × ARwP compatible crosses and wPip Wolbachia density in females.

Data trend is displayed via a polynomial trend-line together with the associated function.

https://doi.org/10.1371/journal.pone.0121813.g003

Results of qPCR analysis relative to wAlbA and wAlbB densities in males from incompatible crosses are reported in Fig. 4 along with the observed CI levels (CI_corr index). Our results showed that induction of strong CI (CI_corr index ≅ 100) corresponded to males with wAlbA titers ranging from 0.01 to 0.11 wAlbA/actin (Fig. 4A). It is worth highlighting that almost all crosses showing incomplete CI (<80%) involved older males and some young males apparently harboring low wAlbA density (<0.001 wAlbA/actin) since emergence. Furthermore, old males with relatively high wAlbA titers (>0.01 wAlbA/actin) induced complete or approximately 100% CI.

Download:

Fig 4. Correlation between wAlbA (a) or wAlbB (b) Wolbachia density and expressed CI levels (CI_corr index) in Ae. albopictus males when crossed with ARwP females at three different ages (3 ± 1; 11 ± 1; 19 ± 1).

Data trend is displayed via a polynomial trend-line together with the associated function.

https://doi.org/10.1371/journal.pone.0121813.g004

Male wAlbA densities were clustered into three classes <0.001, 0.001−0.010, and >0.010 wAlbA/actin, and their related mean CI levels were calculated (Table 1). The average CI level (CI_corr = 68.06 ± 6.20) induced by males with lowest wAlbA density was significantly lower than that in other classes (Kruskal–Wallis and Dunn multicomparison test P < 0.05).

Download:

Table 1. Density of wAlbA Wolbachia and associated CI level expressed by Ae. albopictus males when crossed with ARwP females.

https://doi.org/10.1371/journal.pone.0121813.t001

In contrast, male wAlbB titer showed no apparent correlation with CI penetrance in crosses involving ARwP females (Fig. 4B). To evaluate the specific role of wAlbB as a CI inducer, we restricted the correlation analysis to cases in which wAlbA density was low (<0.001 wAlbA/actin). By following this method, no significant correlation was observed between wAlbB concentration and CI expression (Fig. 5).

Download:

Fig 5. Correlation between wAlbB density and expressed CI level (CI_corr index) in Ae. albopictus males characterized by very low wAlbA titer when crossed with ARwP females.

Data trend is displayed via a polynomial trend-line together with the associated function.

https://doi.org/10.1371/journal.pone.0121813.g005

Wolbachia density in wild caught males

Results of qPCR analysis of wild collected Ae. albopictus are reported in Fig. 6 based on the same wAlbA density classes defined in the previous experiment. In September 2013, approximately 50% males collected from Crevalcore were infected with very low wAlbA titers (0.0001−0.001 wAlbA/actin). The males belonging to the above density class decreased to 46.15% in samples collected in July 2014 in favor of males infected with higher wAlbA titers (>0.01 wAlbA/actin). Among Ae. albopictus males collected from Anguillara Sabazia, proportion of males with infection titers below the lowest Wolbachia density class (0.0001−0.001 wAlbA/actin) ranged between 36.36% in September 2013 and 30.77% in July 2014.

Download:

Fig 6. Density of wAlbA Wolbachia in wild caught Ae. albopictus males from two Italian sites (Crevalcore, CRE, Bologna; Anguillara Sabazia, ANG, Rome) and two periods (September 2013; July 2014).

Density values have been clustered in three density classes. The individuals belonging to each density class are reported as percentages of the whole amount. CI level is expected to decrease to about 68% in crosses between ARwP females and males with wAlbA density values <0.001 wAlbA/act.

https://doi.org/10.1371/journal.pone.0121813.g006

Overall, males infected with wAlbA titers compatible with the expected complete or almost complete CI (>0.001 wAlbA/actin) ranged from 50.00% to 69.23% depending on the site and period.

Discussion

Various strategies exploiting Wolbachia infection are currently being investigated for vector control [16]. Of these, Wolbachia-induced CI has been proposed as a method to produce functionally sterile males that can be released in field for vector suppression, in accordance with the principles of SIT [13,14]. Wolbachia transinfection techniques have been used to establish new laboratory lines of important vector species to obtain biological traits suitable for this purpose [46]. Specifically, CI relationships with wild-type populations, male competitiveness in comparison with wild-type males, and fitness parameters contributing to efficient mass rearing should be studied carefully.

Nearly five years after its generation, the wPip-transfected line of Ae. albopictus (ARwP) is presently being evaluated for developing the most appropriate CI-based suppression strategy against this mosquito species. In previous studies, we have ascertained that ARwP line shows some desirable traits of an ideal IIT effector against Ae. albopictus (full CI, male mating competitiveness and female fitness not significantly different from wild-type Ae. albopictus) [32,34,35]. However, we also found that although CI between ARwP males and wild-type females was always complete, naturally infected males were not equally strong CI inducers towards ARwP females [34]. Moreover, although wild-type males were coinfected with wAlbA and wAlbB Wolbachia strains, only wAlbA strain determined a pattern of complete bidirectional incompatibility with wPip-infected females. In addition, male aging seemed to be the factor responsible for the reduction in CI level [34]. In this study, we analyzed wAlbA and wAlbB titers in age-controlled males collected from two sites and the associated CI levels after crossing these males with ARwP females. In fact, establishment of a correlation between Wolbachia density and CI would help in evaluating the risk of bidirectional CI failure by sampling wild-type males before hypothetical field releases.

Results of qPCR supported the results of previous studies [40,41], confirming that wAlbA is constantly maintained at a lower density than wAlbB both in males and females. In addition, density patterns of both the Wolbachia strains were strongly dependent upon sex, with females showing the highest densities and males exhibiting a dramatic decrease in wAlbA titers with age. Although we clearly confirmed that adult age was a fundamental factor influencing the density of both the bacterial strains, we realized that in general, the density of Wolbachia strains in naturally infected Ae. albopictus was an unpredictable individual feature that was partially related to the geographical origin of the population [40] and to environmental conditions (temperature and food availability) in which the larvae developed [42].

Differences observed at the population level were not significant in our study and did not involve the general trend of infection dynamics but confirmed that bacterial load may vary locally and depending on the period of the year. In fact, larger differences between populations have been observed in previous studies investigating wAlbA and wAlbB densities in Ae. albopictus populations from Reunion island, Greece, and Corsica [40].

Correlation studies between CI and Wolbachia density have shown that incompatibility level is positively related to male wAlbA density. However, this correlation is not linear. In fact, only males with very low wAlbA titers (<0.0010 wAlbA/actin) induced significantly lower levels of CI while those with higher wAlbA density did not show CI weakening. These findings relative to wAlbA partially agree with the model proposed by Breeuwer and Werren [47], which states that decrease in CI penetrance in old males is directly proportional to Wolbachia density in the testes or sperm cysts in general [45,48–56]. Notably, a marked decrease in incompatibility level was previously observed when 10-day-old wAlbA mono-infected males were crossed with Wolbachia-free females [41].

In contrast, wAlbB, the most abundant Wolbachia strain in superinfected males, did not show any correlation with the observed CI in crosses with ARwP females. In fact, the observed decrease in the CI level countered the initial increase in wAlbB density in aging males, which was possibly mediated by a concurrent decrease in wAlbA density. Moreover, in males with very low wAlbA titers, CI expression did not change in response to changes in wAlbB density. These results agree with those of a previous study, which showed that crosses between ARwP females and wAlbB or wAlbA mono-infected males were approximately 20% fertile and completely unfertile, respectively [34].

Previous studies have shown that Wolbachia infected various host tissues but was mostly (at least 20 folds more) concentrated in Ae. albopictus gonads [57]. Particularly, in Koh Samui and Mauritius strains, mono-infected with wAlbA, Wolbachia cannot be detected in non-reproductive tissues [58]. In addition, wAlbA and wAlbB strains showed a highly similar tissue tropism [57,58]. Thus, although our data concerned total body molecular quantification, we can confirm that wAlbA plays the main role in determining the complete bidirectionality in CI pattern occurring between native and ARwP Ae. albopictus. Under these premises, ascertaining wAlbA titer in wild collected males is crucial because changing environmental conditions may cause Wolbachia density to differ significantly. In fact, analysis of males from Crevalcore and Anguillara sites showed very low wAlbA titer in almost half of all the males (on average 45% falling in <0.0001 wAlbA/actin density group), which was in accordance with previous field data [40]. This means that ARwP females could be fertile not only with ARwP males but also (at least partially) with some wild-type males, presumably gaining a reproductive advantage compared with wild-type females.

Therefore, the unintentional but repeated release of ARwP females during IIT should be approached with caution for a series of reasons.

First, the lack of a sexing system that guarantees the total absence of females in the released population increases the vulnerability of any autocidal application against vector mosquitoes because released females can blood feed and transmit diseases [13,15]. This vulnerability further increases when we apply the IIT strategy because accidentally released females can mate with the released males and reproduce. In the specific case of the ARwP strain, the weakness of the bi-CI, that we have highlighted in this work, must be considered as further factor affecting the IIT failsafe.

On the other hand, based on the modeling approach [39,59], presence of two reciprocally completely incompatible mosquito populations at one site is desirable because the competition decreases the number of biting females expected in the long-term for a single population. In addition, a recent paper reported about the stable coexistence in a site of two molecular Wolbachia strains of Culex pipiens, characterized, like ARwP and wild Ae. albopictus, by partially bidirectional CI relationships [60] However, the outcome of a partial failure in the bidirectionality of the CI pattern between ARwP and wild-type Ae. albopictus has not yet been properly investigated by experimental validation of specific theoretical models.

For the reasons above stated, awaiting for the advent of a very efficient sexing system, other strategies could be developed to exploit the favorable properties of the ARwP strain.

An advancement of the IIT-based suppression strategy could be the concurrent or sequential release of males belonging to two reciprocally incompatible lines. This approach could drastically reduce the possibility of establishing a transinfected line because of incompatible crosses that would occur in most cases [15]; however, this should be experimentally validated. Suitable Ae. albopictus lines are already available [61,62]; nevertheless, other lines can be specifically established for this purpose.

Furthermore, an alternative strategy using a combination of IIT and SIT involving irradiation of incompatible males at radiation doses just high enough to induce sterility in any females that are not removed but not sufficient to affect male fitness should be considered [16,63,64]. This approach is expected to be promising if, life most insect species, Ae. albopictus females are more radiosensitive than males [65]. An important advantage of this modified IIT strategy over the traditional SIT strategy would be the absence of any residual male fertility, necessarily tolerated by the latter strategy to save male mating competitiveness [66,67].

As long as a method to release large amounts of only males is available and/or specific population dynamic models are experimentally validated, the latter strategy seems to be the easiest to test and propose. Overall, the goal of field application of the ARwP line to control Ae. albopictus based on IIT/SIT approaches seems achievable. However, as shown in this study, a series of necessary steps need to be considered to ensure that the application strategy is effective and safe in the long-term.

Supporting Information

S1 Fig. Evaluation of PCR sensitivity in relation to increasing wAlbA titers, calculated as wAlbA/actin copy numbers by qPCR on independent mosquito extracts: (1) 0.0001; (2) 0.0010; (3) 0.0039; (4) 0.0108; (5) 0.0582; (6) 0.1671; (7) 2.2938; (8) no template PCR control.

For wspA gene amplification 328F and 691R oligonucleotides were used, according to the following PCR program: 95°C for 3 min; then 35 cycles at 95°C for 30 s, 55°C for 30 s and 72°C for 35 s; finally an elongation step at 72°C for 7 min. The expected wspA amplicon was of 382 bp.

https://doi.org/10.1371/journal.pone.0121813.s001

(TIF)

Acknowledgments

The authors are grateful to Eugenio Benvenuto who provided us with some of the qPCR analysis tools and Orsio Allegrucci for supporting in the maintenance of the insect colonies. We thank Marta Piscitelli, Elena Lampazzi and Francine Tankeu Nzufo for technical assistance. We also thank the anonymous reviewers for their valuable comments on the manuscript.

Author Contributions

Conceived and designed the experiments: MC. Performed the experiments: FM AD MC. Analyzed the data: MC AD FM. Contributed reagents/materials/analysis tools: AP. Wrote the paper: MC RM.

References

1. Gratz NG. Critical review of the vector status of Aedes albopictus. Med Vet Entomol. 2014; 18: 215–227.
- View Article
- Google Scholar
2. Benedict MQ, Levine RS, Hawley WA, Lounibos LP. Spread of the tiger: global risk of invasion by the mosquito Aedes albopictus. Vector Borne Zoonotic Dis. 2007; 7: 76–85. pmid:17417960
- View Article
- PubMed/NCBI
- Google Scholar
3. Mitchell CJ. Vector competence of North and South American strains of Aedes albopictus for certain arboviruses: a review. J Am Mosq Control Assoc. 1991; 7: 446–451. pmid:1791455
- View Article
- PubMed/NCBI
- Google Scholar
4. Effler P, Pang L, Kitsutani P, Vorndam V, Nakata M, Ayers T, et al. Dengue fever, Hawaii, 2001–2002. Emerg Infect Dis. 2005; 11: 742–749. pmid:15890132
- View Article
- PubMed/NCBI
- Google Scholar
5. Issack MI, Pursem VN, Barkham TMS, Ng LC, Inoue M, Manraj SS. Reemergence of dengue in Mauritius. Emerg Infect Dis. 2001; 16: 716–718.
- View Article
- Google Scholar
6. Xu G, Dong H, Shi N, Liu S, Zhou A, Cheng Z, et al. An outbreak of dengue virus serotype 1 infection in Cixi, Ningbo, People’s Republic of China, 2004, associated with a traveler from Thailand and high density of Aedes albopictus. Am J Trop Med Hyg. 2007; 76: 1182–1188. pmid:17556633
- View Article
- PubMed/NCBI
- Google Scholar
7. Tsetsarkin KA, Weaver SC. Sequential adaptive mutations enhance efficient vector switching by chikungunya virus and its epidemic emergence. PLOS Pathog. 2011; 7: e1002412. pmid:22174678
- View Article
- PubMed/NCBI
- Google Scholar
8. Josseran L, Paquet C, Zehgnoun A, Caillere N, Le Tertre A, Solet J-L, et al. Chikungunya disease outbreak, Reunion Island. Am J Trop Med Hyg. 2006; 12: 1994–1995.
- View Article
- Google Scholar
9. Rezza G, Nicoletti L, Angelini R, Romi R, Finarelli AC, Panning M, et al. Infection with chikungunya virus in Italy: an outbreak in a temperate region. Lancet. 2007; 370: 1840–1846. pmid:18061059
- View Article
- PubMed/NCBI
- Google Scholar
10. Grandadam M, Caro V, Plumet S, Thiberge JM, Souarès Y, Failloux AB, et al. Chikungunya virus, southeastern France. Emerg Infect Dis. 2011; 17: 910–913. pmid:21529410
- View Article
- PubMed/NCBI
- Google Scholar
11. Weaver SC. Arrival of chikungunya virus in the new world: prospects for spread and impact on public health. PLoS Negl Trop Dis. 2014; 8: e2921. pmid:24967777
- View Article
- PubMed/NCBI
- Google Scholar
12. Ruiz-Moreno D, Vargas IS, Olson KE, Harrington LC. Modeling dynamic introduction of chikungunya virus in the united states. PLOS Negl Trop Dis. 2012; 6(11): e1918. pmid:23209859
- View Article
- PubMed/NCBI
- Google Scholar
13. Dyck VA, Hendrichs J, Robinson AS. Sterile insect technique. Principles and practice in area-wide integrated pest management. Springer. Dordrecht, The Netherlands; 2005. https://doi.org/10.1007/1-4020-4051-2
14. Alphey L, Benedict M, Bellini R, Clark GG, Dame DA, Service MW, et al. Sterile-insect methods for control of mosquito-borne diseases: an analysis. Vector Borne Zoonotic Dis. 2010; 10: 295–311. pmid:19725763
- View Article
- PubMed/NCBI
- Google Scholar
15. Bourtzis K, Robinson AS. Insect pest control using Wolbachia and/or radiation. In: Bourtzis K, Miller TA, editors. Insect Symbiosis, Volume 2. Boca Raton, FL: CRC Press; 2006. pp. 225–246.
16. Bourtzis K, Dobson SL, Xi Z, Rasgon JL, Calvitti M, Moreira LA, et al. Harnessing mosquito-Wolbachia symbiosis for vector and disease control. Acta Trop. 2014; 132 Suppl: S150–S163. pmid:24252486
- View Article
- PubMed/NCBI
- Google Scholar
17. Oliva CF, Vreysen MJB, Dupé S, Lees RS, Gilles JRL, Gouagna LC, et al. Current status and future challenges for controlling malaria with the sterile insect technique: technical and social perspectives. Acta Trop. 2014; 132 Suppl: S130–S139. pmid:24295892
- View Article
- PubMed/NCBI
- Google Scholar
18. Ponlawat A, Scott JG, Harrington LC. Insecticide susceptibility of Aedes aegypti and Aedes albopictus in Central Africa. J Med Entomol. 2005; 42: 821–825. pmid:16363166
- View Article
- PubMed/NCBI
- Google Scholar
19. Kamgang B, Marcombe S, Chandre F, Nchoutpouen E, Nwane P, Etang J, et al. Insecticide susceptibility of Aedes aegypti and Aedes albopictus in Central Africa. Parasit Vectors. 2011; 4: 79. pmid:21575154
- View Article
- PubMed/NCBI
- Google Scholar
20. Floate KD, Kyei-Poku GK, Coghlin PC. Overview and relevance of Wolbachia bacteria in biocontrol research. Biocontrol Sci Technol. 2006; 16: 767–788.
- View Article
- Google Scholar
21. Zug R, Hammerstein P. Still a host of hosts for Wolbachia: Analysis of recent data suggests that 40% of terrestrial arthropod species are infected. PLOS ONE. 2012; 7(6): e38544. pmid:22685581
- View Article
- PubMed/NCBI
- Google Scholar
22. Werren JH, Baldo L, Clark ME. Wolbachia: master manipulators of invertebrate biology. Nat Rev Microbiol. 2008; 6: 741–751. pmid:18794912
- View Article
- PubMed/NCBI
- Google Scholar
23. Yen JH, Barr AR. New hypothesis of the cause of cytoplasmic incompatibility in Culex pipiens L. Nature. 1971; 232: 657–658. pmid:4937405
- View Article
- PubMed/NCBI
- Google Scholar
24. Werren JH. Biology of Wolbachia. Annu Rev Entomol. 1997; 42: 587–609. pmid:15012323
- View Article
- PubMed/NCBI
- Google Scholar
25. Bourtzis K, Dobson SL, Braig HR, O’Neill SL. Rescuing Wolbachia have been overlooked. Nature. 1998; 391: 852–853. pmid:9495337
- View Article
- PubMed/NCBI
- Google Scholar
26. Bourtzis K, Braig HR. The many faces of Wolbachia. In: Raoult D, Hackstadt T, editors. Rickettsiae and rickettsial diseases at the turn of the third millennium. Paris: Elsevier; 1999. pp.199–219.
27. Stouthamer R, Breeuwer JA, Hurst GD. Wolbachia pipientis: microbial manipulator of arthropod reproduction. Annu Rev Microbiol. 1999; 53: 71–102. pmid:10547686
- View Article
- PubMed/NCBI
- Google Scholar
28. Marshall JF, Staley J. Some notes regarding the morphological and biological differentiation of Culex pipiens Linnaeus and Culex molestus Forskàl (Diptera, Culicidae). Proc R Entomol Soc London Ser A, Gen Entomol. 2009; 12: 17–26.
- View Article
- Google Scholar
29. Ghelelovitch S. Genetic determinism of sterility in the cross-breeding of various strains of Culex autogenicus Roubaud. C R Hebd Seances Acad Sci. 1952; 234: 2386–2388. pmid:12979357
- View Article
- PubMed/NCBI
- Google Scholar
30. Laven H. Eradication of Culex pipiens fatigans through cytoplasmic incompatibility. Nature. 1967; 216: 383–384. pmid:4228275
- View Article
- PubMed/NCBI
- Google Scholar
31. Bellini R, Medici A, Puggioli A, Balestrino F, Carrieri M. Pilot field trials with Aedes albopictus irradiated sterile males in Italian urban areas. J Med Entomol. 2013; 50: 317–325. pmid:23540120
- View Article
- PubMed/NCBI
- Google Scholar
32. Calvitti M, Moretti R, Lampazzi E, Bellini R, Dobson SL. Characterization of a new Aedes albopictus (Diptera: Culicidae)-Wolbachia pipientis (Rickettsiales: Rickettsiaceae) symbiotic association generated by artificial transfer of the wPip strain from Culex pipiens (Diptera: Culicidae). J Med Entomol. 2010; 47: 179–187. pmid:20380298
- View Article
- PubMed/NCBI
- Google Scholar
33. Atyame CM, Labbé P, Dumas E, Milesi P, Charlat S, Fort P, et al. Wolbachia divergence and the evolution of cytoplasmic incompatibility in Culex pipiens. PLOS ONE. 2014; 9: e87336. pmid:24498078
- View Article
- PubMed/NCBI
- Google Scholar
34. Calvitti M, Moretti R, Skidmore AR, Dobson SL. Wolbachia strain wPip yields a pattern of cytoplasmic incompatibility enhancing a Wolbachia-based suppression strategy against the disease vector Aedes albopictus. Parasit Vectors. 2012; 5: 254. pmid:23146564
- View Article
- PubMed/NCBI
- Google Scholar
35. Moretti R, Calvitti M. Male mating performance and cytoplasmic incompatibility in a wPip Wolbachia trans-infected line of Aedes albopictus (Stegomyia albopicta). Med Vet Entomol. 2013; 27: 377–386. pmid:23171418
- View Article
- PubMed/NCBI
- Google Scholar
36. Zabalou S, Apostolaki A, Livadaras I, Franz G, Robinson AS, Savakis C, et al. Incompatible insect technique: incompatible males from a Ceratitis capitata genetic sexing strain. Entomol Exp Appl. 2009; 132: 232–240.
- View Article
- Google Scholar
37. O’Connor L, Plichart C, Sang AC, Brelsfoard CL, Bossin HC, Dobson SL. Open release of male mosquitoes infected with a Wolbachia biopesticide: field performance and infection containment. PLOS Negl Trop Dis. 2012; 6: e1797. pmid:23166845
- View Article
- PubMed/NCBI
- Google Scholar
38. Balestrino F, Puggioli A, Gilles JRL, Bellini R. Validation of a new larval rearing unit for Aedes albopictus (Diptera: Culicidae) mass rearing. PLOS ONE. 2014; 9: e91914. pmid:24647347
- View Article
- PubMed/NCBI
- Google Scholar
39. Dobson SL, Fox CW, Jiggins FM. The effect of Wolbachia-induced cytoplasmic incompatibility on host population size in natural and manipulated systems. Proc Biol Sci. 2002; 269: 437–445. pmid:11886634
- View Article
- PubMed/NCBI
- Google Scholar
40. Tortosa P, Charlat S, Labbé P, Dehecq JS, Barré H, Weill M. Wolbachia age-sex-specific density in Aedes albopictus: a host evolutionary response to cytoplasmic incompatibility? PLOS ONE. 2010; 5: e9700. pmid:20300514
- View Article
- PubMed/NCBI
- Google Scholar
41. Kittayapong P, Mongkalangoon P, Baimai V, O’Neill SL. Host age effect and expression of cytoplasmic incompatibility in field populations of Wolbachia-superinfected Aedes albopictus. Heredity. 2002; 88: 270–274. pmid:11920134
- View Article
- PubMed/NCBI
- Google Scholar
42. Sinkins SP, Braig HR, O’Neill SL. Wolbachia superinfections and the expression of cytoplasmic incompatibility. Proc Biol Sci. 1995; 261: 325–330. pmid:8587875
- View Article
- PubMed/NCBI
- Google Scholar
43. Weeks AR, Reynolds TK, Hoffmann AA. Wolbachia dynamics and host effects: what has (and has not) been demonstrated? Trends Ecol Evol. 2002; 17: 257–262.
- View Article
- Google Scholar
44. Zhou W, Rousset F, O’Neil S. Phylogeny and PCR-based classification of Wolbachia strains using wsp gene sequences. Proc Biol Sci. 1998; 265: 509–515. pmid:9569669
- View Article
- PubMed/NCBI
- Google Scholar
45. Poinsot D, Bourtzis K, Markakis G, Savakis C, Merçot H. Wolbachia transfer from Drosophila melanogaster into D. simulans: Host effect and cytoplasmic incompatibility relationships. PLOS ONE. 1998; 150: 227–237.
- View Article
- Google Scholar
46. Hughes GL, Rasgon JL. Transinfection: a method to investigate Wolbachia-host interactions and control arthropod-borne disease. Insect Mol Biol. 2014; 23: 141–151. pmid:24329998
- View Article
- PubMed/NCBI
- Google Scholar
47. Breeuwer JA, Werren JH. Cytoplasmic incompatibility and bacterial density in Nasonia vitripennis. Genetics. 1993; 135: 565–574. pmid:8244014
- View Article
- PubMed/NCBI
- Google Scholar
48. Hoffmann AA, Turelli M, Simmons GM. Unidirectional incompatibility between populations of Drosophila simulans. Evolution. 1986; 40: 692–701.
- View Article
- Google Scholar
49. Binnington KC, Hoffmann AA. Wolbachia-like organisms and cytoplasmic incompatibility in Drosophila simulans. J Invertebr Pathol. 1989; 54: 344–352.
- View Article
- Google Scholar
50. Bressac C, Rousset F. The reproductive incompatibility system in Drosophila simulans: DAPI-staining analysis of the Wolbachia symbionts in sperm cysts. J Invertebr Pathol. 1993; 61: 226–230. pmid:7689622
- View Article
- PubMed/NCBI
- Google Scholar
51. Turelli M, Hoffmann AA. Cytoplasmic incompatibility in Drosophila simulans: dynamics and parameter estimates from natural populations. Genetics. 1995; 140: 1319–1338. pmid:7498773
- View Article
- PubMed/NCBI
- Google Scholar
52. Bourtzis K, Nirgianaki A, Markakis G, Savakis C. Wolbachia infection and cytoplasmic incompatibility in Drosophila species. Genetics. 1996; 144: 1063–1073. pmid:8913750
- View Article
- PubMed/NCBI
- Google Scholar
53. Jamnongluk W, Kittayapong P, Baisley KJ, O’Neill SL. Wolbachia infection and expression of cytoplasmic incompatibility in Armigeres subalbatus (Diptera: Culicidae). J Med Entomol. 2000; 37: 53–57. pmid:15218907
- View Article
- PubMed/NCBI
- Google Scholar
54. Noda H, Koizumi Y, Zhang Q, Deng K. Infection density of Wolbachia and incompatibility level in two planthopper species, Laodelphax striatellus and Sogatella furcifera. Insect Biochem Mol Biol. 2001; 31: 727–737. pmid:11267910
- View Article
- PubMed/NCBI
- Google Scholar
55. Veneti Z, Clark ME, Zabalou S, Karr TL, Savakis C, Bourtzis K. Cytoplasmic incompatibility and sperm cyst infection in different Drosophila-Wolbachia associations. Genetics. 2003; 164: 545–552. pmid:12807775
- View Article
- PubMed/NCBI
- Google Scholar
56. Veneti Z, Clark ME, Karr TL, Savakis C, Bourtzis K. Heads or tails: host-parasite interactions in the Drosophila-Wolbachia system. Appl Environ Microbiol. 2004; 70: 5366–5372. pmid:15345422
- View Article
- PubMed/NCBI
- Google Scholar
57. Zouache K, Voronin D, Tran-Van V, Mousson L, Failloux AB, Mavingui P. Persistent Wolbachia and cultivable bacteria infection in the reproductive and somatic tissues of the mosquito vector Aedes albopictus. PLOS ONE. 2009; 4(7): e6388. pmid:19633721
- View Article
- PubMed/NCBI
- Google Scholar
58. Dobson SL, Bourtzis K, Braig HR, Jones BF, Zhou W, Rousset F, et al. Wolbachia infections are distributed throughout insect somatic and germ line tissues. Insect Biochem Mol Biol. 1999; 29: 153–160. pmid:10196738
- View Article
- PubMed/NCBI
- Google Scholar
59. Dobson SL, Marsland EJ, Rattanadechakul W. Mutualistic Wolbachia infection in Aedes albopictus: accelerating cytoplasmic drive. Genetics. 2002; 160: 1087–1094. pmid:11901124
- View Article
- PubMed/NCBI
- Google Scholar
60. Atyame CM, Labbé P, Rousset F, Beji M, Makoundou P, Duron O, et al. Stable coexistence of incompatible Wolbachia along a narrow contact zone in mosquito field populations. Molecular Ecology. 2015; 24(2): 508–21 pmid:25482270
- View Article
- PubMed/NCBI
- Google Scholar
61. Xi Z, Khoo CCH, Dobson SL. Interspecific transfer of Wolbachia into the mosquito disease vector Aedes albopictus. Proc R Soc B. 2006; 273: 1317–1322. pmid:16777718
- View Article
- PubMed/NCBI
- Google Scholar
62. Blagrove MSC, Arias-Goeta C, Di Genua C, Failloux AB, Sinkins SP. A Wolbachia wMel transinfection in Aedes albopictus is not detrimental to host fitness and inhibits chikungunya virus. PLoS Negl Trop Dis. 2013; 7(3): e2152. pmid:23556030
- View Article
- PubMed/NCBI
- Google Scholar
63. Curtis CF. Testing systems for the genetic control of mosquitoes. Proceeding XV International Congress of Entomology. Washington DC; 1976. pp. 106–116.
64. Brelsfoard CL, St Clair W, Dobson SL. Integration of irradiation with cytoplasmic incompatibility to facilitate a lymphatic filariasis vector elimination approach. Parasit Vectors. 2009; 2: 38. pmid:19682363
- View Article
- PubMed/NCBI
- Google Scholar
65. Bakri A, Mehta K, Lance DR. Sterilizing insects with ionizing radiation. In: Dyck VA, Hendrichs J, Robinson AS, editors. Sterile Insect Technique. Principles and practice in area-wide integrated pest management. Dordrecht, The Netherlands: Springer; 2005. pp. 233–268.
66. Oliva CF, Jacquet M, Gilles J, Lemperiere G, Maquart PO, Quilici S, et al. The sterile insect technique for controlling populations of Aedes albopictus (Diptera: Culicidae) on Reunion Island: mating vigour of sterilized males. PLOS ONE. 2012; 7: e49414. pmid:23185329
- View Article
- PubMed/NCBI
- Google Scholar
67. Balestrino F, Medici A, Candini G, Carrieri M, Maccagnani B, Calvitti M, et al. Gamma ray dosimetry and mating capacity studies in the laboratory on Aedes albopictus males. J Med Entomol. 2010; 47: 581–591. pmid:20695273
- View Article
- PubMed/NCBI
- Google Scholar

[ref1] 1. Gratz NG. Critical review of the vector status of Aedes albopictus. Med Vet Entomol. 2014; 18: 215–227.
View Article
Google Scholar

[2] View Article

[3] Google Scholar

[ref2] 2. Benedict MQ, Levine RS, Hawley WA, Lounibos LP. Spread of the tiger: global risk of invasion by the mosquito Aedes albopictus. Vector Borne Zoonotic Dis. 2007; 7: 76–85. pmid:17417960
View Article
PubMed/NCBI
Google Scholar

[5] View Article

[6] PubMed/NCBI

[7] Google Scholar

[ref3] 3. Mitchell CJ. Vector competence of North and South American strains of Aedes albopictus for certain arboviruses: a review. J Am Mosq Control Assoc. 1991; 7: 446–451. pmid:1791455
View Article
PubMed/NCBI
Google Scholar

[9] View Article

[10] PubMed/NCBI

[11] Google Scholar

[ref4] 4. Effler P, Pang L, Kitsutani P, Vorndam V, Nakata M, Ayers T, et al. Dengue fever, Hawaii, 2001–2002. Emerg Infect Dis. 2005; 11: 742–749. pmid:15890132
View Article
PubMed/NCBI
Google Scholar

[13] View Article

[14] PubMed/NCBI

[15] Google Scholar

[ref5] 5. Issack MI, Pursem VN, Barkham TMS, Ng LC, Inoue M, Manraj SS. Reemergence of dengue in Mauritius. Emerg Infect Dis. 2001; 16: 716–718.
View Article
Google Scholar

[17] View Article

[18] Google Scholar

[ref6] 6. Xu G, Dong H, Shi N, Liu S, Zhou A, Cheng Z, et al. An outbreak of dengue virus serotype 1 infection in Cixi, Ningbo, People’s Republic of China, 2004, associated with a traveler from Thailand and high density of Aedes albopictus. Am J Trop Med Hyg. 2007; 76: 1182–1188. pmid:17556633
View Article
PubMed/NCBI
Google Scholar

[20] View Article

[21] PubMed/NCBI

[22] Google Scholar

[ref7] 7. Tsetsarkin KA, Weaver SC. Sequential adaptive mutations enhance efficient vector switching by chikungunya virus and its epidemic emergence. PLOS Pathog. 2011; 7: e1002412. pmid:22174678
View Article
PubMed/NCBI
Google Scholar

[24] View Article

[25] PubMed/NCBI

[26] Google Scholar

[ref8] 8. Josseran L, Paquet C, Zehgnoun A, Caillere N, Le Tertre A, Solet J-L, et al. Chikungunya disease outbreak, Reunion Island. Am J Trop Med Hyg. 2006; 12: 1994–1995.
View Article
Google Scholar

[28] View Article

[29] Google Scholar

[ref9] 9. Rezza G, Nicoletti L, Angelini R, Romi R, Finarelli AC, Panning M, et al. Infection with chikungunya virus in Italy: an outbreak in a temperate region. Lancet. 2007; 370: 1840–1846. pmid:18061059
View Article
PubMed/NCBI
Google Scholar

[31] View Article

[32] PubMed/NCBI

[33] Google Scholar

[ref10] 10. Grandadam M, Caro V, Plumet S, Thiberge JM, Souarès Y, Failloux AB, et al. Chikungunya virus, southeastern France. Emerg Infect Dis. 2011; 17: 910–913. pmid:21529410
View Article
PubMed/NCBI
Google Scholar

[35] View Article

[36] PubMed/NCBI

[37] Google Scholar

[ref11] 11. Weaver SC. Arrival of chikungunya virus in the new world: prospects for spread and impact on public health. PLoS Negl Trop Dis. 2014; 8: e2921. pmid:24967777
View Article
PubMed/NCBI
Google Scholar

[39] View Article

[40] PubMed/NCBI

[41] Google Scholar

[ref12] 12. Ruiz-Moreno D, Vargas IS, Olson KE, Harrington LC. Modeling dynamic introduction of chikungunya virus in the united states. PLOS Negl Trop Dis. 2012; 6(11): e1918. pmid:23209859
View Article
PubMed/NCBI
Google Scholar

[43] View Article

[44] PubMed/NCBI

[45] Google Scholar

[ref13] 13. Dyck VA, Hendrichs J, Robinson AS. Sterile insect technique. Principles and practice in area-wide integrated pest management. Springer. Dordrecht, The Netherlands; 2005. https://doi.org/10.1007/1-4020-4051-2

[ref14] 14. Alphey L, Benedict M, Bellini R, Clark GG, Dame DA, Service MW, et al. Sterile-insect methods for control of mosquito-borne diseases: an analysis. Vector Borne Zoonotic Dis. 2010; 10: 295–311. pmid:19725763
View Article
PubMed/NCBI
Google Scholar

[48] View Article

[49] PubMed/NCBI

[50] Google Scholar

[ref15] 15. Bourtzis K, Robinson AS. Insect pest control using Wolbachia and/or radiation. In: Bourtzis K, Miller TA, editors. Insect Symbiosis, Volume 2. Boca Raton, FL: CRC Press; 2006. pp. 225–246.

[ref16] 16. Bourtzis K, Dobson SL, Xi Z, Rasgon JL, Calvitti M, Moreira LA, et al. Harnessing mosquito-Wolbachia symbiosis for vector and disease control. Acta Trop. 2014; 132 Suppl: S150–S163. pmid:24252486
View Article
PubMed/NCBI
Google Scholar

[53] View Article

[54] PubMed/NCBI

[55] Google Scholar

[ref17] 17. Oliva CF, Vreysen MJB, Dupé S, Lees RS, Gilles JRL, Gouagna LC, et al. Current status and future challenges for controlling malaria with the sterile insect technique: technical and social perspectives. Acta Trop. 2014; 132 Suppl: S130–S139. pmid:24295892
View Article
PubMed/NCBI
Google Scholar

[57] View Article

[58] PubMed/NCBI

[59] Google Scholar

[ref18] 18. Ponlawat A, Scott JG, Harrington LC. Insecticide susceptibility of Aedes aegypti and Aedes albopictus in Central Africa. J Med Entomol. 2005; 42: 821–825. pmid:16363166
View Article
PubMed/NCBI
Google Scholar

[61] View Article

[62] PubMed/NCBI

[63] Google Scholar

[ref19] 19. Kamgang B, Marcombe S, Chandre F, Nchoutpouen E, Nwane P, Etang J, et al. Insecticide susceptibility of Aedes aegypti and Aedes albopictus in Central Africa. Parasit Vectors. 2011; 4: 79. pmid:21575154
View Article
PubMed/NCBI
Google Scholar

[65] View Article

[66] PubMed/NCBI

[67] Google Scholar

[ref20] 20. Floate KD, Kyei-Poku GK, Coghlin PC. Overview and relevance of Wolbachia bacteria in biocontrol research. Biocontrol Sci Technol. 2006; 16: 767–788.
View Article
Google Scholar

[69] View Article

[70] Google Scholar

[ref21] 21. Zug R, Hammerstein P. Still a host of hosts for Wolbachia: Analysis of recent data suggests that 40% of terrestrial arthropod species are infected. PLOS ONE. 2012; 7(6): e38544. pmid:22685581
View Article
PubMed/NCBI
Google Scholar

[72] View Article

[73] PubMed/NCBI

[74] Google Scholar

[ref22] 22. Werren JH, Baldo L, Clark ME. Wolbachia: master manipulators of invertebrate biology. Nat Rev Microbiol. 2008; 6: 741–751. pmid:18794912
View Article
PubMed/NCBI
Google Scholar

[76] View Article

[77] PubMed/NCBI

[78] Google Scholar

[ref23] 23. Yen JH, Barr AR. New hypothesis of the cause of cytoplasmic incompatibility in Culex pipiens L. Nature. 1971; 232: 657–658. pmid:4937405
View Article
PubMed/NCBI
Google Scholar

[80] View Article

[81] PubMed/NCBI

[82] Google Scholar

[ref24] 24. Werren JH. Biology of Wolbachia. Annu Rev Entomol. 1997; 42: 587–609. pmid:15012323
View Article
PubMed/NCBI
Google Scholar

[84] View Article

[85] PubMed/NCBI

[86] Google Scholar

[ref25] 25. Bourtzis K, Dobson SL, Braig HR, O’Neill SL. Rescuing Wolbachia have been overlooked. Nature. 1998; 391: 852–853. pmid:9495337
View Article
PubMed/NCBI
Google Scholar

[88] View Article

[89] PubMed/NCBI

[90] Google Scholar

[ref26] 26. Bourtzis K, Braig HR. The many faces of Wolbachia. In: Raoult D, Hackstadt T, editors. Rickettsiae and rickettsial diseases at the turn of the third millennium. Paris: Elsevier; 1999. pp.199–219.

[ref27] 27. Stouthamer R, Breeuwer JA, Hurst GD. Wolbachia pipientis: microbial manipulator of arthropod reproduction. Annu Rev Microbiol. 1999; 53: 71–102. pmid:10547686
View Article
PubMed/NCBI
Google Scholar

[93] View Article

[94] PubMed/NCBI

[95] Google Scholar

[ref28] 28. Marshall JF, Staley J. Some notes regarding the morphological and biological differentiation of Culex pipiens Linnaeus and Culex molestus Forskàl (Diptera, Culicidae). Proc R Entomol Soc London Ser A, Gen Entomol. 2009; 12: 17–26.
View Article
Google Scholar

[97] View Article

[98] Google Scholar

[ref29] 29. Ghelelovitch S. Genetic determinism of sterility in the cross-breeding of various strains of Culex autogenicus Roubaud. C R Hebd Seances Acad Sci. 1952; 234: 2386–2388. pmid:12979357
View Article
PubMed/NCBI
Google Scholar

[100] View Article

[101] PubMed/NCBI

[102] Google Scholar

[ref30] 30. Laven H. Eradication of Culex pipiens fatigans through cytoplasmic incompatibility. Nature. 1967; 216: 383–384. pmid:4228275
View Article
PubMed/NCBI
Google Scholar

[104] View Article

[105] PubMed/NCBI

[106] Google Scholar

[ref31] 31. Bellini R, Medici A, Puggioli A, Balestrino F, Carrieri M. Pilot field trials with Aedes albopictus irradiated sterile males in Italian urban areas. J Med Entomol. 2013; 50: 317–325. pmid:23540120
View Article
PubMed/NCBI
Google Scholar

[108] View Article

[109] PubMed/NCBI

[110] Google Scholar

[ref32] 32. Calvitti M, Moretti R, Lampazzi E, Bellini R, Dobson SL. Characterization of a new Aedes albopictus (Diptera: Culicidae)-Wolbachia pipientis (Rickettsiales: Rickettsiaceae) symbiotic association generated by artificial transfer of the wPip strain from Culex pipiens (Diptera: Culicidae). J Med Entomol. 2010; 47: 179–187. pmid:20380298
View Article
PubMed/NCBI
Google Scholar

[112] View Article

[113] PubMed/NCBI

[114] Google Scholar

[ref33] 33. Atyame CM, Labbé P, Dumas E, Milesi P, Charlat S, Fort P, et al. Wolbachia divergence and the evolution of cytoplasmic incompatibility in Culex pipiens. PLOS ONE. 2014; 9: e87336. pmid:24498078
View Article
PubMed/NCBI
Google Scholar

[116] View Article

[117] PubMed/NCBI

[118] Google Scholar

[ref34] 34. Calvitti M, Moretti R, Skidmore AR, Dobson SL. Wolbachia strain wPip yields a pattern of cytoplasmic incompatibility enhancing a Wolbachia-based suppression strategy against the disease vector Aedes albopictus. Parasit Vectors. 2012; 5: 254. pmid:23146564
View Article
PubMed/NCBI
Google Scholar

[120] View Article

[121] PubMed/NCBI

[122] Google Scholar

[ref35] 35. Moretti R, Calvitti M. Male mating performance and cytoplasmic incompatibility in a wPip Wolbachia trans-infected line of Aedes albopictus (Stegomyia albopicta). Med Vet Entomol. 2013; 27: 377–386. pmid:23171418
View Article
PubMed/NCBI
Google Scholar

[124] View Article

[125] PubMed/NCBI

[126] Google Scholar

[ref36] 36. Zabalou S, Apostolaki A, Livadaras I, Franz G, Robinson AS, Savakis C, et al. Incompatible insect technique: incompatible males from a Ceratitis capitata genetic sexing strain. Entomol Exp Appl. 2009; 132: 232–240.
View Article
Google Scholar

[128] View Article

[129] Google Scholar

[ref37] 37. O’Connor L, Plichart C, Sang AC, Brelsfoard CL, Bossin HC, Dobson SL. Open release of male mosquitoes infected with a Wolbachia biopesticide: field performance and infection containment. PLOS Negl Trop Dis. 2012; 6: e1797. pmid:23166845
View Article
PubMed/NCBI
Google Scholar

[131] View Article

[132] PubMed/NCBI

[133] Google Scholar

[ref38] 38. Balestrino F, Puggioli A, Gilles JRL, Bellini R. Validation of a new larval rearing unit for Aedes albopictus (Diptera: Culicidae) mass rearing. PLOS ONE. 2014; 9: e91914. pmid:24647347
View Article
PubMed/NCBI
Google Scholar

[135] View Article

[136] PubMed/NCBI

[137] Google Scholar

[ref39] 39. Dobson SL, Fox CW, Jiggins FM. The effect of Wolbachia-induced cytoplasmic incompatibility on host population size in natural and manipulated systems. Proc Biol Sci. 2002; 269: 437–445. pmid:11886634
View Article
PubMed/NCBI
Google Scholar

[139] View Article

[140] PubMed/NCBI

[141] Google Scholar

[ref40] 40. Tortosa P, Charlat S, Labbé P, Dehecq JS, Barré H, Weill M. Wolbachia age-sex-specific density in Aedes albopictus: a host evolutionary response to cytoplasmic incompatibility? PLOS ONE. 2010; 5: e9700. pmid:20300514
View Article
PubMed/NCBI
Google Scholar

[143] View Article

[144] PubMed/NCBI

[145] Google Scholar

[ref41] 41. Kittayapong P, Mongkalangoon P, Baimai V, O’Neill SL. Host age effect and expression of cytoplasmic incompatibility in field populations of Wolbachia-superinfected Aedes albopictus. Heredity. 2002; 88: 270–274. pmid:11920134
View Article
PubMed/NCBI
Google Scholar

[147] View Article

[148] PubMed/NCBI

[149] Google Scholar

[ref42] 42. Sinkins SP, Braig HR, O’Neill SL. Wolbachia superinfections and the expression of cytoplasmic incompatibility. Proc Biol Sci. 1995; 261: 325–330. pmid:8587875
View Article
PubMed/NCBI
Google Scholar

[151] View Article

[152] PubMed/NCBI

[153] Google Scholar

[ref43] 43. Weeks AR, Reynolds TK, Hoffmann AA. Wolbachia dynamics and host effects: what has (and has not) been demonstrated? Trends Ecol Evol. 2002; 17: 257–262.
View Article
Google Scholar

[155] View Article

[156] Google Scholar

[ref44] 44. Zhou W, Rousset F, O’Neil S. Phylogeny and PCR-based classification of Wolbachia strains using wsp gene sequences. Proc Biol Sci. 1998; 265: 509–515. pmid:9569669
View Article
PubMed/NCBI
Google Scholar

[158] View Article

[159] PubMed/NCBI

[160] Google Scholar

[ref45] 45. Poinsot D, Bourtzis K, Markakis G, Savakis C, Merçot H. Wolbachia transfer from Drosophila melanogaster into D. simulans: Host effect and cytoplasmic incompatibility relationships. PLOS ONE. 1998; 150: 227–237.
View Article
Google Scholar

[162] View Article

[163] Google Scholar

[ref46] 46. Hughes GL, Rasgon JL. Transinfection: a method to investigate Wolbachia-host interactions and control arthropod-borne disease. Insect Mol Biol. 2014; 23: 141–151. pmid:24329998
View Article
PubMed/NCBI
Google Scholar

[165] View Article

[166] PubMed/NCBI

[167] Google Scholar

[ref47] 47. Breeuwer JA, Werren JH. Cytoplasmic incompatibility and bacterial density in Nasonia vitripennis. Genetics. 1993; 135: 565–574. pmid:8244014
View Article
PubMed/NCBI
Google Scholar

[169] View Article

[170] PubMed/NCBI

[171] Google Scholar

[ref48] 48. Hoffmann AA, Turelli M, Simmons GM. Unidirectional incompatibility between populations of Drosophila simulans. Evolution. 1986; 40: 692–701.
View Article
Google Scholar

[173] View Article

[174] Google Scholar

[ref49] 49. Binnington KC, Hoffmann AA. Wolbachia-like organisms and cytoplasmic incompatibility in Drosophila simulans. J Invertebr Pathol. 1989; 54: 344–352.
View Article
Google Scholar

[176] View Article

[177] Google Scholar

[ref50] 50. Bressac C, Rousset F. The reproductive incompatibility system in Drosophila simulans: DAPI-staining analysis of the Wolbachia symbionts in sperm cysts. J Invertebr Pathol. 1993; 61: 226–230. pmid:7689622
View Article
PubMed/NCBI
Google Scholar

[179] View Article

[180] PubMed/NCBI

[181] Google Scholar

[ref51] 51. Turelli M, Hoffmann AA. Cytoplasmic incompatibility in Drosophila simulans: dynamics and parameter estimates from natural populations. Genetics. 1995; 140: 1319–1338. pmid:7498773
View Article
PubMed/NCBI
Google Scholar

[183] View Article

[184] PubMed/NCBI

[185] Google Scholar

[ref52] 52. Bourtzis K, Nirgianaki A, Markakis G, Savakis C. Wolbachia infection and cytoplasmic incompatibility in Drosophila species. Genetics. 1996; 144: 1063–1073. pmid:8913750
View Article
PubMed/NCBI
Google Scholar

[187] View Article

[188] PubMed/NCBI

[189] Google Scholar

[ref53] 53. Jamnongluk W, Kittayapong P, Baisley KJ, O’Neill SL. Wolbachia infection and expression of cytoplasmic incompatibility in Armigeres subalbatus (Diptera: Culicidae). J Med Entomol. 2000; 37: 53–57. pmid:15218907
View Article
PubMed/NCBI
Google Scholar

[191] View Article

[192] PubMed/NCBI

[193] Google Scholar

[ref54] 54. Noda H, Koizumi Y, Zhang Q, Deng K. Infection density of Wolbachia and incompatibility level in two planthopper species, Laodelphax striatellus and Sogatella furcifera. Insect Biochem Mol Biol. 2001; 31: 727–737. pmid:11267910
View Article
PubMed/NCBI
Google Scholar

[195] View Article

[196] PubMed/NCBI

[197] Google Scholar

[ref55] 55. Veneti Z, Clark ME, Zabalou S, Karr TL, Savakis C, Bourtzis K. Cytoplasmic incompatibility and sperm cyst infection in different Drosophila-Wolbachia associations. Genetics. 2003; 164: 545–552. pmid:12807775
View Article
PubMed/NCBI
Google Scholar

[199] View Article

[200] PubMed/NCBI

[201] Google Scholar

[ref56] 56. Veneti Z, Clark ME, Karr TL, Savakis C, Bourtzis K. Heads or tails: host-parasite interactions in the Drosophila-Wolbachia system. Appl Environ Microbiol. 2004; 70: 5366–5372. pmid:15345422
View Article
PubMed/NCBI
Google Scholar

[203] View Article

[204] PubMed/NCBI

[205] Google Scholar

[ref57] 57. Zouache K, Voronin D, Tran-Van V, Mousson L, Failloux AB, Mavingui P. Persistent Wolbachia and cultivable bacteria infection in the reproductive and somatic tissues of the mosquito vector Aedes albopictus. PLOS ONE. 2009; 4(7): e6388. pmid:19633721
View Article
PubMed/NCBI
Google Scholar

[207] View Article

[208] PubMed/NCBI

[209] Google Scholar

[ref58] 58. Dobson SL, Bourtzis K, Braig HR, Jones BF, Zhou W, Rousset F, et al. Wolbachia infections are distributed throughout insect somatic and germ line tissues. Insect Biochem Mol Biol. 1999; 29: 153–160. pmid:10196738
View Article
PubMed/NCBI
Google Scholar

[211] View Article

[212] PubMed/NCBI

[213] Google Scholar

[ref59] 59. Dobson SL, Marsland EJ, Rattanadechakul W. Mutualistic Wolbachia infection in Aedes albopictus: accelerating cytoplasmic drive. Genetics. 2002; 160: 1087–1094. pmid:11901124
View Article
PubMed/NCBI
Google Scholar

[215] View Article

[216] PubMed/NCBI

[217] Google Scholar

[ref60] 60. Atyame CM, Labbé P, Rousset F, Beji M, Makoundou P, Duron O, et al. Stable coexistence of incompatible Wolbachia along a narrow contact zone in mosquito field populations. Molecular Ecology. 2015; 24(2): 508–21 pmid:25482270
View Article
PubMed/NCBI
Google Scholar

[219] View Article

[220] PubMed/NCBI

[221] Google Scholar

[ref61] 61. Xi Z, Khoo CCH, Dobson SL. Interspecific transfer of Wolbachia into the mosquito disease vector Aedes albopictus. Proc R Soc B. 2006; 273: 1317–1322. pmid:16777718
View Article
PubMed/NCBI
Google Scholar

[223] View Article

[224] PubMed/NCBI

[225] Google Scholar

[ref62] 62. Blagrove MSC, Arias-Goeta C, Di Genua C, Failloux AB, Sinkins SP. A Wolbachia wMel transinfection in Aedes albopictus is not detrimental to host fitness and inhibits chikungunya virus. PLoS Negl Trop Dis. 2013; 7(3): e2152. pmid:23556030
View Article
PubMed/NCBI
Google Scholar

[227] View Article

[228] PubMed/NCBI

[229] Google Scholar

[ref63] 63. Curtis CF. Testing systems for the genetic control of mosquitoes. Proceeding XV International Congress of Entomology. Washington DC; 1976. pp. 106–116.

[ref64] 64. Brelsfoard CL, St Clair W, Dobson SL. Integration of irradiation with cytoplasmic incompatibility to facilitate a lymphatic filariasis vector elimination approach. Parasit Vectors. 2009; 2: 38. pmid:19682363
View Article
PubMed/NCBI
Google Scholar

[232] View Article

[233] PubMed/NCBI

[234] Google Scholar

[ref65] 65. Bakri A, Mehta K, Lance DR. Sterilizing insects with ionizing radiation. In: Dyck VA, Hendrichs J, Robinson AS, editors. Sterile Insect Technique. Principles and practice in area-wide integrated pest management. Dordrecht, The Netherlands: Springer; 2005. pp. 233–268.

[ref66] 66. Oliva CF, Jacquet M, Gilles J, Lemperiere G, Maquart PO, Quilici S, et al. The sterile insect technique for controlling populations of Aedes albopictus (Diptera: Culicidae) on Reunion Island: mating vigour of sterilized males. PLOS ONE. 2012; 7: e49414. pmid:23185329
View Article
PubMed/NCBI
Google Scholar

[237] View Article

[238] PubMed/NCBI

[239] Google Scholar

[ref67] 67. Balestrino F, Medici A, Candini G, Carrieri M, Maccagnani B, Calvitti M, et al. Gamma ray dosimetry and mating capacity studies in the laboratory on Aedes albopictus males. J Med Entomol. 2010; 47: 581–591. pmid:20695273
View Article
PubMed/NCBI
Google Scholar

[241] View Article

[242] PubMed/NCBI

[243] Google Scholar

Figures

Abstract

Introduction

Materials and Methods

Ethics Statement

Mosquito lines and rearing

Age- and sex-specific Wolbachia density in SCRE and SANG mosquito lines

Crosses for CI strength assessment

Wolbachia density in wild caught males

Wolbachia genotyping and qPCR

DNA purification and qPCR.

Construction of plasmids for obtaining qPCR standard curves.

Statistical analysis and CI computation.

Results

Age- and sex-specific wAlbA and wAlbB densities

Effect of male wAlbA and wAlbB densities on CI expression

Wolbachia density in wild caught males

Discussion

Supporting Information

S1 Fig. Evaluation of PCR sensitivity in relation to increasing wAlbA titers, calculated as wAlbA/actin copy numbers by qPCR on independent mosquito extracts: (1) 0.0001; (2) 0.0010; (3) 0.0039; (4) 0.0108; (5) 0.0582; (6) 0.1671; (7) 2.2938; (8) no template PCR control.

Acknowledgments

Author Contributions

References

Age- and sex-specific Wolbachia density in S_CRE and S_ANG mosquito lines