Paratransgenesis to control malaria vectors: a semi-field pilot study
- Maria Vittoria Mancini†1,
- Roberta Spaccapelo†2,
- Claudia Damiani1,
- Anastasia Accoti1,
- Mario Tallarita2,
- Elisabetta Petraglia1,
- Paolo Rossi1,
- Alessia Cappelli1,
- Aida Capone1,
- Giulia Peruzzi2,
- Matteo Valzano1,
- Matteo Picciolini2,
- Abdoulaye Diabaté3,
- Luca Facchinelli2,
- Irene Ricci1 and
- Guido Favia1Email author
© Mancini et al. 2016
Received: 21 December 2015
Accepted: 5 March 2016
Published: 10 March 2016
Malaria still remains a serious health burden in developing countries, causing more than 1 million deaths annually. Given the lack of an effective vaccine against its major etiological agent, Plasmodium falciparum, and the growing resistance of this parasite to the currently available drugs repertoire and of Anopheles mosquitoes to insecticides, the development of innovative control measures is an imperative to reduce malaria transmission. Paratransgenesis, the modification of symbiotic organisms to deliver anti-pathogen effector molecules, represents a novel strategy against Plasmodium development in mosquito vectors, showing the potential to reduce parasite development. However, the field application of laboratory-based evidence of paratransgenesis imposes the use of more realistic confined semi-field environments.
Large cages were used to evaluate the ability of bacteria of the genus Asaia expressing green fluorescent protein (Asaia gfp), to diffuse in Anopheles stephensi and Anopheles gambiae target mosquito populations. Asaia gfp was introduced in large cages through the release of paratransgenic males or by sugar feeding stations. Recombinant bacteria transmission was directly detected by fluorescent microscopy, and further assessed by molecular analysis.
Here we show the first known trial in semi-field condition on paratransgenic anophelines. Modified bacteria were able to spread at high rate in different populations of An. stephensi and An. gambiae, dominant malaria vectors, exploring horizontal ways and successfully colonising mosquito midguts. Moreover, in An. gambiae, vertical and trans-stadial diffusion mechanisms were demonstrated.
Our results demonstrate the considerable ability of modified Asaia to colonise different populations of malaria vectors, including pecies where its association is not primary, in large environments. The data support the potential to employ transgenic Asaia as a tool for malaria control, disclosing promising perspective for its field application with suitable effector molecules.
KeywordsAsaia Anopheles Paratransgenesis Large cages trials
The emergence of drug resistant parasites and insecticide resistant mosquito strains, together with several eco-environmental concerns related to the use of most chemicals, require the development of additional control methods for mosquito-borne diseases . In addition to transgenic mosquitoes engineered to replace or suppress wild vector populations [2–4], a parallel approach aimed at producing paratransgenic tools to control vector-borne diseases has been developed, providing concrete possibilities for innovative control strategies [5–7].
Paratransgenesis is commonly defined as the use of symbiotic organisms, naturally inhabiting mosquito midgut and rapidly spreading among vector population, to deliver anti-pathogen effector molecules [8–10]. In the last decade several studies focusing on effective paratransgenic-based malaria control protocols have been published and a few bacterial symbionts have been already selected as potentially useful tools, although all related studies have been performed in small laboratory cages [11–15]. The transition from small laboratory cages to open field trials is a critical step to effectively set-up an in-depth control approach . In this context, the intermediate step of confined semi-field conditions represents an ideal tool to evaluate the potential of paratransgenesis technology to be employed to counteract malaria and other mosquito-borne diseases. At the same time, it gives the possibility to develop predictive models and comprehensive risk assessment related to the use of paratransgenic mosquitoes. The use of large cages allows a wider picture of the actual transmission potential of selected symbiont(s), together with preliminary behavioural ecology insights of paratransgenic mosquitoes, in a specifically arranged environment, simulating the near-natural ecosystem conditions . To our knowledge, no complete surveys in large cages have been yet performed for paratransgenesis.
The acetic acid bacterium Asaia is one of the most promising mosquito symbionts for paratransgenic approach. Asaia investigations in Anopheles stephensi, where it represents the dominant commensal genus, disclosed its ability to spread with high efficiency in recipient populations and throughout following generations, as demonstrated in small laboratory cages . Moreover, the association between Asaia and field collected An. gambiae was reported . Its intrinsic biological characteristics, easy transformability and capability to be transmitted through horizontal and vertical transmission routes in small cages, together with its colonisation throughout the mosquito life-cycle, as well as its co-localisation in Plasmodium invasion hot-spots, are invaluable features that make this bacterium a suitable candidate for symbiotic control strategies [20, 21].
Laboratory-strain colonies of Anopheles gambiae (G3) and Anopheles stephensi (SD500) were used. Mosquitoes were reared at 27 °C at a relative humidity of 70 %. Larvae were reared in deionised water to which 0.3 g/liter of artificial sea salts were added, and were fed daily with a diet provided as a slurry of 2:2:1 bovine liver powder, tuna meal and Vanderzant vitamin mix . Adults were maintained with wet cottons pads soaked with a 5 % sucrose solution. For 16S rRNA gene pyrosequencing analysis, to evaluate the effect of kanamycin on mosquito microbiota, An. gambiae were reared in bugdorms and maintained for 10 days with 5 % sucrose solution plus kanamycin (100 μg/ml).
Asaia sp. growth
Asaia sp. SF2.1 (GFP)  (hereafter, Asaia gfp) was grown 24 h at 30 °C in GLY medium (25 ml/L glycerol, 10 g/L yeast extract; pH 5.0). Cells were grown to OD600 = 1.0 (108 recombinant bacterial cells/ml), precipitated, washed three times in 0.9 % NaCl and resuspended in 5 % sucrose solution. For monitoring long-term colonisation, the suspension was supplemented with 100 μg/ml of kanamycin to avoid plasmid loss from bacterial cells.
Semi-field set up
The study was carried out at the confined release facility of the Department of Experimental Medicine, University of Perugia. Three large experimental cages (A-B-C) of 15.9 m3 each were located in a 6.68 × 3.80 × 3.00 m chamber, with complete control of environmental conditions. Briefly, a 24 h light cycle provided by four ceiling lights, dawn lasted for 30 min, full light lasted for 11.5 h and twilight lasted for 1 h and 30 min of fading ceiling light from full light to minimum power simulating sunset. Each large cage was equipped with clay resting shelters kept humid and swarming stimuli consisted of a square arena made of contrasting black and white ground marks [22, 24].
Asaia sp. horizontal and vertical transmission
Horizontal spreading of Asaia gfp through mosquito populations was achieved by either the release in the large cages of colonised males or by infected cotton pads. Paratransgenic An. stephensi and An. gambiae mosquitoes were obtained by oral infection of newly emerged males in bugdorms with cotton pads soaked with 5 % sucrose solution enriched with 108 recombinant Asaia gfp cells/ml and kanamycin (100 μg/ml) for 5 days. Colonised males were marked with pink fluorescent powder prior to their release in the large cages, as previously described . Non-colonised males from the same batch were maintained in the same conditions. The three semi-field cages were populated with An. stephensi as follows: 200 newly emerged females and 200 5 day-old non-colonised males prior the release of 12 and 36 Asaia gfp-colonised males, in cage A and B, respectively. Experiments with An. gambiae mosquitoes were performed using the same experimental design, but only with the lowest amount of paratransgenic males (12 males in cage A). Cage C represents the negative control without insertion of paratransgenic males.
Mosquitoes were maintained with cotton pads soaked with a solution composed of 5 % sucrose, 10 % peach juice, kanamycin (100 μg/ml) and methylparaben (0.1 %) as preservative. To assess Asaia gfp horizontal transmission, 50 females and 50 males from each cage sampled at 5, 12 and 20 days after the start of the experiment were dissected under a stereomicroscope and their guts investigated by fluorescent microscopy (Nikon Eclipse TE 2000-U); some samples were additionally analysed by PCR as described below.
To evaluate Asaia gfp horizontal transmission in An. gambiae population by infected cotton pads, a cohort of newly emerged 350 males or 350 females was introduced in cage A and B, respectively. Cage C hosted 350 females and 350 males from several cohorts of individuals of different ages: in order to establish it, newly emerged adults were constantly reintroduced to maintain the density and the age distribution as stable as possible, for a month and a half before the start of the experiment. Each semi-field cage contained two uninfected cotton pads soaked with a solution composed of 5 % sucrose, 10 % peach juice, kanamycin (100 μg/ml) and methylparaben (0.1 %), and one supplemented with 108 Asaia gfp cells/ml. Bacterial spread was assessed after 5 and 12 days after the release. Paternal and maternal contributions to Asaia gfp transmission were analysed by releasing 5 day-old naïve females or males into cages A and B, respectively. Prior to the release, all the infected feeding stations were removed from cages in order to restrict co-feeding transmission routes. Mosquitoes were allowed to mate for 24 h.
To assess Asaia gfp vertical transmission route at the end of the above described trials, females were collected and blood-fed through a Hemotek PS5 membrane feeder (Discovery Workshops, UK) at 37 °C. Engorged female mosquitoes were provided with wet filter paper for oviposition: laid eggs were floated in larval pans with breeding water containing 100 μg/ml of kanamycin, and maintained as above. Adults and 4th instar larvae, collected immediately after eclosion, were sampled and screened by fluorescent microscopy and by PCR. Data were validated by statistical analysis using G test  and Bonferroni post-hoc test and performed in R (http://www.r-project.org/).
DNA extraction, PCR analysis and metagenomic library preparation
Prior to DNA extraction, mosquitoes were surface-sterilised by immersion in 70 % ethanol and washed in PBS for three times, then processed with an automatic tissue homogeniser (Precellys 24, Bertin Technologies SAS, Villeurbanne, France). Genomic DNA was extracted using a JetFlex Genomic DNA Purification kit (Genomed, Lohne, Germany) according to the manufacturer’s instructions. Samples of 5 individuals were pooled together for 16S DNA pyrosequencing. Molecular analysis of Asaia gfp-infected mosquitoes was performed by PCR using the following Asaia gfp specific oligonucleotide primers (200 nM): FOR: 5'-CAA GAG TGC CAT GCC CGA AGG-3' and REV: 5'-GAC AGG GCC ATC GCC AAT TGG-3'. PCR was performed using the DreamTaq DNA polymerase kit (Thermo Fisher Scientific, Waltham, Massachusetts, USA) according to the manufacturer’s protocol; 50 ng of genomic DNA was amplified with an initial denaturation at 94 °C for 3 min, followed by 30 cycles consisting of denaturation at 94 °C for 30 s, annealing at 60 °C for 30 s, extension at 72 °C 30 for s, and ultimately a final step at 72 °C for 10 min.
Paired-end 16S community sequencing on the Illumina MiSeq platform was performed by Polo d’Innovazione di Genomica, Genetica e Biologia, Perugia, using bacteria/archaeal degenerate primers 515 F/806R  to target the 16S V4 regions. 50 ng of gDNA was used for PCR amplification in 25 μl reaction volume, containing 2 μM AmpliTaq Gold 360 Master Mix (Applied Biosystem, Foster City, California, USA) and 5 μM each of the oligonucleotide primers. All amplifications were performed in a T100 Thermal Cycler (BIO-RAD, CA, USA) with an initial step at 94 °C for 3', followed by 35 cycles: 94 °C for 45 s, 55 °C for 1 min, and 72 °C for 1.5 min, and a final step at 72 °C for 10 min. Metagenomic libraries were prepared using the Nextera XT protocol (Illumina, San Diego, California, USA). Briefly, 1 ng of the purified amplicons were tagmented by the transposon, amplified via a limited-cycle PCR program that also adds index 1 (i7) and index 2 (i5), size-exclusion purified by Agencourt beads (Beckman Coulter, Brea, California, USA), normalised and pooled. The samples libraries were sequenced in a 2 × 250 PE using MiSeq Reagent Kit v2.
Metagenomic data analysis
Metagenomic raw data were cleaned first by removing Phix contaminants using bowtie2 , then the sequence of the adapters and low quality scores (<20) were trimmed by Trimmomatic . Cleaned reads were used to assemble the amplicons using Pear . Amplicons were dereplicated, sorted and clustered to identify the OTU by vsearch (https://github.com/torognes/vsearch). OTU taxonomy was determined by a basic local alignment with BLASTn of amplicons, against the SILVA database v.119 . The abundance of all OTUs identified was calculated by the alignment of the raw amplicons to the OTU references using vsearch. Normalisation and evaluation of relative abundance were performed by R (http://www.r-project.org/).
Results and discussion
The trial involving horizontal bacterial transmission through sugar feeding station provided outcomes of a notable diffusion rate of Asaia in An. gambiae population and in the environment. Two uninfected and one Asaia gfp-infected feeding stations were introduced in each large cage. After two days all the feeding stations were screened for Asaia gfp infection, resulting all positive (data not shown). Cages A, B and C were populated with only females, males or assorted stable-age mosquito population, respectively. The rate of Asaia gfp-infected mosquitoes increased over time and reached 91 %, 95 % and on average 79 % respectively, at 12 days post-release, thus indicating a successful horizontal transmission within the recipient population (G = 15.1, 1 d.f., P = 9.71 e-05) (Fig. 3b).
Overall, our data are consistent with the preliminary observations obtained in previous studies performed in small cages with An. stephensi . Additionally, this study contributes to a better understanding transmission routes and employed vectors. The strong ability of modified Asaia to be horizontally spread in different populations, through the release of previously infected males or through feeding stations in populations of both An. stephensi and An. gambiae is demonstrated. At the same time, our data suggest intrinsic behavioural and ecological differences between the two vector systems. The slightly lower and delayed rate of infection of An. stephensi compared to An. gambiae has been addressed to the internal arrangement of the large cages, optimised for swarming and mating behaviour of An. gambiae. Thus, since horizontal spreading of Asaia gfp mainly relies on co-feeding and mating, the behaviour of the mosquitoes in this semi-field condition may have limited the second route of infection. For these reasons, the analysis of the first generation of An. stephensi has been prevented.
The possibility to release non-biting paratransgenic males in open field will circumvent the concerns of releasing bacteria-transmitting females, being consistent with safety requirements related to the use of paratransgenesis to reduce vector competence. Nevertheless, complexities could still arise given the fact that this practice may result in the overall increase of the mosquito population density in a given area. Therefore, we also propose an alternative approach for Asaia gfp transmission assessment, whose introduction does not imply the release of mosquitoes, and recombinant bacteria were introduced in large cages by means of feeding stations. Both pathways definitely demonstrate the high efficiency of Asaia to diffuse and colonise mosquito populations in large environment with respect to small cages. This effectiveness explores both horizontal, mainly by synergistic co-feeding and mating, and vertical diffusion pathways of both paternal and maternal contribution, despite the introduction procedures applied. Our data, obtained by exploiting fluorescence marked bacteria, lay the foundation for further applications of Asaia as paratransgenic tool.
The ability of paratransgenic approaches to control malaria and other mosquito-borne diseases is very promising, disclosing a concrete applicative prospect. We report here results from the first known paratransgenic trial performed in large cages aimed at testing the feasibility of this approach. The success of paratransgenesis obviously depends on a variety of factors. Nevertheless, our findings support the applied perspective involving the use of Asaia as a promising tool and further demonstrate the great utility of confined environments to define the most efficient methodologies for an in-depth evaluation of technologies transition from laboratory to field employment. The field release of paratransgenic mosquitoes imposes a rigorous risk assessment framework coherent with a strict regulatory system, appropriate to national and international guidelines. Evaluation of the risks and benefits of this strategy is required. Investigation of hazards and safety related concerns , together with implementation of authorised ongoing projects (transgenic and/or Wolbachia-transinfected mosquitoes ) will lay the basis for a solid regulatory oversight of the paratransgenic program, and ultimately, to allow its field trials. Since Asaia has been recently detected in several insect vectors [34–37], these data provide crucial clues applicable toward multiple paratransgenic targets in the control of a wide spectrum of vector borne diseases.
This paper is dedicated to our friend and colleague Thanasis Loukeris, who died on 30 May 2014. We would like to thank Sheila Beatty for editing the English of the manuscript. The work was supported by grants (to GF) from the Italian Ministry of Education, University and Research (MIUR) (Prin 2012 protocol 2012T85B3R), the EU-FP7 Capacities-Infrastructure 2008 (grant 228421) and (to IR) from the European Union Seventh Framework Programme ([FP7/2007-2013] under grant agreement n. 281222. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
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- Scott TW, Takken W, Knols BG, Boëte C. The ecology of genetically modified mosquitoes. Science. 2002;298:117–9.View ArticlePubMedGoogle Scholar
- Knols BG, Bossin HC, Mukabana WR, Robinson AS. Transgenic mosquitoes and the fight against malaria: managing technology push in a turbulent GMO world. Am J Trop Med Hyg. 2007;77(6 Suppl):232–42.PubMedGoogle Scholar
- Coutinho-Abreu IV, Zhu KY, Ramalho-Ortigao M. Transgenesis and paratransgenesis to control insect-borne diseases: current status and future challenges. Parasitol Int. 2010;59(1):1–8.View ArticlePubMedPubMed CentralGoogle Scholar
- Bourtzis K, Lees RS, Hendrichs J, Vreysen MJ. More than one rabbit out of the hat: radiation, transgenic and symbiont-based approaches for sustainable management of mosquito and tsetse fly populations. Acta Trop. 2016;157:115–30.View ArticlePubMedGoogle Scholar
- Hurwitz I, Fieck A, Read A, Hillesland H, Klein N, Kang A, et al. Paratransgenic control of vector borne diseases. Int J Biol Sci. 2011;7(9):1334–44.View ArticlePubMedPubMed CentralGoogle Scholar
- Ren X, Hoiczyk E, Rasgon JL. Viral paratransgenesis in the malaria vector Anopheles gambiae. PLoS Pathog. 2008;4(8):e1000135.View ArticlePubMedPubMed CentralGoogle Scholar
- Hurwitz I, Hillesland H, Fieck A, Das P, Durvasula R. The paratransgenic sand fly: a platform for control of Leishmania transmission. Parasit Vectors. 2011;4:82.View ArticlePubMedPubMed CentralGoogle Scholar
- Wilke AB, Marrelli MT. Paratransgenesis: a promising new strategy for mosquito vector control. Parasit Vectors. 2015;8:342.View ArticlePubMedPubMed CentralGoogle Scholar
- Ricci I, Valzano M, Ulissi U, Epis S, Cappelli A, Favia G. Symbiotic control of mosquito borne disease. Pathog Glob Health. 2012;106(7):380–5.View ArticlePubMedPubMed CentralGoogle Scholar
- Villegas LM, Pimenta PF. Metagenomics, paratransgenesis and the Anopheles microbiome: a portrait of the geographical distribution of the anopheline microbiota based on a meta-analysis of reported taxa. Mem Inst Oswaldo Cruz. 2014;109(5):672–84.View ArticlePubMedPubMed CentralGoogle Scholar
- Dinparast Djadid N, Jazayeri H, Raz A, Favia G, Ricci I, Zakeri S. Identification of the midgut microbiota of An. stephensi and An. maculipennis for their application as a paratransgenic tool against malaria. PLoS One. 2011;6(12):e28484.View ArticlePubMedPubMed CentralGoogle Scholar
- Riehle MA, Moreira CK, Lampe D, Lauzon C, Jacobs-Lorena M. Using bacteria to express and display anti-Plasmodium molecules in the mosquito midgut. Int J Parasitol. 2007;37(6):595–603.View ArticlePubMedGoogle Scholar
- Favia G, Ricci I, Damiani C, Raddadi N, Crotti E, Marzorati M, et al. Bacteria of the genus Asaia stably associate with Anopheles stephensi, an Asian malarial mosquito vector. Proc Natl Acad Sci U S A. 2007;104(21):9047–51.View ArticlePubMedPubMed CentralGoogle Scholar
- Wang S, Jacobs-Lorena M. Genetic approaches to interfere with malaria transmission by vector mosquitoes. Trends Biotechnol. 2013;31(3):185–93.View ArticlePubMedPubMed CentralGoogle Scholar
- Chavshin AR, Oshaghi MA, Vatandoost H, Pourmand MR, Raeisi A, Terenius O. Isolation and identification of culturable bacteria from wild Anopheles culicifacies, a first step in a paratransgenesis approach. Parasit Vectors. 2014;7:419.View ArticlePubMedPubMed CentralGoogle Scholar
- WHO/TDR and FNIH The Guidance Framework for testing genetically modified mosquitoes. 2014. ISBN 978 92 4 150748 6.Google Scholar
- Knols BG, Njiru BN, Mathenge EM, Mukabana WR, Beier JC, Killeen GF. MalariaSphere: a greenhouse-enclosed simulation of a natural Anopheles gambiae (Diptera: Culicidae) ecosystem in western Kenya. Malar J. 2002;1:19.View ArticlePubMedPubMed CentralGoogle Scholar
- Damiani C, Ricci I, Crotti E, Rossi P, Rizzi A, Scuppa P, et al. Paternal transmission of symbiotic bacteria in malaria vectors. Curr Biol. 2008;18(23):R1087–1088.View ArticlePubMedGoogle Scholar
- Damiani C, Ricci I, Crotti E, Rossi P, Rizzi A, Scuppa P, et al. Mosquito-bacteria symbiosis: the case of Anopheles gambiae and Asaia. Microb Ecol. 2010;60(3):644–54.View ArticlePubMedGoogle Scholar
- Ricci I, Damiani C, Capone A, DeFreece C, Rossi P, Favia G. Mosquito/microbiota interactions: from complex relationships to biotechnological perspectives. Curr Opin Microbiol. 2012;15(3):278–84.View ArticlePubMedGoogle Scholar
- Capone A, Ricci I, Damiani C, Mosca M, Rossi P, Scuppa P, et al. Interactions between Asaia, Plasmodium and Anopheles: new insights into mosquito symbiosis and implications in malaria symbiotic control. Parasit Vectors. 2013;6(1):182.View ArticlePubMedPubMed CentralGoogle Scholar
- Facchinelli L, Valerio L, Lees RS, Oliva CF, Persampieri T, Collins CM, et al. Stimulating Anopheles gambiae swarms in the laboratory: application for behavioural and fitness studies. Malar J. 2015;14:271.View ArticlePubMedPubMed CentralGoogle Scholar
- Damiens D, Benedict MQ, Wille M, Gilles JR. An inexpensive and effective larval diet for Anopheles arabiensis (Diptera: Culicidae): eat like a horse, a bird, or a fish? J Med Entomol. 2012;49(5):1001–11.View ArticlePubMedGoogle Scholar
- Valerio L, Facchinelli L, Ramsey JM, Bond JG, Scott TW. Dispersal of male Aedes aegypti in a coastal village in southern Mexico. Am J Trop Med Hyg. 2012;86(4):665–76.View ArticlePubMedPubMed CentralGoogle Scholar
- Sokal RR, Rohlf FJ. Biometry. San Francisco: W.H. Freeman and Company; 1981.Google Scholar
- Caporaso JG, Lauber CL, Walters WA, Berg-Lyons D, Huntley J, Fierer N, et al. Ultra-high-throughput microbial community analysis on the Illumina HiSeq and MiSeq platforms. ISME J. 2012;6:1621–4.View ArticlePubMedPubMed CentralGoogle Scholar
- Langmead B, Salzberg SL. Fast gapped-read alignment with Bowtie 2. Nat Methods. 2012;9(4):357–9.View ArticlePubMedPubMed CentralGoogle Scholar
- Bolger AM, Lohse M, Usadel B. Trimmomatic: a flexible trimmer for Illumina sequence data. Bioinformatics. 2014;30(15):2114–20.View ArticlePubMedPubMed CentralGoogle Scholar
- Zhang J, Kobert K, Flouri T, Stamatakis A. PEAR: a fast and accurate Illumina Paired-End reAd mergeR. Bioinformatics. 2014;30(5):614–20.View ArticlePubMedPubMed CentralGoogle Scholar
- Quast C, Pruesse E, Yilmaz P, Gerken J, Schweer T, Yarza P, et al. The SILVA ribosomal RNA gene database project: improved data processing and web-based tools. Nucleic Acids Res. 2013;41(Database issue):D590–596.View ArticlePubMedPubMed CentralGoogle Scholar
- Kukutla P, Lindberg BG, Pei D, Rayl M, Yu W, Steritz M, et al. Insights from the genome annotation of Elizabethkingia anophelis from the malaria vector Anopheles gambiae. PLoS One. 2014;9(5):e97715.View ArticlePubMedPubMed CentralGoogle Scholar
- De Freece C, Paré Toé L, Esposito F, Diabaté A, Favia G. Preliminary assessment of framework conditions for release of genetically modified mosquitoes in Burkina Faso. Int Health. 2014;6:263–5.View ArticlePubMedGoogle Scholar
- Nguyen TH, Nguyen HL, Nguyen TY, Vu SN, Tran ND, Le TN, et al. Field evaluation of the establishment potential of wMelPop Wolbachia in Australia and Vietnam for dengue control. Parasit Vectors. 2015;8:563.View ArticlePubMedPubMed CentralGoogle Scholar
- De Freece C, Damiani C, Valzano M, D'Amelio S, Cappelli A, Ricci I, et al. Detection and isolation of the α-proteobacterium Asaia in Culex mosquitoes. Med Vet Entomol. 2014;28(4):438–42.View ArticlePubMedGoogle Scholar
- Mee PT, Weeks AR, Walker PJ, Hoffmann AA, Duchemin JB. Detection of low level Cardinium and Wolbachia infections in Culicoides. Appl Environ Microbiol. 2015;81(18):6177–88.View ArticlePubMedPubMed CentralGoogle Scholar
- Sant'Anna MR, Diaz-Albiter H, Aguiar-Martins K, Al Salem WS, Cavalcante RR, Dillon VM, et al. Colonisation resistance in the sand fly gut: Leishmania protects Lutzomyia longipalpis from bacterial infection. Parasit Vectors. 2014;7:329.View ArticlePubMedPubMed CentralGoogle Scholar
- Crotti E, Damiani C, Pajoro M, Gonella E, Rizzi A, Ricci I, et al. Asaia, a versatile acetic acid bacterial symbiont, capable of cross-colonizing insects of phylogenetically distant genera and orders. Environ Microbiol. 2009;11(12):3252–64.View ArticlePubMedGoogle Scholar