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. 2019 Nov 5;4:11. Originally published 2019 Jan 24. [Version 4] doi: 10.12688/wellcomeopenres.15005.4

Detection of diverse Wolbachia 16S rRNA sequences at low titers from malaria vectors in Kayin state, Myanmar

Sunisa Sawasdichai 1, Victor Chaumeau 1,2,a, Tee Dah 1, Thithiworada Kulabkeeree 1, Ladda Kajeechiwa 1, Monthicha Phanaphadungtham 1, Muesuwa Trakoolchengkaew 1, Praphan Kittiphanakun 1, Yanada Akararungrot 1, Kyi Oo 1, Gilles Delmas 1,2, Nicholas J White 2,3, François H Nosten 1,2
PMCID: PMC6892426  PMID: 31828225

Version Changes

Revised. Amendments from Version 3

In this revised version of the manuscript, the wording "Wolbachia infection" was changed in the title and throughout the main text in order to address reviewer's concern that the detection of Wolbachia DNA sequences with PCRs is not a demonstration of an actual infection of the insect host with the bacteria. Primer sequences and amplicon length were added in the Methods section. The possibility that Wolbachia DNA detected in this study may come from some sort of environmental contamination given the high number of arthropod species infected with this bacteria has been made more explicit by adding a specific statement and the suggested reference in the Discussion section.

Abstract

Background : Natural Wolbachia infections in malaria mosquitoes were recently reported in Africa, and negatively correlated with the development of Plasmodium falciparum in the vectors. The occurrence and effects of Wolbachia infections outside Africa have not been described and may have been underestimated.

Methods : Mosquitoes were collected by human-landing catch during May and June 2017 in ten villages in Kayin state, Myanmar. Closely related species of malaria vectors were identified with molecular assays. 16S rRNA Wolbachia DNA sequences were detected with quantitative real-time PCR.

Results: Low titer of Wolbachia DNA was detected in 13/370 samples in six malaria vector species. Sequences were diverse and different from those described in the African malaria mosquitoes.

Conclusion: The detection of Wolbachia DNA in malaria mosquitoes from Kayin state warrants further investigations to understand better the ecology and biology of Anopheles- Wolbachia interactions in Southeast Asia.

Keywords: Wolbachia, Anopheles, Plasmodium, 16S rRNA, entomological inoculation rate, Southeast Asia, Kayin state, wAnga

Introduction

Wolbachia are intracellular bacteria that infect a wide variety of arthropods and filarial nematodes. Symbiotic relationship that results from the infection have a broad range of phenotypic effects on the infected hosts, from mutualism (beneficial) to commensalism (neutral) and parasitism (harmful) 1. In mosquitoes, Wolbachia can invade the germline and induce cytoplasmic incompatibilities between the sperm from infected males and oocytes from uninfected females 2. Hence, mass-releases of Wolbachia-infected male mosquitoes were attempted to extinguish mosquito populations 3, 4. Cytoplasmic incompatibilities produce a fitness advantage of Wolbachia-infected over uninfected female mosquitoes, thereby driving the spread of Wolbachia-infected females in the population. In addition, Wolbachia can interfere with the development of some pathogens in the mosquito host, including dengue virus 5, Plasmodium malaria parasites 6 and filarial nematodes 7. Therefore, the release of Wolbachia-infected female mosquitoes is proposed for transmission-blocking of some mosquito-borne diseases 8.

Most diversions of mosquito- Wolbachia interactions for controlling vector-borne diseases were conducted with mosquitoes artificially infected with the endosymbiont. Natural Wolbachia infections may have important effects on mosquito populations and dynamics of diseases transmission but they are less well described 9. Wolbachia DNA was detected by PCR in 27 mosquito genera including the medically important Aedes, Armigeres, Culex, Mansonia and Stegomiya 9– 18. Interestingly, this organism was not detected in malaria mosquitoes until recent observations of naturally infected anopheline vectors in Africa 9, 13, 19– 24.

Only one study assessed the effects of natural Wolbachia infection on the reproductive fitness anopheline mosquitoes, namely the dominant African malaria vector Anopheles coluzzi 22. The authors did not observe cytoplasmic incompatibilities, differences in the number of eggs laid or progeny sex ratio, but infected females laid eggs more rapidly. Two studies demonstrated the negative effects of Wolbachia infections on the development of P. falciparum 20, 22. Shaw et al. observed a negative correlation between Wolbachia infection and the development of P. falciparum in naturally blood-fed females. Gomes et al. obtained similar results on the sporozoite stage by screening large numbers of mosquitoes identified as An. gambiae sensu stricto and An. coluzzi. In addition to their field investigations, Gomes et al. infected a laboratory-adapted An. coluzzi colony with a local strain of Wolbachia, and performed artificial transmission studies with cultured gametocytes of P. falciparum strain NF54. They observed a moderate yet significant positive correlation between Wolbachia infection and oocyst development, and a negative correlation between Wolbachia infection and the number of sporozoites that subsequently invaded the salivary glands.

Natural Wolbachia infections in Southeast Asian malaria vectors have not been reported. Their potential effects on Anopheles mosquitoes and dynamics of malaria transmission are not known. The objective of this study was therefore to assess the presence of Wolbachia in malaria vector populations in Kayin state, Myanmar.

Methods

Study sites and entomological collections

Entomological surveys were conducted in May and June 2017 in ten villages in Kayin state, Myanmar ( Figure 1). Each survey consisted of five consecutive nights of collection from 06:00 pm to 06:00 am as described previously 25. In each village, five traditional houses were selected for mosquito sampling with human-landing catches. Collectors were asked to collect every mosquitoes landing on their uncovered legs for 50 min per hour and allowed to rest for 10 min per hour. Mosquitoes were shipped to Mae Sot (Thailand) at the end of each survey.

Figure 1. Map of the study area.

Figure 1.

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Malaria vectors identification

Mosquitoes were immediately identified at the genus level by morphology and Anopheles specimen were stored individually at -20°C in 1.5 mL plastic tubes containing silica gel. Anopheles were identified at the Group or Complex level using the key developed by Rattanarithikul et al. 26. Closely related species in the Funestus, Maculatus and Leucopshyrus Groups were discriminated in a subsample of the total number of collected mosquito using allele-specific PCR assays (AS-PCR) adapted from Garros et al. and Walton et al. 27– 29. Single whole mosquitoes were crushed in 200 μl of cetyl-trimethylammonium bromide solution 2% (TrisHCl pH = 8, 20mM; EDTA 10mM; NaCl, 1.4 mM; N-cetyl-N,N,N-trimethyl ammonium bromide 2%) with a TissueLyser II™ (Qiagen) set on 29 movements /second for 3 minutes. Samples were then warmed at 65°C for 5 minutes and 200 μl of chloroform were added. The aqueous phase was collected and DNA was precipitated with 200 μl of isopropanol. After centrifugation at 20,000 g for 15 minutes, the pellet was washed twice with 200 µl of 70% ethanol and suspended in 50 μl of PCR grade water 30. The PCR mix was composed of 1X Goldstar™ DNA polymerase (Eurogentec, Seraing, Belgium) and 400 nM of each primer (Funestus assay: ITS2A 5’-TGT GAA CTG CAG GAC ACA T-3’, MIA 5’-CCC GTG CGA CTT GAC GA-3’, MIC 5’-GTT CAT TCA GCA ACA TCA GT-3’, ACO 5’-ACA GCG TGT ACG TCC AGT-3’, PAM 5’-TGT ACA TCG GCC GGG GTA-3’, VAR 5’-TTG ACC ACT TTC GAC GCA-3’; Maculatus assay: 5.8F 5’-TGT GAA CTG CAG GAC ACA T-3’, MAC 5’-CCC GTG CGA CTT GAC GA-3’, PSEU 5’-GTT CAT TCA GCA ACA TCA GT-3’, SAW 5’-ACA GCG TGT ACG TCC AGT-3’, K 5’-TGT ACA TCG GCC GGG GTA-3’, DRAV 5’-TTG ACC ACT TTC GAC GCA-3’ and Leucopshyrus assay: D-AC 5’-CAC AGC GAC TCC ACA CG-3’, D-B 5’-CGG GAT ATG GGT CGG CC-3’, D-D 5’-GCG CGG GAC CGT CCG TT-3’, D-F 5’-AAC GGC GGT CCC CTT TG-3’, D-AC 5’-CAC AGC GAC TCC ACA CG-3’). The PCR was conducted in a total reaction volume of 25 μl (1 μl of DNA template and 24 μl of PCR mix). The thermocycling protocol consisted in an initial activation step of 1 minute at 94°C, followed by 40 amplification cycles of 20 seconds at 94°C, 20 seconds at the appropriate annealing temperature (45°C for the Funestus assay, and 55°C for the Maculatus and Leucosphyrus assays), and 30 seconds at 72°C. The length of the PCR product was determined by gel electrophoresis in 2% agarose for 70 minutes at 120V. In case AS-PCR gave a negative result, amplification of ITS2 was performed using the primer pair ITS2A (5'-TGT GAA CTG CAG GAC ACA T-3') and ITS2B (5'-ATG CTT AAA TTY AGG GGG T-3') described by Beebe and Saul 31. The PCR mix was composed of 1X Goldstar™ DNA polymerase (Eurogentec, Seraing, Belgium) and 400 nM of each primer. The PCR was conducted in a total reaction volume of 25 μl (1 μl of DNA template and 24 μl of PCR mix). The thermocycling protocol consisted in an initial activation step of 1 minute at 94°C, followed by 40 amplification cycles of 20 seconds at 94°C, 20 seconds at 51°C and 30 seconds at 72°C. PCR products were purified on site using the Illustra™ ExoStar™ PCR and Sequence Reaction Clean-Up Kit (GE Healthcare) according to manufacturer’s instruction. Macrogen (Seoul, South Korea) sequenced the purified PCR products off site with the ITS2A primer. Sequences were blasted against the National Center for Biotechnology Information nucleotide database in order to determine the corresponding species (accession numbers MK358471 - MK358807).

Detection of Wolbachia DNA by quantitative real-time PCR

Two primer sets were considered for Wolbachia screening in mosquito samples: W-Specf/W-Specr (5’-CAT ACC TAT TCG AAG GGA TAG-3’ and 5’- AGC TTC GAG TGA AAC CAA TTC-3’) amplified a 438 bp conserved region of the 16S rRNA genes and W-Specf/W16S (5’- CAT ACC TAT TCG AAG GGA TAG -3’ and 5’- TTG CGG GAC TTA ACC CAA CA -3’) amplified a shorter fragment of the same region (102 bp). These two sets were selected because they were previously used by other in order to detect Wolbachia in Anopheles mosquitoes 20. Without a priori knowledge on Wolbachia DNA sequences detected in this study, the W-Specf/W-Specr primer set was selected for its ability to detect most Wolbachia strains infecting insects and to establish phylogenetic relationships among isolates 32.

The performances of the primers W-Specf/W-Specr for the detection and quantitation of Wolbachia in mosquito samples were compared to that of the primers W-Specf/W16S as described previously 30. Briefly, a published strain of laboratory-reared Aedes aegypti artificially infected with Wolbachia strain wMel were used as a reference material 33. The optimal conditions for the PCR (hybridization temperature for primers annealing, and concentration of MgCl 2 and primers) were determined during a single gradient experiment in order to take into account cross-interactions between the different parameters. The range tested were 55–62°C for the hybridization temperature, 2.5–4.5 mM of MgCl 2 and 100–400 nM of each primers. The reaction conditions that gave the smallest CP (optimal conditions) were selected for all subsequent experiments. Serial-dilution experiments were then carried out in order to verify PCR efficiency (EFF) and to estimate the standard curve parameters.

All experiments were conducted with a CFX-96® (Biorad) device. Reactions were conducted in 20µl of EVAGreen qPCR Mix Plus® (Euromedex); 5µl of DNA template was used in a total reaction volume of 25µl. The PCR mix was composed of 1X HOT FIREPol™ EvaGreen™ qPCR Mix Plus (Solis BioDyne, Tartu, Estonia) and 200 nM of each primer. The thermocycling protocol consisted in an activation step at 95°C for 15 minutes followed by 45 amplification cycles at 95°C for 15 seconds, 58°C for 15 seconds and 72°C for 20 seconds. PCR products were characterized by analyzing amplicon melt curve (95°C for 15 seconds, 68°C for 1 minute, 80°C for 15 seconds, 60°C for 15 seconds, then 60°C to 90°C with an increment of 0.2°C per second). No template and positive controls were included in all runs. All samples and controls were tested in triplicates.

Specificity of the PCR was confirmed by Sanger sequencing with both W-Specf/W-Specr primers for all samples that give at least 1/3 positive reaction. Positive reaction was defined by the presence of a PCR product with the same melting temperature than the positive control at the end of the thermocycling. Macrogen (Seoul, South Korea) performed both PCR product purification and sequencing off site to avoid contamination of our facilities with post-PCR amplicons. The sequences were used for phylogenetic analysis (accession numbers MK336794 - MK336806).

Data analysis

Human-biting rate was defined as the number of collected mosquitoes divided by the corresponding number of collection-nights. Poisson confidence intervals were calculated using the epitools package version 0.5–10 in R software. Human-biting rate for sensu stricto species in the Funestus, Maculatus and Leucopshyrus Groups was estimated using the relative proportion of the species in the corresponding group.

The limit of detection of the qPCR assay (LOD) was defined as the highest dilution (lowest concentration) that gave 100% of positive reactions. The performances of the two primer sets at low concentrations of Wolbachia were also compared by scoring the proportion of positive reactions as described previously 30, 34. Crossing-point (CP) values were determined using the regression algorithm of the analysis software of the PCR device (CFX Biorad Manager version 3.01, Biorad). CP values of standard samples in the serial-dilution experiments were used to set-up the standard curve of the assay. The best fit-line and the subsequent values of the slope and y-intercept were estimated by performing least-square analysis of the linear portion of the curve (Pearson’s coefficient r 2>0.990). PCR efficiency was estimated with the formula EFF = 10 (-1/slope)–1.

For the phylogenetic analysis, chimeric PCR products were detected with the DECIPHER software version 2.0 and excluded from subsequent analysis (4/17 samples with a positive PCR result). 16S ssuRNA sequences were blasted against the National Center for Biotechnology Information nucleotide database and the most similar sequence was downloaded. Reference Rickettsiales sequences were added and alignment was performed using the DECIPHER package version 2.10 in R software. DNA sequences were converted into RNA sequences and then aligned using the AlignSeqs() function set with default parameters in order to take into account base pairing and to use single-base and double-base substitution matrices. Tamura-Nei genetic distance model and neighbor-joining tree were computed with the ape package version 5.2 of the R software. There was 373 positions in the final dataset.

Ethical considerations

This project was approved through the ethics review committee on medical research involving human beings from Myanmar, Ministry of Health and Sports, Department of Medical Research (lower Myanmar): 73/Ethics 2014. All participants provided their written consent to participate in this study.

Results

qPCR assay validation for the detection of Wolbachia in mosquitoes

Optimal reaction conditions were similar for both primer sets: 58°C for primer annealing (range tested= 55–62°C), 2.5 mM of MgCl 2 (range tested= 2.5–4.5 mM) and 200 nM of each primers (range tested= 100–400 nM). In these conditions, PCR efficiency was 108 and 110% with the primer sets W-Specf/W-Specr and W-Specf/W16S respectively, and the linear dynamic spanned over six orders of magnitude ( r²=0.998 and 0.999) ( Table 1). There was a one-log decrease in the LOD of the assay when using W16S as a reverse primer instead of W-Specr, and the assay scored better at low concentrations of Wolbachia (16/18 and 12/18 positive reactions respectively, χ 2= 2.5714, P=0.109). Typical amplification and melt curves are shown in the Figure 2.

Table 1. Results of the serial dilution experiments.

Primers (%EFF, r 2) a Parameter Value of the parameter at the indicated dilution Score (%) e
Not diluted 10 -1 10 -2 10 -3 10 -4 10 -5 10 -6 10 -7
W-Specf/W-Specr
(108%, 0.998) 		Nb. pos. / Nb. tested b 9/9 9/9 9/9 9/9 9/9 7/9 5/9 0/9 12/18
(66%)
Mean CP value 18.81 21.26 24.68 28.01 30.73 34.48 34.95 -
Intra-assay SD c 0.07 0.04 0.03 0.10 0.25 1.64 0.33 -
Inter-assay SD d 0.11 0.03 0.02 0.09 0.25 1.45 0.71 -
W-Specf/W16S
(110%, 0.999) 		Nb. pos. / Nb. tested 9/9 9/9 9/9 9/9 9/9 9/9 7/9 0/9 16/18
(88%)
Mean CP value 17.80 20.71 23.98 27.29 29.90 33.40 34.86 -
Intra-assay SD 0.05 0.03 0.02 0.10 0.24 0.65 1.01 -
Inter-assay SD 0.01 0.06 0.14 0.18 0.09 0.10 1.26 -
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a %EFF : efficiency (EFF) of the PCR was calculated with the formula EFF = 10 (-1/slope) - 1 and expressed as a percentage. An efficiency of 100% corresponds to a slope of -3.32 and means that the number of amplicons doubles after each cycle of amplification; r 2: Pearson’s correlation coefficient expressing the intensity of the relationship between the logarithm of the concentration and the mean CP value. r 2 varies between 0 (no correlation) and 1 (perfect correlation), a value >0.990 testify of the linearity of the method (over a defined linear range) and allow an accurate quantification. r 2 and EFF have been calculated on the linear dynamic of each curve (bold cells).

b Nb. pos. / Nb. tested: number of positive reactions (amplification of the PCR DNA target) / total of reactions performed at a given dilution.

c Intra-assay SD : intra-assay standard deviation (SD), calculated as the average SD of the mean CP value measured for each dilution during the same experiment.

d Inter-assay SD : inter-assay standard deviation (SD), calculated as the SD of the means CP values measured during two independent experiments.

e score of the proportion of positive reactions at low concentrations of Wolbachia (score was calculated on dilutions 10 -5 and 10 -6); an example of the calculation of the score is given here : the maximum hit for the score is 18 reactions (9 at the dilution 10 -5, +9 at the dilution 10 -6), the score obtained with the primer pair W-Specf/W-Specr is 66% (12/18=(7+5)/18).

Figure 2. Typical result of the qPCR assay used for Wolbachia detection in mosquito samples.

Figure 2.

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A) W-Specf/W-Specr primers; B) W-Specf/W16S primers. Left panels show amplification curves and right panels show the melt curve of the PCR products. (*) primer dimers, (**) PCR DNA target.

Biodiversity of Anopheles mosquitoes

Four thousand seven hundreds forty-three Anopheles were collected during 500 person-nights of collection. We report the occurrence of 12 Anopheles taxa among which nine were groups of closely related or sibling species (Maculatus, Funestus, Jamesii, Leucosphyrus, Annularis, Barbirostris, Subpictus, Hyrcanus and Asiaticus Groups) and only three were sensu stricto species ( An. karwari, An. kochi and An. tessellatus). A subsample of 1098 mosquitoes in the Maculatus, Funestus and Leucosphyrus Groups were identified at the species level with molecular assays. The most frequently detected species were An. maculatus ( s.s.), An. sawadwongporni and An. pseudowillmori in the Maculatus Group, An. minimus ( s.s.), An. culicifacies B and An. jeyporiensis in the Funestus Group and An. baimaii in the Leucosphyrus Group ( Table 2).

Table 2. Village-collated human-biting rate estimates of Anopheles mosquitoes.

Group Species Human-biting rate estimate (95%CI) in the indicated dilution, expressed in number of bites/person/month
HD-3634 HG-369 LK-350 MK-3633 MK-3635 MM-3631 NT-361 TG-357 TP-339 WM-367
Annularis An. annularis
( s.l.) b 		40.2
(31.2-51.1)		 28.8
(21.2-38.2)		 24
(17.1-32.7)		 29.4
(21.8-38.9)		 4.2
(1.7-8.7)		 30
(22.3-39.6)		 40.2
(31.2-51.1)		 65.4
(53.7-78.9)		 7.2
(3.7-12.6)		 28.8
(21.2-38.2)
Asiaticus An. asiaticus ( s.l.) 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0.6
(0-3.3)		 0
(0-2.2)
Barbirostris An. barbirostris
( s.l.) b 		27
(19.7-36.1)		 2.4
(0.7-6.1)		 16.8
(11.2-24.3)		 3.6
(1.3-7.8)		 5.4
(2.5-10.3)		 10.2
(5.9-16.3)		 3.6
(1.3-7.8)		 24.6
(17.7-33.4)		 6
(2.9-11)		 2.4
(0.7-6.1)
Funestus An. aconitus b 3
(1-7)		 1.2
(0.1-4.3)		 9.6
(5.5-15.6)		 6
(2.9-11)		 3.6
(1.3-7.8)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)
An. culicifacies A c 9
(5-14.8)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 3.6
(1.3-7.8)		 6
(2.9-11)		 4.2
(1.7-8.7)		 1.8
(0.4-5.3)		 0
(0-2.2)
An. culicifacies B 36.6
(28-47)		 7.2
(3.7-12.6)		 0
(0-2.2)		 73.8
(61.3-88.1)		 12.6
(7.8-19.3)		 123.6
(107.3-141.7)		 94.2
(80-110.1)		 7.8
(4.2-13.3)		 1.8
(0.4-5.3)		 26.4
(19.2-35.4)
An. harrisoni 3
(1-7)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 4.2
(1.7-8.7)		 0
(0-2.2)		 0.6
(0-3.3)
An. jeyporiensis c 3
(1-7)		 1.2
(0.1-4.3)		 12.6
(7.8-19.3)		 6
(2.9-11)		 44.4
(34.9-55.7)		 3.6
(1.3-7.8)		 3
(1-7)		 78.6
(65.7-93.3)		 3.6
(1.3-7.8)		 0
(0-2.2)
An. minimus a 161.4
(142.7-181.9)		 46.8
(37-58.4)		 240
(217.1-264.7)		 185.4
(165.3-207.3)		 96
(81.7-112.1)		 167.4
(148.3-188.2)		 141.6
(124.1-160.9)		 259.2
(235.3-284.8)		 141
(123.5-160.2)		 52.2
(41.8-64.4)
An. varuna 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 4.2
(1.7-8.7)		 0
(0-2.2)		 0
(0-2.2)
Hyrcanus An. hyrcanus
( s.l.) c 		0.6
(0-3.3)		 0.6
(0-3.3)		 2.4
(0.7-6.1)		 0.6
(0-3.3)		 0
(0-2.2)		 0
(0-2.2)		 2.4
(0.7-6.1)		 0.6
(0-3.3)		 0
(0-2.2)		 0
(0-2.2)
Jamesii An. jamesii ( s.l.) c 140.4
(123-159.6)		 27
(19.7-36.1)		 9
(5-14.8)		 15.6
(10.2-22.9)		 8.4
(4.6-14.1)		 15.6
(10.2-22.9)		 46.2
(36.5-57.7)		 141.6
(124.1-160.9)		 119.4
(103.4-137.2)		 44.4
(34.9-55.7)
Kochi An. kochi 37.8
(29-48.4)		 1.8
(0.4-5.3)		 3
(1-7)		 0.6
(0-3.3)		 55.8
(45-68.4)		 6.6
(3.3-11.8)		 15.6
(10.2-22.9)		 36
(27.5-46.3)		 54
(43.4-66.4)		 2.4
(0.7-6.1)
Leucosphyrus An. baimaii a 31.8
(23.8-41.6)		 1.8
(0.4-5.3)		 24.6
(17.7-33.4)		 10.2
(5.9-16.3)		 13.2
(8.3-20)		 18.6
(12.6-26.4)		 15
(9.7-22.1)		 24
(17.1-32.7)		 11.4
(6.9-17.8)		 6.6
(3.3-11.8)
An. dirus a 2.4
(0.7-6.1)		 0
(0-2.2)		 0
(0-2.2)		 1.2
(0.1-4.3)		 0
(0-2.2)		 1.8
(0.4-5.3)		 0
(0-2.2)		 1.8
(0.4-5.3)		 0.6
(0-3.3)		 0
(0-2.2)
An. introlatus 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0.6
(0-3.3)		 0
(0-2.2)
Maculatus An. maculatus a 241.2
(218.2-266)		 40.8
(31.7-51.7)		 88.2
(74.5-103.7)		 26.4
(19.2-35.4)		 59.4
(48.3-72.3)		 61.8
(50.4-75)		 226.8
(204.5-250.9)		 1112.4
(1062.3-
1164.2)		 315
(288.6-343.1)		 39
(30.1-49.7)
An.
pseudowillmori b 		6.6
(3.3-11.8)		 3.6
(1.3-7.8)		 0
(0-2.2)		 4.2
(1.7-8.7)		 15
(9.7-22.1)		 1.2
(0.1-4.3)		 3.6
(1.3-7.8)		 19.2
(13.1-27.1)		 0
(0-2.2)		 0
(0-2.2)
An. rampae 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 0.6
(0-3.3)		 0
(0-2.2)		 0
(0-2.2)		 0
(0-2.2)		 38.4
(29.6-49)		 0
(0-2.2)		 0
(0-2.2)
An.
sawadwongporni a 		42
(32.7-53.1)		 12.6
(7.8-19.3)		 25.2
(18.2-34.1)		 21.6
(15.1-29.9)		 3.6
(1.3-7.8)		 16.8
(11.2-24.3)		 66
(54.2-79.5)		 307.2
(281.2-335)		 42
(32.7-53.1)		 12
(7.3-18.5)
Maculatus-
related		 An. karwari c 15
(9.7-22.1)		 2.4
(0.7-6.1)		 0
(0-2.2)		 2.4
(0.7-6.1)		 1.2
(0.1-4.3)		 0
(0-2.2)		 3
(1-7)		 11.4
(6.9-17.8)		 3
(1-7)		 0.6
(0-3.3)
Subpictus An. subpictus
( s.l.) c 		79.8
(66.8-94.6)		 40.8
(31.7-51.7)		 41.4
(32.2-52.4)		 13.2
(8.3-20)		 5.4
(2.5-10.3)		 58.8
(47.7-71.7)		 137.4
(120.2-156.4)		 24.6
(17.7-33.4)		 182.4
(162.5-204.1)		 119.4
(103.4-137.2)
Tessellatus An. tessellatus c 3.6
(1.3-7.8)		 0
(0-2.2)		 24
(17.1-32.7)		 2.4
(0.7-6.1)		 0
(0-2.2)		 14.4
(9.2-21.4)		 8.4
(4.6-14.1)		 1.2
(0.1-4.3)		 3.6
(1.3-7.8)		 0.6
(0-3.3)
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a primary malaria vectors; b secondary malaria vectors, c efficient malaria vector species in some areas that were never reported infected with human malaria parasites on the Thailand-Myanmar border 25. Human-biting rates of sensu stricto anopheline species in the Funestus, Maculatus and Leucosphyrus Groups were estimates from the relative proportion of each species in the corresponding Group assessed with molecular assays.

Detection of Wolbachia DNA in malaria vectors

The presence of Wolbachia DNA was assessed in six Anopheles species namely An. maculatus ( s.s.), An. sawadwongporni, An. pseudowillmori (Maculatus Group), An. minimus ( s.s.) (Funestus Group, Minimus Complex), An. dirus ( s.s.) and An. baimaii (Leucosphyrus Group, Dirus Complex). Wolbachia DNA was detected in 13/370 samples ( Table 3). Eight unique 16S rRNA sequences were identified ( Figure 3). 16S rRNA sequences clustered with that of Wolbachia strains in the supergroups B, D and F ( Figure 4).

Table 3. Results of the screening for natural Wolbachia infections in the ten villages.

Group Species Nb. pos / Nb. tested ( Wolbachia supergroup) in the indicated village and species
HD-3634 HG-369 LK-350 MK-3633 MK-3635 MM-3631 NT-361 TG-357 TP-339 WM-367
Funestus An. minimus 0/10 2/10 (F) 0/10 0/10 0/10 1/10 (D) 0/10 1/10 (D) 0/10
Maculatus An. maculatus 0/10 0/10 0/10 2/10 (B) 0/10 0/10 1/10 (F) 0/11 1/9 (B)
An. pseudowillmori 0/1 0/1 0/1 1/7 (B) 0/1
An. sawadwongporni 0/8 0/3 0/9 0/10 1/1 (B) 0/10 0/11 0/10 0/4 0/2
Leucopshyrus An. baimaii 0/10 0/2 0/10 1/10 (D) 0/11 0/10 0/10 0/10 1/16 (B) 0/10
An. dirus 0/4 0/2 0/2 0/3 1/1 (B)
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Figure 3. Multiple alignment of 16S RNA sequences used to build the Tamura-Nei genetic distance model and neighbor-joining tree.

Figure 3.

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Figure 4. Phylogenetic analysis based on the alignment of a conserved region of the 16S rRNA gene using Wolbachia-specific primer pair W-Specf/W-Specr.

Figure 4.

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Sequences of the PCR products were blasted against the NCBI nucleotide database and the most similar result was downloaded. A phylogenetic tree was reconstructed using a Tamura-Nei genetic distance model and neighbor joining. Sequences from other non- Wolbachia proteobacteria were also included, and the sequence from Rickettsia japonica was used as the reference outgroup. There was 373 positions in the final dataset. Nodes with bootstrap support <50% were collapsed. Study samples were labeled with the host name and the study village, and the accession number reported into the brackets. Formally named Wolbachia strains were labeled with their abbreviation: wNo is a symbiont of Drosophila simulans, wCne of Ctenocephalides felis, wAlbB of Aedes albopictus, wAnga of An. gambiae, wMel of Drosophila melanogaster, wPeJe1 of Penicillidia jenynsii and wBru of Brugia malayi.

Crossing-point values ranged from 31.0 to 40.6 amplification cycles and Wolbachia DNA titers were generally close or below the limit of detection of the assay (only one sample gave 3/3 positive reactions) ( Table 4).

Table 4. qPCR results of the Wolbachia-infected samples detected during the screening.

Sample ID Village Species Nb pos CP1 CP2 CP3 Supergroup
1 HG-369 An. minimus 1 35.8 F
2 HG-369 An. minimus 1 33.0 F
3 MK-3633 An. baimaii 1 35.6 D
4 MK-3635 An. maculatus 1 34.3 B
5 MK-3635 An. maculatus 1 34.3 B
6 MK-3635 An. pseudowillmori 1 37.6 B
7 MK-3635 An. sawadwongporni 2 34.5 32.8 B
8 NT-361 An. minimus 3 36.8 35.8 36.6 D
9 TG-357 An. maculatus 1 34.2 F
10 TP-339 An. baimaii 3 33.0 31.0 32.3 B
11 TP-339 An. dirus 1 34.1 B
12 TP-339 An. minimus 1 40.6 D
13 WM-367 An. maculatus 1 32.6 B
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Discussion

Wolbachia DNA was detected for the first time in Southeast Asian malaria vectors, including An. maculatus ( s.s.), An. sawadwongporni, An. pseudowillmori (Maculatus Group), An. dirus ( s.s.) and An. baimaii (Dirus Complex, Leucosphyrus Group).

CP values reported in this study suggest that Wolbachia DNA titers were very low, usually close or below the limit of detection of our assay. This result is not compatible with the integration of Wolbachia DNA in the mosquito genome, which would have given much lower CP values 35. Important precautions were taken to ensure the quality of our molecular data 36. This was the first study on Wolbachia in our facilities. The 16S DNA sequences detected in the screened samples were different from that of the reference material, hence excluding cross-contaminations. In addition, all experiments were conducted with the real-time PCR technology (which allows amplification and detection of the PCR DNA target in a closed system) and great care was taken to perform all handlings of PCR products off site. These precautions, combined with the good laboratory practices relevant to molecular diagnosis (eg. dedicated facilities with unidirectional workflow, experiment conducted by qualified laboratory technicians and appropriate quality controls), drastically limited the risk of false positive by contamination. The risk of false positive results due to low specificity of the assay was ruled out by sequencing the PCR product in all positive samples. It is probable that some results were falsely negative due to limited sensitivity, given that most positive samples were infected at a density close of below the detection of the assay. In this study, we have shown that using the W16S as a reverse primer increases the analytical sensitivity of the qPCR assay in the optimal reaction conditions. However, in the absence of a priori data on the Wolbachia DNA sequences detected in this study, we selected the W-Specf/W-Specr primers to perform the screening because of their availability to detect a wide variety of Wolbachia infecting insects and to establish phylogenetic relationships among field isolates 32. Molecular phylogeny based on 16S rRNA sequences revealed a high diversity of Wolbachia strains, which belonged to different lineages than those recently reported in the African malaria vectors 19– 24. Eight out of thirteen sequences reported in this study were unique. The DNA extracts were also used to assess Plasmodium infection rates in the mosquito population (data not shown), precluding multi locus sequence typing of the Wolbachia strains because there was no material remaining after the screening.

The significance of these findings regarding the biology and ecology of Wolbachia-Anopheles interactions must be interpreted cautiously as the detection of low titers of Wolbachia DNA by PCR is not unequivocal of an actual symbiosis between Wolbachia and the mosquito. The detection of Wolbachia in the supergroup D and F suggests that some DNA extracts were contaminated with Wolbachia endosymbionts of filarial nematodes rather than reflecting actual Wolbachia infections in mosquitoes. Chrostek and Gerth further argued that the high diversity of Wolbachia sequences combined with the very low titers detected was incompatible with the notion of a stable, intraovarially-transmitted Wolbachia symbiont in An. gambiae 37. Given that most arthropods are infected with Wolbachia 38, we cannot exclude that the DNA sequences detected in this study come from some sort of environmental contamination. An alternative explication could be that horizontal transfers of Wolbachia happen at a much higher frequency than previously thought, for example via plants 39 or via ectoparasitic mites 40, 41. Additional experiments would be of great interest to demonstrate actual infection, e.g. showing intracellular localization of the sequences and maternal transmission of the bacteria. Finally, we did not assess the effects of the presence of Wolbachia DNA on the phenotype of mosquitoes and dynamics of malaria transmission. In Kayin state, malaria transmission is low, seasonal and unstable. Plasmodium infection rate is usually less than 1% and often nil in the mosquito populations 25. Therefore, it was not possible to establish direct correlations between Plasmodium and the presence of Wolbachia DNA in the mosquito vectors. In this setting, the effect of possible Wolbachia infections on malaria transmission may be better assessed by performing artificial infections of field-collected mosquitoes with Plasmodium malaria parasites.

Conclusion

The detection Wolbachia DNA in malaria vectors from Kayin state warrants further investigations to understand better the ecology and biology of Anopheles- Wolbachia interactions in Southeast Asia.

Data availability

The data is available upon request to the Mahidol Oxford Tropical Medicine Research Unit Data Access Committee ( http://www.tropmedres.ac/data-sharing) and following the Mahidol Oxford Tropical Medicine Research Unit data access policy ( http://www.tropmedres.ac/_asset/file/data-sharing-policy-v1-0.pdf).

Acknowledgments

We thank to the communities from the study villages for their support to the study, and to the SMRU Entomology Department for their work. Wolbachia-infected reference samples were kindly provided by Dr. Lauren Carrington from the Oxford University Clinical Research Unit, Ho Chi Minh City, Vietnam. SMRU is part of the Mahidol Oxford University Research Unit, supported by the Wellcome Trust of Great Britain.

Funding Statement

This work was supported by the Wellcome Trust [101148]; the Bill and Melinda Gates Foundation [GH OPP 1081420] and the Global Fund [THA-M-DDC].

The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.

[version 4; peer review: 2 approved, 1 approved with reservations]

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Wellcome Open Res. 2019 Dec 3. doi: 10.21956/wellcomeopenres.17037.r36934

Reviewer response for version 4

Ewa Chrostek 1

Thank you for the revisions. The problematic "Wolbachia infection" has been mostly replaced, except from one place in the discussion. Also, I disagree with calling the Wolbachia DNA from previous reports an infection in the abstract and the introduction of this paper. In my opinion, this study is not defficient compared to the previous ones on Wolbachia in Anopheles. It only has the potential to be more carefully worded.

I agree with the author's interpretation of the qPCR results. However, as primer dimers contribute to the total SYBR green fluorescence in the qPCR reaction this assay cannot be quantitative. Also, only running the qPCR products on the agarose gel can reveal if the bands can indeed be distinguished by a conventional PCR. As the authors do not aim at PCR-identification or qPCR quantification of Wolbachia here this is not crucial. It is curious though, and suggests that the qPCR assay could be optimized further.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard.

Wellcome Open Res. 2019 Oct 15. doi: 10.21956/wellcomeopenres.16939.r36677

Reviewer response for version 3

Ewa Chrostek 1

In this study, the authors provide evidence for qPCR amplification of a fragment of Wolbachia 16S sequence from 13 out of 370 field-collected Southeast-Asian mosquitoes. However, a single, low level, hard to amplify (despite extensive optimisation) fragment of Wolbachia rDNA does not provide sufficient evidence for the Wolbachia infection in these malaria vectors. The environment is severely contaminated with Wolbachia as most arthropods are infected with this symbiont (Weinert et al. 2015 1). To show an infection, additional experiments are necessary, e.g. showing intracellular localization of the sequences and maternal transmission of the bacteria (see Chrostek and Gerth 2019 2 for further discussion). I suggest changing the wording throughout, from “natural Wolbachia infections” to “Wolbachia 16S/wSpec sequence amplification”. I also recommend changing the title to: “Detection of low-density Wolbachia 16S sequences from malaria vectors in Kayin state, Myanmar”.

In the results section, the authors identify the supergroups D and F Wolbachia sequences as likely environmental/parasitic contaminants (6 out of 13 positive samples). It is unclear why they do not use the same caution while interpreting data on the remaining 7 sequences from supergroup B.

Figure 2 shows an interesting property of wSpec primer set. The amplification from the positive and negative samples produces similar amplification curves (Fig. 2A, green and red lines) with very similar CPs (crossing-points), that can be distinguished from each other by the shape of their melt curved (Fig. 2B). This figure shows, that when using wSpec primer set for a PCR, negative samples produce an amplicon as well. Whether the positive and negative amplicons can be distinguished by agarose gel electrophoresis, and therefore if classical PCR with wSpec has any diagnostic value when used under this conditions, remain to be determined. The comment on this in the text would help future studies embarking on Wolbachia identification projects.

Minor comments:

  1. The information on the origin of the laboratory-reared wMel-infected A. aegypti is missing. Were they made by the authors or are they the published strains?

  2. The sequence of W16S primer, the amplicon sizes for both qPCR reactions and the rationale behind trying both primer sets should be added.

  3. In the discussion, p7/28, first line: the integration into the chromosome would have produced “higher CP values”, and not as it is - “lower”.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to state that I do not consider it to be of an acceptable scientific standard, for reasons outlined above.

References

  • 1. : The incidence of bacterial endosymbionts in terrestrial arthropods. Proc Biol Sci.2015;282(1807) : 10.1098/rspb.2015.0249 20150249 10.1098/rspb.2015.0249 [DOI] [PMC free article] [PubMed] [Google Scholar]
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Wellcome Open Res. 2019 Nov 1.
Victor Chaumeau 1

We thank to reviewer for her useful comments on the manuscript. The wording “natural infection” was changed in the title and throughout the manuscript as per reviewer’s suggestion. In addition, part of the discussion was rewritten and suggested reference was added in order to better emphasize this study limitation in the revised version of the manuscript. The possibility of environmental contamination for all sequences including that of the supergroup B was explicitly discussed in the Discussion section. The amplicon produced in no-template controls and negative samples certainly results from the formation of primer dimers, which are unlikely to reach the size of 438 bp. Therefore, it should be possible to discriminate between primer dimers and amplification of the PCR DNA target using gel electrophoresis. Nevertheless, the reviewer is right to question the value of conventional PCR in the field of molecular diagnostic given the high risk of false positive by contamination [1]. This issue has been extensively discussed in the manuscript and we do not wish to add more details in the current version of the manuscript. Point-by-point answers to specific comments are listed below:

1.         The information on the origin of the laboratory-reared wMel-infected A. aegypti is missing. Were they made by the authors or are they the published strains?

Wolbachia-infected reference samples were kindly provided by Dr. Lauren Carrington from the Oxford University Clinical Research Unit, Ho Chi Minh City, Vietnam. Corresponding reference [2] was added in the methods section of the revised version of the manuscript.

2.         The sequence of W16S primer, the amplicon sizes for both qPCR reactions and the rationale behind trying both primer sets should be added.

The sequence of W16S primer, the amplicon sizes for both qPCR reactions and the rationale behind trying both primer sets were added in the Methods section in the revised version of the manuscript.

3.         In the discussion, p7/28, first line: the integration into the chromosome would have produced “higher CP values”, and not as it is - “lower”.

The integration of Wolbachia genome into the mosquito chromosome would have resulted in a higher copy number of PCR DNA target in the sample (several millions versus a few dozens), and therefore in a lower CP value given the negative correlation that exists between the CP value and the logarithm of the concentration of PCR DNA target in the sample.

1.            Apfalter P, Reischl U, Hammerschlag MR. In-house nucleic acid amplification assays in research: how much quality control is needed before one can rely upon the results? J Clin Microbiol. 2005;43 12:5835-41; doi: 10.1128/jcm.43.12.5835-5841.2005. http://dx.doi.org/10.1128/jcm.43.12.5835-5841.2005.

2.            Carrington LB, Tran BCN, Le NTH, Luong TTH, Nguyen TT, Nguyen PT, et al. Field- and clinically derived estimates of Wolbachia-mediated blocking of dengue virus transmission potential in Aedes aegypti mosquitoes. Proc Natl Acad Sci U S A. 2018;115 2:361-6; doi: 10.1073/pnas.1715788115. http://dx.doi.org/10.1073/pnas.1715788115.

Wellcome Open Res. 2019 Sep 6. doi: 10.21956/wellcomeopenres.16771.r35969

Reviewer response for version 2

Francesco Baldini 1

The second version of this manuscript provides greater clarity and detailed methodological information. Limitations of the study are also discussed. Overall the manuscript contains novel insights on natural Wolbachia strains in Anopheles species from Myanmar, calling for further investigations on the role of these endosymbionts on the biology of malaria mosquitoes.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard.

Wellcome Open Res. 2019 Jul 22. doi: 10.21956/wellcomeopenres.16771.r35968

Reviewer response for version 2

Thomas Walker 1

Figure 2 has been added to show the typical result of the qPCR asay used for Wolbachia detection in mosquito samples. The Wspec results (panel A) show that CP values were generated from no template controls (some below 35) and melt curve analysis was required to differentiate primer dimers from genuine target amplification. With Wolbachia gene qPCR I would not have expected no template control CPs to come up at below 35 cycles and this could be due to trying to amplify a 16S gene fragment >400 base pairs (the first report of this to my knowledge). According to Table 1, the threshold CP value for using the Wspec primer set appears to be between 30-33 cycles given only 7/9 of the 10 -5 dilution amplified. This would be further supported by the CP of 10 -6 (34.95 +/-1.64) not being statistically higher that to the 10 -5 dilution (34.48 +/-0.33). The authors rightly conclude that all of the samples in this study are at threshold detection (CP values >31 cycles) and the authors correctly used both melt curve analysis and sequencing to confirm this was genuine amplification. Further details of the qPCR analysis strengthens the manuscript but I’m not entirely sure how the authors estimated bacterial load. The sentence ‘Given that DNA was extracted from whole mosquitoes, it was possible to estimate the bacterial load in single mosquitoes without using a calibrator to normalize the signal’ needs further clarification, particularly given the title has changed to include reference to ‘low density strains’. The CP values would suggest low density infections but in order to make the comparison to wMel in Aedes aegypti you would need to account for body size and/or extraction efficiency as these factors will influence Wolbachia CP values and therefore estimating bacterial load. Lab-reared Aedes aegypti adults will likely be larger in size than wild caught Anopheles species so that comparison is not possible without either normalising to host genes or measuring the total DNA extracted. I also would suggest a better measure of prevalence rates would have resulted from using the Wspec primer set in the conventional PCR format (Werren & Windsor, 2000 1) which has been routinely used to screen mosquito populations.  

Figure 4 shows the phylogeny of the Wolbachia 16S gene and it appears that the sequences are quite diverse within individual species (eg. An. maculates appearing to have four different strains) and the same strains appear across multiple species (eg. An. minimus and An. baimaii). Although Wolbachia superinfections exist in mosquito species, having the same strain of Wolbachia (based on 16S sequences) in multiple species would need confirmation from additional gene sequencing as this seems unlikely for endosymbiotic Wolbachia bacteria. The discussion does now contain an explanation that no material was left for MLST due to assessing Plasmodium infection rates despite having 50 μL of eluted DNA. Overall the reliance on only 16S sequences (some of which appear to have identical sequences across multiple Anopheles species) is still problematic in my opinion and I would think a title that contains ‘genetically diverse’ is inappropriate based on sequencing of only one Wolbachia gene fragment.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard, however I have significant reservations, as outlined above.

References

  • 1. : Wolbachia infection frequencies in insects: evidence of a global equilibrium?. Proc Biol Sci.2000;267(1450) : 10.1098/rspb.2000.1139 1277-85 10.1098/rspb.2000.1139 [DOI] [PMC free article] [PubMed] [Google Scholar]
Wellcome Open Res. 2019 Sep 18.
Victor Chaumeau 1

We thank to reviewer for his useful comments on the manuscript. Reviewer’s concern on the comparison of Wolbachia DNA titers measured in wild Anopheles and laboratory-reared Aedes aegypti was addressed by removing the corresponding sentences in the Methods, Results and Discussion sections. Our data clearly demonstrate that wild Anopheles are infected with very low Wolbachia densities given the high CP values of the screened samples (>30 cycles) and the very low CP values of the reference Aedes aegypti samples (<19 cycles). The reference to “low-density infections” in the title is therefore valid, and does not overlook low densities as a specific feature of the strains detected in this study. We agree with reviewer’s comment on the limitation of not having additional sequence data on other markers, and regret that it was not possible to generate such data in the current study. Although it not possible to assess accurately the phylogenic relationships between different Wolbachia strains only with 16S rRNA sequences, our data clearly demonstrate that the Wolbachia strains detected in this study are genetically diverse. Therefore, we wish to keep the title of the manuscript unchanged.

Wellcome Open Res. 2019 Mar 4. doi: 10.21956/wellcomeopenres.16370.r34658

Reviewer response for version 1

Francesco Baldini 1

This work identified Wolbachia strains in Anopheles species from Myanmar. To my knowledge this is the first evidence of identification of Wolbachia infection in Anopheles mosquito populations in Asia. This work opens novel questions on the potential role of these infections on the vectorial capacity of the vector host. Indeed, recently identified natural Wolbachia infections in Anopheles species in Africa have been shown to potentially influence the vectorial capacity of the infected vectors.

The authors state that Wolbachia infections were identified by using quantitative PCR approach followed by sequencing of the Wolbachia 16S gene, although some details are missing and the methodology should be clarified. Phylogenetic analysis is also not completely detailed. Specific comments to the manuscript are listed below.

Introduction:

The authors state that cytoplasmic incompatibility (CI) ‘enhances the “vertical” transmission of Wolbachia’. From my knowledge this is incorrect, as CI does not directly affect “vertical” transmission, but rather increases ‘indirectly’ the fitness of the progeny of Wolbachia infected mothers.

The authors state that ‘Shaw et al. observed a negative correlation between Wolbachia infection and the development of P. falciparum oocysts in naturally blood-fed females’. This is not completely correct, as in this work the authors have quantified the prevalence of P. falciparum in resting blood fed females inside house 5 days after collection/blood feeding, without any prior knowledge on the infectious status of the mosquitoes; thereby, the stage of parasite infection (oocyst or sporozoite) was not investigated.  

Methods:

Authors indicate Shaw et al. and Gomes et al. as references for the qPCR using W-Spec primers against Wolbachia 16S. This is where I am getting confused, as Shaw et al. did not use qPCR and Gomes et al. use a different primer set for qPCR. The methodology should be clarified.

If W-Spec primers were used, the expected product size is >400 base pairs (bp); this bp size is often too large for qPCR, as large amplicons tend to produce secondary structures during the dissociation steps, thus resulting in multiple melting peaks. If possible, it would be informative to provide more details on the optimization of this assay (in case additional reagents were added, for example) and to show the dissociation profile of the obtained amplicons, as this would enable troubleshooting of the technique if others will try to replicate the work and/or use the same methodology. Also, it should be specified how sequencing was performed, e.g. direct purification after qPCR (how?), which primer was used, etc.

In the phylogenetic analysis the authors should state what was the sequence size used to build the tree. Novel obtained sequences should also be uploaded and their unique identifier indicate in the article.

Statistical analysis used in Table 1-3 should be indicated (if any).

Results:

The authors state the ‘we assessed species diversity, Plasmodium and Wolbachia infection rates in these Groups’. This is incorrect as Plasmodium infection rates are not shown.

Captions in Table 1-3 are missing information on what each column indicates. Although these tables can be generally informative, I wonder if showing species diversity using pie-charts (for each species group) over imposed on the map in Figure 1 would provide a more direct illustration of the species composition and abundances of the Anopheles species in the study area.

As indicated in the methods, the size of the sequence used for Figure 2 should be indicated. It would be informative to include the alignment use for the tree figure.

Figure 3b shows the overall prevalence of Wolbachia in different villages without specifying the species, so I am not sure what is the purpose of illustrating the result in this way. If this is too show that some villages have higher prevalence over others this should be indicated only if statistical analysis supports it (although I doubt this is the case if species distribution is included as a variable).

The authors state that ‘Crossing-point values ranged from 31.0 to 40.6 amplification cycles. Infected specimens were generally infected at a density close or below the limit of detection of the assay (only one sample gave 3/3 positive reactions).’ More details should be given regarding the rational for inclusion (or exclusion) of an infected/amplified sample; it is not clear to me if ‘reactions’ refers to technical replicates in the same qPCR run or in different qPCR assays. This should be described with more details. It would also be informative to normalize the quantity of the amplified Wolbachia 16S using a mosquito housekeeping gene, for example. Indeed, as ‘density [was] close or below the limit of detection of the assay’ normalization would provide information on the likelihood of false negatives in samples, as you would expect if the total DNA is very low (for example due to inefficient DNA extraction).

Discussion:

The authors state that ‘Our data and African studies confirm that the occurrence of natural Wolbachia infections has been underestimated in malaria mosquitoes.’ As direct assessment of Wolbechia prevalcence on samples previously identified as uninifected was not performed here (nor in African samples) it cannot be ruled out that previous Wolbachia negative samples were not true negatives, so this work (and others) only suggest possible underestimation in previous works, as they have not directly confirmed it.

The authors state that ‘It was not possible to study more in detail the phylogeny of Wolbachia strain detected in this study by multi locus sequence typing because of the lack of DNA extracts after the screening.’ It is not clear to me if any attempts were made at all or not. If so, please give more details on the targeted genes and discuss why these could not be amplified.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard, however I have significant reservations, as outlined above.

Wellcome Open Res. 2019 Jul 3.
Victor Chaumeau 1

We thank to the reviewer for his useful feedback on the manuscript. Additional details on the qPCR assay used for Wolbachia detection and on the phylogenetic analysis were added to the revised manuscript. Answer to point-specific comments are given below:

 

  • The authors state that cytoplasmic incompatibility (CI) ‘enhances the “vertical” transmission of Wolbachia’. From my knowledge this is incorrect, as CI does not directly affect “vertical” transmission, but rather increases ‘indirectly’ the fitness of the progeny of Wolbachia infected mothers.

Reviewer’s comment has been addressed in the revised version of the manuscript.

  • The authors state that ‘Shaw et al. observed a negative correlation between Wolbachia infection and the development of P. falciparum oocysts in naturally blood-fed females’. This is not completely correct, as in this work the authors have quantified the prevalence of P. falciparum in resting blood fed females inside house 5 days after collection/blood feeding, without any prior knowledge on the infectious status of the mosquitoes; thereby, the stage of parasite infection (oocyst or sporozoite) was not investigated.

Reviewer’s comment has been addressed in the revised version of the manuscript.

  • Authors indicate Shaw et al. and Gomes et al. as references for the qPCR using W-Spec primers against Wolbachia 16S. This is where I am getting confused, as Shaw et al. did not use qPCR and Gomes et al. use a different primer set for qPCR. The methodology should be clarified.

In this study, we adapted an in-house real-time PCR assay with legacy primers describe previously (W-Spec forward and reverse primers). More details and appropriate reference were added in the paragraph on Wolbachia detection in the Methods in section.

  • If W-Spec primers were used, the expected product size is >400 base pairs (bp); this bp size is often too large for qPCR, as large amplicons tend to produce secondary structures during the dissociation steps, thus resulting in multiple melting peaks. If possible, it would be informative to provide more details on the optimization of this assay (in case additional reagents were added, for example) and to show the dissociation profile of the obtained amplicons, as this would enable troubleshooting of the technique if others will try to replicate the work and/or use the same methodology. Also, it should be specified how sequencing was performed, e.g. direct purification after qPCR (how?), which primer was used, etc.

Additional information on the validation of the assay and sequencing of the PCR products were added to the revised version of the manuscript as per reviewer’s suggestion.

  • In the phylogenetic analysis the authors should state what was the sequence size used to build the tree. Novel obtained sequences should also be uploaded and their unique identifier indicate in the article.

The sequence size used to build the tree was added in the figure legend in the revised version of the manuscript. Accession number are given in the method section.

  • Statistical analysis used in Table 1-3 should be indicated (if any).

Tables 1-3 were replaced by a single table presenting human-biting rate instead of relative proportion of each mosquito species in the corresponding group. Appropriate definition of the statistics used to calculate human-biting rate are given in the Methods section.

  • The authors state the ‘we assessed species diversity, Plasmodium and Wolbachia infection rates in these Groups’. This is incorrect as Plasmodium infection rates are not shown.

The inaccurate statement was removed from the revised version of the manuscript.

  • Captions in Table 1-3 are missing information on what each column indicates. Although these tables can be generally informative, I wonder if showing species diversity using pie-charts (for each species group) over imposed on the map in Figure 1 would provide a more direct illustration of the species composition and abundances of the Anopheles species in the study area.

Table 1-3 were removed from the manuscript and replaced by a single table showing human-biting rate estimates collated by village and species. Given the number of study villages and diversity of Anopheles mosquitoes, we do not think that figuring multiple pie-charts on the map will improve the readability of the data.

  • As indicated in the methods, the size of the sequence used for Figure 2 should be indicated. It would be informative to include the alignment use for the tree figure.

The sequence alignment was added to the revised version of the manuscript and the number of positions in the final dataset used to build the tree was added to the figure legend of the revised version of the manuscript.

  • Figure 3b shows the overall prevalence of Wolbachia in different villages without specifying the species, so I am not sure what is the purpose of illustrating the result in this way. If this is too show that some villages have higher prevalence over others this should be indicated only if statistical analysis supports it (although I doubt this is the case if species distribution is included as a variable).

The reviewer is right to question the relevance of our prevalence data. Given the low sample size and the diversity of Wolbachia strains and Anopheles species, prevalence data were removed from the revised version of the manuscript.

  • The authors state that ‘Crossing-point values ranged from 31.0 to 40.6 amplification cycles. Infected specimens were generally infected at a density close or below the limit of detection of the assay (only one sample gave 3/3 positive reactions).’ More details should be given regarding the rational for inclusion (or exclusion) of an infected/amplified sample; it is not clear to me if ‘reactions’ refers to technical replicates in the same qPCR run or in different qPCR assays. This should be described with more details. It would also be informative to normalize the quantity of the amplified Wolbachia 16S using a mosquito housekeeping gene, for example. Indeed, as ‘density [was] close or below the limit of detection of the assay’ normalization would provide information on the likelihood of false negatives in samples, as you would expect if the total DNA is very low (for example due to inefficient DNA extraction).

It is common that at low parasite concentration, only some replicates give a positive result because the distribution of the DNA template in the reaction tube follow a Poisson distribution (Sterkers, Varlet-Marie et al. 2010, Stahlberg and Kubista 2014, Chaumeau, Andolina et al. 2016). This observation does not challenge the validity of our results. A clear statement that some Wolbachia infected sample have probably been missed because bacterial density observed in Anopheles are close or below the limit of detection of the assay that give 95% positive reaction. Misleading interpretations on the prevalence of Wolbachia infection were removed given the small sample size and the possibility of false negative.

 

  • The authors state that ‘Our data and African studies confirm that the occurrence of natural Wolbachia infections has been underestimated in malaria mosquitoes.’ As direct assessment of Wolbechia prevalcence on samples previously identified as uninifected was not performed here (nor in African samples) it cannot be ruled out that previous Wolbachia negative samples were not true negatives, so this work (and others) only suggest possible underestimation in previous works, as they have not directly confirmed it. The authors state that ‘It was not possible to study more in detail the phylogeny of Wolbachia strain detected in this study by multi locus sequence typing because of the lack of DNA extracts after the screening.’ It is not clear to me if any attempts were made at all or not. If so, please give more details on the targeted genes and discuss why these could not be amplified.

The methodology used for Wolbachia detection in this study was described into detail in the Methods section and we did not attempt additional experiments on Wolbachia than that described in the manuscript. In addition to Wolbachia detection and molecular identification of the mosquito species, sample were also screened for Plasmodium infection. There was not DNA material to perform additional experiment after the screening with the W-Specf/ W-Specr primers.

Wellcome Open Res. 2019 Feb 28. doi: 10.21956/wellcomeopenres.16370.r34878

Reviewer response for version 1

Thomas Walker 1

The authors present an interesting study in which Wolbachia strains were detected in Anopheles species from Myanmar. This study is particularly timely given the recent discoveries of natural Wolbachia strains in Anopheles malaria vectors in Sub-Saharan Africa and evidence that these natural endosymbiotic bacteria could be influencing malaria parasite infection prevalence in wild mosquito populations. The study provides evidence for Wolbachia infections using amplification and sequencing of the Wolbachia 16S gene although more clarity is needed on which primer set was used given the authors report undertaking qrtPCR and sequencing of PCR products. The manuscript would be significantly improved with additional Wolbachia gene analysis and to provide the quantitative PCR data. This would provide more information on the Wolbachia strain infections being presented and allow these strains to be put into context with recent discoveries in other Anopheles species.

Introduction

The introduction needs significant improvement in the referencing. For example, the sentence ‘In addition, Wolbachia can interfere with the development of some pathogens, including dengue virus’ contains a reference to a publication that only describes Wolbachia establishment and invasion in an Aedes aegypti laboratory population (not virus inhibition). 

Furthermore, including references 5 & 6 in the context of pathogen blocking is not appropriate given this was work which was proposing to use cytoplasmic incompatibility to reduce Culex mosquito populations and artificial Wolbachia-infected mosquito lines were only established in the mid 2000s. 

The paragraph describing natural Wolbachia infections in mosquitoes also needs further references 1 , 2 , 3.

The final paragraph in the introduction presents the fact that Natural Wolbachia infections in Southeast Asian malaria vectors have not been reported. However, the authors should reference the studies in which screening of Anopheles species for Wolbachia was undertaken despite finding no evidence of natural infections 4.

Methods

The primers used for Wolbachia detection W-Specf (CATACC TATTCGAAGGGATAG) and W-Specr (AGCTTCGAGTGAA ACCAATTC) produce a product size of 438 bp and this (to my knowledge) would not be possible or has not been reported using a qrtPCR format. The authors also reference Gomes et al. 5 which used a different reverse primer (5′-TTGCGGGACTTAACCCAACA-3′) that results in amplification of a smaller fragment of the 16S rRNA gene for qrtPCR. The accession numbers MK336794 - MK336806 refer to sequences with >400 bases indicating W-Specf/W-Specr was used. The authors need to clarify if W-Specf/W-Specr was used on a qrtPCR format or if both were used independently and report the differences in prevalence rates using these two primer sets.  

Results

The inclusion for analysis of only what would be considered ‘primary malaria vectors’ needs more explanation if the authors overall aim was to provide evidence for natural Wolbachia infections in Anopheles species given the mosquitoes were not screened for Plasmodium infection.  

Tables 1-3 provide a breakdown of the species composition collected at the different villages but I think it would be more informative to have all the different species grouped according to villages.  Currently it’s difficult to determine mosquito species prevalence on a village level.

The statistics used in tables 1-3 don’t appear to be explained either in the manuscript methods or in the table legends. For example, I am assuming ‘n/N’ means the species/total number collected but again this would be much easier to understand if species were grouped by villages.

Wolbachia infections

With reference to my previous point raised in the methods, which 16S PCR primer set and format was used to determine the prevalence rates and to generate sequences for Figure 2? 

Figure 2 needs to have more details included such as the number of nucleotide sequences used in the analysis and the total positions in the final dataset.

The authors provide the overall prevalence rate (13/370) and then have Figure 3 to show the individual species. Figure 3a I don’t feel is needed because plotting 1/11 (PSE) and 1/12 (DIR) seems unnecessary and could be in a table that incorporates prevalence rates by species and village.  Having an overall village prevalence rate (Fig 3b) has little biological relevance given you have variable Anopheles species containing what appears to be different Wolbachia strains based on 16S analysis.   

A major limitation of the phylogenetic analysis (and even the prevalence rates) is only using a single Wolbachia gene (16S) but I appreciate that Cp values ranging from 31-40.6 are at the limit of detection. The authors should provide these 16S Cp values to allow the reader to see the variation both between technical and biological replicates.  

Could the authors also provide the rationale for concluding that samples were positive where not all technical replicates produced positive amplification given ‘only one sample gave 3/3 positive reactions’? How do these results fit with the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines 6?

Could the authors not have used another Wolbachia qPCR assay based on a second gene that targets a broad range of strains 7?

The density comparison to laboratory-reared Aedes aegypti artificially-infected with the wMel Wolbachia strain is not particularly informative for several reasons. Firstly, quantifying Wolbachia density without accounting for mosquito body size and/or DNA extraction efficiency is problematic. Secondly, the wMel is a strain that naturally infects Drosophila melanogaster so a better comparison would be to natural infections in mosquito species (such as wPip in Culex quinquefasciatus or even the wAlbA/wAlbB strains in Aedes albopictus ). Therefore, I would question the inclusion of this density data given wMel in Ae. aegypti is an artificial infection.    

Discussion

The sentence ‘It was not possible to study more in detail the phylogeny of Wolbachia strain detected in this study by multi locus sequence typing because of the lack of DNA extracts after the screening’ needs clarification. Do the authors mean that they were unable to amplify any of the Wolbachia MLST genes? Did they try using degenerate primer protocols or nested PCR given the qPCR data would indicate low density infections?

The statement “This is consistent with previous attempts to quantify Wolbachia in naturally infected malaria vectors" is incorrect and refers to some (An. gambiae complex) but not all species analysed in Sub-Saharan Africa 8. The authors should expand this discussion as the low density infections presented in this study are comparable to those strains detected in An. gambiae mosquitoes from Sub-Saharan Africa. Some of these studies have only resulted in 16S gene amplification and sequencing resulting in conflicting phylogenetics which appear incompatible with the traditional criteria for vertically transmitted endosymbionts (reviewed in reference 9 9). The authors should provide some further discussion points on whether their results only amplifying 16S could have resulted from either 1) integration into the mosquito genome or 2) some form of contamination. However, additional Wolbachia gene analysis would allow more confidence in these detected strains given the high 16S qPCR Cp values are at the limit of detection.

I confirm that I have read this submission and believe that I have an appropriate level of expertise to confirm that it is of an acceptable scientific standard, however I have significant reservations, as outlined above.

References

  • 1. : Diverse novel resident Wolbachia strains in Culicine mosquitoes from Madagascar. Sci Rep.2018;8(1) : 10.1038/s41598-018-35658-z 17456 10.1038/s41598-018-35658-z [DOI] [PMC free article] [PubMed] [Google Scholar]
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  • 3. : wFlu: characterization and evaluation of a native Wolbachia from the mosquito Aedes fluviatilis as a potential vector control agent. PLoS One.2013;8(3) : 10.1371/journal.pone.0059619 e59619 10.1371/journal.pone.0059619 [DOI] [PMC free article] [PubMed] [Google Scholar]
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  • 5. : Effect of naturally occurringWolbachia inAnopheles gambiae s.l. mosquitoes from Mali onPlasmodium falciparum malaria transmission. Proceedings of the National Academy of Sciences.2017;114(47) : 10.1073/pnas.1716181114 12566-12571 10.1073/pnas.1716181114 [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 6. : The MIQE guidelines: minimum information for publication of quantitative real-time PCR experiments. Clin Chem.2009;55(4) : 10.1373/clinchem.2008.112797 611-22 10.1373/clinchem.2008.112797 [DOI] [PubMed] [Google Scholar]
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  • 8. : Novel Wolbachia strains in Anopheles malaria vectors from Sub-Saharan Africa. Wellcome Open Res.2018;3: 10.12688/wellcomeopenres.14765.2 113 10.12688/wellcomeopenres.14765.2 [DOI] [PMC free article] [PubMed] [Google Scholar]
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Wellcome Open Res. 2019 Jul 3.
Victor Chaumeau 1

We thank to the reviewer for his useful feedback on the manuscript. More details on the qPCR assay used to detect Wolbachia in this study were provided in the revised version of the manuscript, including the raw quantitative data as per reviewer’s suggestion. We also agree that additional Wolbachia genotyping would have been an important added value to the manuscript. However, there was not enough DNA material to attempt additional genotyping of the Wolbachia strains detected in this study given that mosquito samples were also screened for Plasmodium infection and identified at the species level with molecular assays.

Response to point specific comments are listed below:

  • The introduction needs significant improvement in the referencing. For example, the sentence ‘In addition, Wolbachia can interfere with the development of some pathogens, including dengue virus’ contains a reference to a publication that only describes Wolbachia establishment and invasion in an Aedes aegypti laboratory population (not virus inhibition). Furthermore, including references 5 & 6 in the context of pathogen blocking is not appropriate given this was work which was proposing to use cytoplasmic incompatibility to reduce Culex mosquito populations and artificial Wolbachia-infected mosquito lines were only established in the mid 2000s. The paragraph describing natural Wolbachia infections in mosquitoes also needs further references. The final paragraph in the introduction presents the fact that Natural Wolbachia infections in Southeast Asian malaria vectors have not been reported. However, the authors should reference the studies in which screening of Anopheles species for Wolbachia was undertaken despite finding no evidence of natural infections.

The referencing of the introduction was improved as per reviewer’s suggestions.

  • The primers used for Wolbachia detection W-Specf (CATACC TATTCGAAGGGATAG) and W-Specr (AGCTTCGAGTGAA ACCAATTC) produce a product size of 438 bp and this (to my knowledge) would not be possible or has not been reported using a qrtPCR format. The authors also reference Gomes et al. 5 which used a different reverse primer (5′-TTGCGGGACTTAACCCAACA-3′) that results in amplification of a smaller fragment of the 16S rRNA gene for qrtPCR. The accession numbers MK336794 - MK336806 refer to sequences with >400 bases indicating W-Specf/W-Specr was used. The authors need to clarify if W-Specf/W-Specr was used on a qrtPCR format or if both were used independently and report the differences in prevalence rates using these two primer sets.

Amplification of fragments much longer than 438 bp with real-time PCR technology has been reported previously (Rothfuss, Gasser et al. 2010). Without a priori knowledge on the DNA sequences of the Wolbachia strains detected in this study, the primer W-Specf and W-Specr were chosen for their ability to detect most Wolbachia strains infecting insects and to establish phylogenic relationship among isolates (Werren and Windsor 2000). The results of additional assay optimization and serial dilution experiments with the W-Specf/W16S primers used by Gomes et al. were added to the revised version of the manuscript, although we did not use these primers for the screening of Wolbachia in field mosquito samples.

  • The inclusion for analysis of only what would be considered ‘primary malaria vectors’ needs more explanation if the authors overall aim was to provide evidence for natural Wolbachia infections in Anopheles species given the mosquitoes were not screened for Plasmodium infection.

The vector status of Anopheles species in the Thailand-Myanmar border area has been determined previously (Somboon, Aramrattana et al. 1998, Chaumeau, Fustec et al. 2018). Primary vectors in the study area are An. minimus s.s. (Minimus Complex, Funestus Group), An. maculatus s.s., An. sawadwongporni (Maculatus Group), An. dirus s.s. and An. baimaii (Dirus Complex, Leucosphyrus Group). Proper referencing was added in the revised version of the manuscript.

  • Tables 1-3 provide a breakdown of the species composition collected at the different villages but I think it would be more informative to have all the different species grouped according to villages.  Currently it’s difficult to determine mosquito species prevalence on a village level.

The reviewer is right to question the relevance of presenting specific diversity as a proportion of a given species in the corresponding group. In the revised manuscript, human-biting rates were reported is a single table instead of the relative proportions.

  • The statistics used in tables 1-3 don’t appear to be explained either in the manuscript methods or in the table legends. For example, I am assuming ‘n/N’ means the species/total number collected but again this would be much easier to understand if species were grouped by villages.

In the revised version of the manuscript, table 1-3 were merged in a single table and appropriate description of the statistics used in this table were added to the Methods section.

  • With reference to my previous point raised in the methods, which 16S PCR primer set and format was used to determine the prevalence rates and to generate sequences for Figure 2?

The primer W-Specf and W-Specr were used in a real-team PCR format for both estimation of the prevalence rates and phylogenetic analysis as described in the Methods section.

  • Figure 2 needs to have more details included such as the number of nucleotide sequences used in the analysis and the total positions in the final dataset.

More details were added in the revised phylogenetic tree and the total number of position in the final dataset was stated in the figure legend.

  • The authors provide the overall prevalence rate (13/370) and then have Figure 3 to show the individual species. Figure 3a I don’t feel is needed because plotting 1/11 (PSE) and 1/12 (DIR) seems unnecessary and could be in a table that incorporates prevalence rates by species and village.  Having an overall village prevalence rate (Fig 3b) has little biological relevance given you have variable Anopheles species containing what appears to be different Wolbachia strains based on 16S analysis.

The reviewer is right to question the biological significance of plotting prevalence estimates per species and per village. In the revised manuscript, the screening results collated by village and species are presented in a table, and the two plots were removed.

  • A major limitation of the phylogenetic analysis (and even the prevalence rates) is only using a single Wolbachia gene (16S) but I appreciate that Cp values ranging from 31-40.6 are at the limit of detection. The authors should provide these 16S Cp values to allow the reader to see the variation both between technical and biological replicates.

We agree with the reviewer the analyzing only 16S ssuRNA genes is a limitation of our study. Raw quantitative data were added to the revised version of the manuscript as per reviewer suggestion.

  • Could the authors also provide the rationale for concluding that samples were positive where not all technical replicates produced positive amplification given ‘only one sample gave 3/3 positive reactions’? How do these results fit with the Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines ?

It is common that at low parasite concentration, only some replicates give a positive result because the distribution of the DNA template in the reaction tube follow a Poisson distribution (Sterkers, Varlet-Marie et al. 2010, Stahlberg and Kubista 2014, Chaumeau, Andolina et al. 2016). This observation does not challenge the validity of our results. A clear statement that some Wolbachia infected sample have probably been missed because bacterial density observed in Anopheles are close or below the limit of detection of the assay that give 95% positive reaction. Misleading interpretations on the prevalence of Wolbachia infection were removed given the small sample size and the possibility of false negative.

  • Could the authors not have used another Wolbachia qPCR assay based on a second gene that targets a broad range of strains.

In the absence of a priori knowledge on the Wolbachia strains infecting Anopheles  samples, the W-Specf/ W-Specr primers were chosen for their ability to detect a broad range of strains infecting insects (Werren and Windsor 2000). There was not enough DNA material remaining after the screening with the W-Specf/ W-Specr primers to use another assay. However, the specificity of the PCR was confirmed in all positive sample by Sanger sequencing of the PCR product.

  • The density comparison to laboratory-reared Aedes aegypti artificially-infected with the wMel Wolbachia strain is not particularly informative for several reasons. Firstly, quantifying Wolbachia density without accounting for mosquito body size and/or DNA extraction efficiency is problematic. Secondly, the wMel is a strain that naturally infects Drosophila melanogaster so a better comparison would be to natural infections in mosquito species (such as wPip in Culex quinquefasciatus or even the wAlbA/wAlbB strains in Aedes albopictus ). Therefore, I would question the inclusion of this density data given wMel in Ae. aegypti is an artificial infection.

We agree with the reviewer that there is little biological relevance in comparing the density of Wolbachia infection in artificially infected Aedes aegypti and naturally infected Anopheles. We think that presenting those quantitative data is an added value to support that natural Wolbachia infection in this study actually occur at low density rather than resulting from of a low assay sensitivity. We believe that a calibrator to normalize the signal is not necessary as DNA was extracted from whole mosquitoes and bacterial loads expressed as an (arbitrary) number of bacteria per mosquito rather the a number of bacteria per weight-unit of mosquito body (Varlet-Marie, Sterkers et al. 2014, Chaumeau, Andolina et al. 2016).

  • The sentence ‘It was not possible to study more in detail the phylogeny of Wolbachia strain detected in this study by multi locus sequence typing because of the lack of DNA extracts after the screening’ needs clarification. Do the authors mean that they were unable to amplify any of the Wolbachia MLST genes? Did they try using degenerate primer protocols or nested PCR given the qPCR data would indicate low density infections?

The methodology used for Wolbachia detection in this study was described into detail in the Methods section and we did not attempt additional experiments on Wolbachia than that described in the manuscript. In addition to Wolbachia detection and molecular identification of the mosquito species, samples were also screened for Plasmodium infection (data not shown). There was no DNA material to perform additional experiment after the screening with the W-Specf/ W-Specr primers.

  • The statement “This is consistent with previous attempts to quantify Wolbachia in naturally infected malaria vectors" is incorrect and refers to some (An. gambiae complex) but not all species analysed in Sub-Saharan Africa 8 . The authors should expand this discussion as the low density infections presented in this study are comparable to those strains detected in An. gambiae mosquitoes from Sub-Saharan Africa. Some of these studies have only resulted in 16S gene amplification and sequencing resulting in conflicting phylogenetics which appear incompatible with the traditional criteria for vertically transmitted endosymbionts (reviewed in reference 9 9 ). The authors should provide some further discussion points on whether their results only amplifying 16S could have resulted from either 1) integration into the mosquito genome or 2) some form of contamination. However, additional Wolbachia gene analysis would allow more confidence in these detected strains given the high 16S qPCR Cp values are at the limit of detection.

We thank to the reviewer for his useful feedback on the interpretation of our quantitative data. Suggested edits and references were added in the revised version of the manuscript.

References

Chaumeau, V., C. Andolina, B. Fustec, N. Tuikue Ndam, C. Brengues, S. Herder, D. Cerqueira, T. Chareonviriyaphap, F. Nosten and V. Corbel (2016). "Comparison of the Performances of Five Primer Sets for the Detection and Quantification of Plasmodium in Anopheline Vectors by Real-Time PCR." PLoS One 11(7): e0159160.

Chaumeau, V., B. Fustec, S. Nay Hsel, C. Montazeau, S. Naw Nyo, S. Metaane, S. Sawasdichai, P. Kittiphanakun, P. Phatharakokordbun, N. Kwansomboon, C. Andolina, D. Cerqueira, T. Chareonviriyaphap, F. H. Nosten and V. Corbel (2018). "Entomological determinants of malaria transmission in Kayin state, Eastern Myanmar: A 24-month longitudinal study in four villages." Wellcome Open Research 3(109).

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Stahlberg, A. and M. Kubista (2014). "The workflow of single-cell expression profiling using quantitative real-time PCR." Expert Rev Mol Diagn 14(3): 323-331.

Sterkers, Y., E. Varlet-Marie, S. Cassaing, M. P. Brenier-Pinchart, S. Brun, F. Dalle, L. Delhaes, D. Filisetti, H. Pelloux, H. Yera and P. Bastien (2010). "Multicentric comparative analytical performance study for molecular detection of low amounts of Toxoplasma gondii from simulated specimens." J Clin Microbiol 48(9): 3216-3222.

Varlet-Marie, E., Y. Sterkers, M. P. Brenier-Pinchart, S. Cassaing, F. Dalle, L. Delhaes, D. Filisetti, H. Pelloux, F. Touafek, H. Yera and P. Bastien (2014). "Characterization and multicentric validation of a common standard for Toxoplasma gondii detection using nucleic acid amplification assays." J Clin Microbiol 52(11): 3952-3959.

Werren, J. H. and D. M. Windsor (2000). "Wolbachia infection frequencies in insects: evidence of a global equilibrium?" Proc Biol Sci 267(1450): 1277-1285.

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