Contradictory Results

Detection of Wolbachia in Field-Collected Mosquito Vector, Aedes aegypti

Thaddeus M. Carvajal, Kazuki Hashimoto, Reza Kurniawan Harnandika, Divina M Amalin, Kozo Watanabe
doi: https://doi.org/10.1101/408856

ABSTRACT

It was the impression from past literature that Wolbachia is not naturally found in Ae. aegypti. However, there are have been reports that recently reveals the presence of this endosymbiont in this mosquito vector. With this, our study presents additional support of Wolbachia infection in Ae. aegypti by screening field-collected adult mosquitoes using Wolbachiaspecific 16S rDNA and its surface protein (wsp) makers under optimized PCR conditions. From a total of 672 Ae. aegpyti adult mosquito samples collected in Metropolitan Manila, Philippines, 113 (16.8%) and 89 (13.2%) individual mosquito samples were determined to be Wolbachia infected using the wsp and 16S rDNA markers, respectively. The Ae. aegpyti wsp sample sequences were similar or identical to five known Wolbachia strains belonging to supergroups A or B while majority of 16S rDNA sample sequences were similar to strains belonging to supergroup B. Overall, 80 (11.90%) individual mosquito samples revealed to show positive amplifications in both markers and 69.0% showed congruence in supergroup identification (supergroup B). Our findings illustrate that the infection status of Wolbachia in Ae. aegypti may appear common than previously recognized.

INTRODUCTION

Mosquitoes are considered to be medically important insects because of their capacity to carry notable human disease pathogens1. Among the known mosquito vectors, Aedes aegypti is an efficient and dangerous mosquito vector because of its ability to carry significant arboviral diseases such as Dengue, Chikungunya, Yellow Fever and Zika2,3. Despite the development of vaccines, these arboviral diseases are considered to be the leading cause of global disease burden4and thus, targeting the mosquito vector is deemed to be the primary control and prevention. A considerable number of vector control strategies had been implemented, but the disease burden continues to increase. Novel and newer approaches are being developed that shows promising outcomes in vector and disease control and one of which is the utilization of the intracellular bacterial endosymbiont, Wolbachia7-9.

Wolbachia is a naturally occurring endosymbiont which can be maternally inherited and cause different reproductive alterations in its host to increase their transmission to the next generation10-12. In insects, it is estimated to be naturally present in 60-65% of known species13. As to date, there are 17 identified major clades or supergroups (A-Q) where a majority are known to infect arthropods such as insects, arachnids, and crustaceans14. The pathogenic effects of Wolbachia in its host are well-studied and determined to cause spermegg incompatibility, parthenogenesis, cytoplasmic incompatibility, and feminization11,15. Therefore, utilizing these effects towards medically-important mosquito vectors, such as Ae. aegypti, has taken great research strides in the past two decades. The discovery of a virulent Wolbachia strain (wMelPop) in Drosophila melanogaster was successfully transferred to Ae. aegypti where it reduced the lifespan of the mosquito vector16-18. In addition to this, wMelPop and other Wolbachia strains (e.g., wMel) were able to demonstrate conferring resistance on a wide range of insect viruses, especially to human viral pathogens, such as dengue and chikungunya19-22. The life-shortening capability plus pathogen interference of this Wolbachia strain opened an avenue for its potential use as a biological control agent approach against mosquito-borne diseases. The World Mosquito Program (https://www.worldmosquitoprogram.org), formerly known as the Eliminate Dengue Project, was able to generate stable Wolbachia-infected Ae. aegypti lines that possessed the ability of pathogen interference from dengue viruses under laboratory conditions. These Wolbachia strains showed maternal transmission rates close to 100% and induced high levels of cytoplasmic incompatibility to Ae. aegypti16. Semi-field cage experiments were also conducted to assess the fitness cost effect of the discovered strain towards the mosquito vector and its ability of these strains to invade the mosquito population. These experiments demonstrated the true potential of the endosymbiont because of the reduced fecundity of Wolbachia-infected Ae. aegypti as compared to the uninfected wildtype22. Australia became the first country to release these Wolbachia-infected Ae. aegypti into the wild population where it exhibited promising results23,24. As to date, this methodological strategy against the mosquito vector, Ae. aegypti, is now being tested in eight dengue-endemic countries such as Indonesia, Vietnam, Colombia, and Brazil (https://www.worldmosquitoprogram.org). It claimed this approach is considered to be cost-effective and safer for the environment than conventional insecticide-based measures19,25.

With the recognition of about 65% of known insects to be naturally infected with Wolbachia including those mosquito species from the genera of Aedes, Culex, Mansonia, major mosquito vectors of diseases such as Ae. aegypti and Anopheline mosquitoes were reported not to possess this endosymbiont26-31. It led to the belief that the presence of Wolbachia endosymbiont could be the reason why many of the mosquito species are considered to be weak vectors23. Nonetheless, more recent studies show evidence that Wolbachia infection in Ae. aegypti and Anopheles gambiae may appear to be more common than it was previously recognized. Natural Wolbachia infections have now been reported in adult, larvae and egg populations of An. gambiae32-34.

Lately, studies have reported detecting Wolbachia from field-collected Ae. aegypti samples using either wsp marker35or 16S metabarcoding36-37. Though these studies are commendable, there were still uncertainties in establishing whether the mosquito vector does harbor naturally the endosymbiont. Although metabarcoding studies had a substantial sample size (n=85-270), there were unable to report an accurate estimate of the infection rate because mosquito adult or larval samples were pooled from each location. In contrast, wsp detection in Ae. aegypti larval samples35were screened individually, thus, was able to report the infection rate (50.0%). However, it was difficult to affirm or ascertain its true prevalence since the sample size was small (n=16 individuals). Moreover, there is possibility of a potential bias in reporting a high infection rate if larval samples were collected from the same water container due to the sampling of mosquito siblings from the same female mosquito. Nevertheless, these studies further suggest the likelihood of Wolbachia to be naturally associated with Ae. aegypti, thus, opening an avenue to re-visit or re-examine its infection status.

Our study aims to present additional support of Wolbachia infection found from fieldcollected Ae. aegypti adult mosquitoes using Wolbachia-specific 16S rDNA and the Wolbachia surface protein (wsp) markers. Based on the limitations presented from previous studies, two considerations were applied in addressing these gaps. First, Wolbachia screening was done over a large sample size (n=672) and used an individual-based detection of adult Ae. aegypti mosquitoes to gain a better estimate of its prevalence in this mosquito vector. Secondly, two molecular markers were used to confirm the detection status and infer the type of Wolbachia strains found in Ae aegypti.

METHODS

Study area and Mosquito collection

The study area is the National Capital Region of the Philippines or also known as Metropolitan Manila. Located on the Eastern shore of Manila Bay in Southwestern Luzon Island (14°50′ N Latitude, 121°E Longitude), it is considered to be one of the highly urbanized and densely populated areas in the Philippines. Dengue disease is endemic in this region where it accounts for 15%-25% of the total number of reported Dengue cases annually in 2009 201438. Vector control programs are being implemented in various localities of the region. Insecticide application and cleaning of the surroundings have been extensively used however its effectiveness is in question because of the constant and unchanging burden of the disease. As to date, the Philippines, especially Metropolitan Manila, has never conducted any Wolbachia-based program against Ae. aegypti.

Adult mosquito samples were collected using a commercial branded mosquito UVlight trap (Jocanima®) installed in the outdoor premises of 138 residential households (sampling sites) from May 2014 – January 2015 (Figure 1a). Collected samples were then sorted and identified as Ae. aegypti using available keys39. Each sample was then placed in a tube with 99.5% ethanol for preservation. A total of 672 Ae. aegypti adult mosquito samples were collected, identified, labeled accordingly (See Supplementary Table 1) and stored at 20°C for subsequent processing.

Fig 1.
Fig 1.

(a) Spatial distribution of the sampling sites (n=138) for collecting adult Ae. aegypti. Wolbachia-positive sampling sites based on wsp (b) and 16S rDNA (c). Details of the number of Wolbachia-positive mosquitoes per sampling site is found in the Table S1.

DNA Extraction, Polymerase Chain Reaction, and Sequencing

Total genomic DNA of each mosquito individual was extracted using the QIAGEN Blood and Tissue DNEasy Kit© following a modified protocol40. Our study used two molecular markers for detecting Wolbachia infection namely; wsp41 and 16S rDNA42. The primer sequences are as follows: wsp 81F (5′TGG TCC AAT AAG TGA TGA AGA AAC) and wsp 691R (5′ AAA AAT TAA ACG CTA CTC CA) for wsp marker while Wspecf (AGC TTC GAG TGA AAC CAA TTC) and Wspecr (GAA GAT AAT GAC GGT ACT CAC) for 16S rDNA.

For wsp gene amplification, we followed the standard wsp protocol30where the suggested annealing temperature and number of cycles were 55 °C and 30 cycles respectively. In order to conduct an individual-based detection, we initially performed this protocol in Culex quinquefasciatus as our positive control. Certain modifications were made in the standard protocol based on the results where the annealing temperature was set to 57 °C and the number of cycles increased to 35 cycles. This initial modified protocol was performed in individual Ae. aegypti samples where it yielded positive faint bands. It prompted us to remodify again the protocol where the annealing temperature is set at 59 °C with 40 cycles and the addition of 10% DMSO (Sigma-Aldrich®) that led to desirable results necessary for sequencing. Therefore, a 10 μl final reaction volume was used and composed of 10X buffer (TAKARA®), 25 mM MgCl2, 10 mM of each dNTPs, 10 μ M forward and reverse primers, 10% DMSO (Sigma-Aldrich®) and 5.0U/ μl of Taq DNA polymerase (TAKARA®). The final thermal profile consisted an initial denaturation of 95 °C for 3 minutes, followed by another denaturation temperature of 95 °C for 1 minute, an annealing temperature of 59 °C for 1 minute and an extension temperature of 72 °C for 1 minute for 40 cycles, and accompanied by a final extension temperature at 72 °C for 3 minutes.

On the other hand, 16S rDNA gene amplification used a 10 μl final reaction volume and composed of 10X buffer (TAKARA), 25 mM MgCl2, 10 mM of each dNTPs, 10 μM forward and reverse primers, 10% DMSO (Sigma-Aldrich®) and 5.0U/ μL of Taq DNA polymerase (TAKARA®). Thermal profiles follow the protocol of Simões et al42with initial denaturation temperature at 95 °C for 2 minutes, followed by two cycles of 95 °C for 2 minutes of denaturation, annealing temperature of 60 °C for 1 minutes and extension temperature of 72 °C for 1 minute, afterwards 35 cycles of denaturation of 95 °C for 30 seconds, annealing temperature of 60 °C for 1 minute and extension temperature of 72 °C for45 seconds and final extension at 72 °C for 10 minutes.

All PCR amplification experiments included positive and negative controls. The positive control is a Wolbachia-infected Cu. quinquefasciatus sample while the negative control consisted of water as the template. The product size of each molecular marker was checked through electrophoresis with 1.5% agarose gel set at 100 volts for 30 minutes. The size of the amplified wsp gene is 610 bp while the 16S rDNA gene is 438 bp. PCR amplification process underwent two replicates to validate the results obtained (See Supplementary Table 1). A third screening was performed for selected individual samples that had conflicting results based on the two prior replicates. Therefore, the criteria set in reporting the certainty for Wolbachia infection is based on two successful amplification of the molecular markers. Amplified PCR products from each molecular marker were sent for sequencing to Eurofins, Operon – Tokyo.

Identity of Wolbachia strains and their positions in phylogroups

All sequences were subjected to the Nucleotide Basic Local Alignment Search Tool (BLAST) and compared to deposited Wolbachia sequences in GENBANK. Next, selected sequences of Wolbachia strains (Table 1) and those obtained in the study underwent multiple alignment using Clustal W in MEGA 643. After editing, the final length used for phylogenetic inference analyses was 398 bp and 732 bp for wsp and 16S rDNA respectively. The identities and relationships of the Wolbachia strains obtained in our study were determined by performing the Bayesian method in PhyML 3.0 software with 1000 bootstrap replicates44. The Smart Model Selection45 was also utilized to set the parameters for wsp as GTR+G (number of estimated parameters k = 232, Akaike Information Criterion (AIC) = 4897.31702) and 16S rDNA as GTR+G+1 (number of estimated parameters k = 207, Akaike Information Criterion (AIC) = 5332.88688). All sample sequences were submitted to GENBANK with Accession numbers____ - _____.

View this table:
Table 1.

Representative Wolbachia type sequences from different insect hosts in wsp and 16S rDNA molecular markers.

Statistical Analysis

A Clark-Evans test was performed to determine if the spatial distribution of Wolbachia-positive mosquito samples from each molecular marker have a pattern of complete spatial randomness. The test uses the aggregation index (R) where a value of > 1 suggests an ordered distribution and a value of < 1 suggests clustering. This analysis was performed using R program version 3.3.546under package spatstat 46

RESULTS

Detection of Wolbachia through wsp and its phylogeny

From a total of 672 adult Ae. aegypti screened, 113 (16.8%) individual adult mosquito samples are infected with Wolbachia using the wsp marker (Table 2). Based on the study’s criterion (See methods), only 17 samples demonstrated one successful amplification, thus excluding them for further analysis. In addition, female/male ratio is 0.82 (Table 2). All sequenced amplicons resulted in a high degree of similarity (>98.0%) with deposited wsp sequences in GENBANK. The spatial distribution showed that 60 (43.0%) sampling sites (Figure 1b) contained Wolbachia positive mosquitoes with 1 – 8 individuals. Further analysis showed that the distribution of wsp-positive mosquito samples was significantly clustered (R = 0.003,p < 0.001). Figure 2 and Figure S1 show the phylogeny of Wolbachia sequences based on wsp sequences. Majority of the sequences were found in supergroup B (n=84) while the remaining were clustered in supergroup A (n=29). Based on descending order of sample sizes, sample sequences in supergroup B were identical (>99.0%) to Wolbachia type strains from selected hosts of Ae. albopictus (wAlbB) (n= 51), Cu. quinquefasciatus, Cu. pipiens (wPip), Ae. aegypti wMel strain (n= 23) and Ephestia cautella (wCau) (n= 10). The sample sequences from supergroup A were either similar (98.0-99.0%) (n = 8) or identical (>99.0%) (n= 21) to the type strain (wAlbA) from host Ae. albopictus.

View this table:
Table 2.

Summary of wsp and 16S rDNA detection results in Ae. aegypti

Fig 2.
Fig 2.

Phylogenic analysis of wsp. The alignment was analyzed in PhyML. Sample sequences of Ae.aegypti collected in Metropolitan Manila are in red, labeled as AAML (Ae. aegypti Metropolitan ManiLa) and alphanumeric values indicate the unique code assigned to each Ae. aegypti individual sample. Merging (triangles) of sample and representative Wolbachia sequences was done to show degree of similarity (98-100%). Supergroups were indicated as A – C depending on the representative sequences used. The phylogenetic trees are re-drawn for better visualization, thus an expanded version can be viewed in Supplemental Figure S1. Please refer to Table 1 for the Wolbachia type sequences (ingroup and outgroup) for both markers.

Detection of Wolbachia through 16S rDNA and its phylogeny

For 16S rDNA, 89 (13.2%) individual adult mosquito samples were infected with Wolbachia (Table 2). 20 individual mosquito samples generated one successful 16S rDNA amplification, thus, excluding them for further analysis. Furthermore, female/male ratio is 0.85 (Table 2). 50 (36.0%) sampling sites (Figure 1c) contained Wolbachia-positive mosquitoes ranging from 1-8 individuals and the distribution of 16SrDNA-positive individuals revealed to be clustered or aggregated (R = 0.001,p < 0.001). All sequenced amplicons resulted in a high degree of similarity (>98%) with deposited 16S rDNA Wolbachia sequences in GENBANK. Nearly all 16S rDNA sample sequences (n=85) (Figure 3, Figure S2) were grouped in supergroup B. Only one sample sequence was identical to Nasonia vitripennis while the remaining sample sequences were up to 99% similar from the selected hosts of the supergroup. The remaining sample sequences (n=4) were grouped in supergroup C & J. One sample sequence was highly similar (>99%) with Dirofilaria immitis while the remaining were 98-99% similar from the selected hosts of the supergroup.

Fig 3.
Fig 3.

Phylogenic analysis of 16S rDNA. The alignment was analyzed in PhyML. Sample sequences of Ae.aegypti collected in Metropolitan Manila are in red, labeled as AAML (Ae. aegypti Metropolitan ManiLa) and alphanumeric values indicate the unique code assigned to each Ae. aegypti individual sample. Merging (triangles) of sample and representative Wolbachia sequences was done to show degree of similarity (98-100%). Supergroups were indicated as A – J depending on the representative sequences used. The phylogenetic trees are re-drawn for better visualization, thus an expanded version can be viewed in Supplemental Figure S2. Please refer to Table 1 for the Wolbachia type sequences (ingroup and outgroup) for both markers.

Comparison of 16S rDNA and wsp for Wolbachia detection and phylogeny

From the 113 and 89 positively detected mosquito individuals from wsp and 16S rDNA respectively, 80 (11.90%) individual samples yielded positive amplification in both markers (Table 2). In wsp positive detection (n=113), 80 had two successful amplification of the 16S rDNA marker while 27 had only one amplification of 16S rDNA and the remaining 6 had no successful amplification on 16S rDNA marker. On the other hand, the 89 individual samples deemed 16S rDNA positive for Wolbachia showed 80 individuals had two successful amplification of the wsp marker while 9 had only one successful amplification on the said marker. Next, we focus on the supergroup classification of the 80 individual samples based on the wsp and 16S rDNA phylogeny. It was shown that 55 (69%) had the same classification in supergroup B while the remaining 25 (31%) showed a disparity in supergroup classification. Such difference, for example, showed that wsp identified the individual sample as supergroup A, but 16S rDNA reveals to be either supergroup B or C & J.

DISCUSSION

Our study was able to demonstrate the detection of the endosymbiont Wolbachia in field-caught adult Ae. aegypti. Notably, the main reason for the positive detection, especially in wsp, is because of the procedural modifications or optimization in the amplification of the said marker. A case in point, for example, why optimization is necessary is the evidence presented in the malaria mosquito vector, An. gambiae. Previous studies had reported no observed natural Wolbachia infection in this mosquito vector26-31; however, the endosymbiont was successfully detected in An. gambiae from Burkina Faso, West Africa using an optimized wsp protocol32,33. Another potential reason for a positive detection was the study’s sample size. Based on several literature on assessing the prevalence of Wolbachia in different mosquito species, the highest number of Ae. aegypti individuals screened was 11930which resulted in non-detection of the endosymbiont. As compared to the actual study (n= 672), the sample sizes from previous studies were low; thus, larger sample size would provide a more accurate estimate of the prevalence of Wolbachia infection. Similarly, these reasons were clearly emphasized by recent studies on why earlier investigations may have underestimated the actual incidence of Wolbachia infection from different insect hosts48,49.

Our study acknowledges the uncertainties associated with conventional PCR detection such as high false positive detection rates. With this in mind, the study was cautious in affirming a positive infection in each Ae. aegypti adult sample. First, the selection of markers is based on the recommendation of Simoes et al.42that two of its preferred primer sets (e.g. Wspecf and Wspecr) was determined to produce the lowest false positive and false negative rates. Secondly, our study performed replications with a stringent criterion for a successful Wolbachia infection on each mosquito sample. Although there are several genetic markers (e.g. MLST genes) and techniques (e.g. IFA, FISH or whole-genome sequencing) available, the primary intention of this study is to detect Wolbachia infection in Ae. aegypti initially using this PCR-Based approach.

Linking our findings with the previous studies35-37which reported Wolbachia in Ae. aegypti may incidentally provide a clear picture of its infection status. First, the probable density of the endosymbiont found in this mosquito vector may be low. Even though our study did not measure the actual density, a 40-cycle PCR amplification procedure or a long PCR run50may detect a small amount of Wolbachia present. It partly supports the results presented from metabarcoding studies36,37where a low number (2-4) of Wolbachia sequence reads were detected in both the larvae and adult Ae. aegypti mosquito. These can be another potential reason why earlier prevalence studies were not able to detect Wolbachia in Ae. aegypti samples. Moreover, the low probable density of the endosymbiont may also translate to the observed low infection rate (13-16%) found in our study. This again partly supports metabarcoding studies36,37where only two Ae. aegypti mosquito pools had the presence of these low number Wolbachia sequences. On the other hand, our results are in contrast with the report from Ae. aegypti larvae (n=16 individuals) in Malaysia which resulted in a 50% infection rate35. However, there could be some uncertainties to this estimate because of its small sample size and, more importantly, the collected larval samples may be siblings from the same female Ae. aegypti mosquito. The limitation as mentioned earlier prompted us to conduct an individual-based adult mosquito detection so that it can present a better and explicit estimation of the infection rate. Secondly, we assume that the Wolbachia strain/s found in Ae. aegypti can be maternally-inherited due to the following reasons: (a) reported positive infections in larval samples from the previous studies35-37and (b) detecting positive infections in male Ae. aegypti mosquitoes (our study, Table 2). However, there is still a need to present direct evidence of maternal transmission of this endosymbiont during thedevelopmental stages of Ae. aegypti since all studies, including ours, were performed independently.

Lastly, the Wolbachia strains infecting Ae. aegypti have been shown in our study belong to supergroups A and B. Both wsp and 16S rDNA phylogeny showed that majority of the individual samples belong to supergroup B while a small number of individual samples were found in supergroup A (based on wsp). Detecting different Wolbachia strains in a single mosquito species is relatively common especially in medically important mosquitoes, Ae. albopictus51,52 and An. gambiae32, and other insect host species (e.g. Drosophila species51). Since our study presented a majority of our sample sequences belonging to supergroup B, this was also the same observation reported by previous studies35-37. Dipterans, especially mosquitoes, are commonly infected by these Wolbachia strains from supergroups, A and B. It has been shown to cause parasitism towards its insect host by producing phenotype effects such as cytoplasmic incompatibility, male killing, and feminization11,53. Nevertheless, whether the identified Wolbachia strains in Ae. aegypti possess these phenotypic effects remains unclear. Also, further studies are needed to ascertain the pathogenic impact of this local endosymbiont to the mosquito vector. More importantly, it is very essential to determine whether these identified Wolbachia strains could render Ae. aegypti a less effective vector by blocking key arboviruses such as dengue. It is also worth mentioning that some individual samples have shown to be similar with Wolbachia strains found in supergroups C and J based on 16S rDNA. These two supergroups are not generally found in dipterans especially in mosquitoes. It is likely that our 16S rDNA amplified the Wolbachia strain residing in the roundworm, Dirofilaria immitis. Ae. aegpyti mosquitoes are also known to carry this parasitic nematode to certain mammals, such as dogs54. This observation was also reported in one of the metabarcoding studies37that showed sequences of Wolbachia from Dirofilaria immitis. However, when these 16S rDNA results were compared to the wsp results in our study, it showed the Wolbachia wsp sample sequence of the same mosquito individuals belong to supergroup B. We can only infer that the inconsistent results observed in our study may stem towards the sensitivity and specificity of the markers used. The wsp gene marker has been likened to antigen protein typing in screening pathogenic bacteria where it can be a perfect diagnostic tool for detecting Wolbachia infection55,56. However, it is unsuitable for phylogenetic analysis or deeper taxonomic relationship because of its extensive recombination and strong diversifying selection11,57,58. 16S rDNA, on the other hand, is known to be a conserved gene highly suited in bacterial identification and phylogeny, but its use in detecting Wolbachia infection has demonstrated varying results depending on the specific 16S rDNA primers42. It was emphasized that “no single protocol” can ultimately ensure the specificity and accuracy of 16S rDNA to detect Wolbachia infection56. Thus, further claiming that 16S rDNA markers in Wolbachia detection may be far from optimal56.

We consider our findings to be crucially important especially if the Philippines would implement or approve two scenarios in the release of: (a) Wolbachia-infected (e.g. wMelPop or wMel) mosquitoes or (b) local Wolbachia strains found by our study in dengue-endemic areas. In the first scenario, a vital consideration is the presence of “bidirectional incompatibility” mechanism between the intended Wolbachia strain (e.g. wMelPop or wMel) to be released and the present local strain found in the mosquito. There are instances that two strains in one host cannot stably coexist with each other because the naturally occurring strain is preventing the intended strain to reach fixation or establishment59-61. It would serve as an impediment to the intentional spread of Wolbachia strain to the mosquito population. It was suggested that to overcome this incompatibility is to remove the existing natural strain inhabiting the mosquito vector or to perform a “superinfection” where the intended Wolbachia strain induces unidirectional incompatibility with the natural strain62. Nevertheless, it very important to re-examine the infection status of Wolbachia in Ae. aegypti mosquitoes in intended areas prior a mass release program. If the second scenario, utilizing the release of local Wolbachia strains, is implemented, there are specific considerations that should be addressed for a successful population replacement. The first and most important consideration is to determine whether these local strains may exhibit the same phenotypic effects and pathogen blocking of wMel strain to Ae. aegypti. Currently, these characteristics are still unknown and therefore crucial if utilized for mass release. Another consideration is endosymbiont’s density in the mosquito vector. Mosquito species naturally infected with Wolbachia are not ideal candidates due to the changing molecular interactions between Wolbachia and the host over time63. The result of this symbiosis is the amount of bacterial density found in the mosquito host where it can influence the intensity of Wolbachia-induced phenotypic or anti-viral effects22,62,64,65. Newer infections (e.g. tansinfections) are shown to produce high bacterial density while natural infections lead to lower bacterial density due to the adaptation of the host to the endosymbiont infection over time. In our study, we infer that the local Wolbachia strains are in low density inside its host, Ae. aegypti. If this is the case, it will result in a reduced physiological and anti-viral impact of the strain to the mosquito vector. However, high Wolbachia density which also possesses strong inhibitory effects against insect viruses had been observed from natural Wolbachia strains with a long-term association from its host66,67. The last consideration is the low infection rate. It raises the question, more importantly to the population replacement approach, if any of the local Wolbachia strains could be sustained for an extended period or possess the ability to infect the mosquito population thoroughly. Studies had suggested that a successful strain used in population replacement or invasion should reach an infection rate of >90% and should remain at this rate over an extended period of time68-70. Thus, utmost consideration in the infection status of Wolbachia and its role in Ae. aegypti is necessary for a Wolbachia-based vector control program to be successful, efficient and, as well as, effective.

AUTHOR CONTRIBUTIONS

T.M.C., D.M.A. and K.W. designed the experiments. T.M.C., K.H., and R.K.H. performed the experiments. T.M.C., K.H., and R.K.H. performed the sequencing while T.M.C., K.W. and D.M.A. accomplished the phylogenetic analysis. T.M.C. wrote the manuscript along with and K.W. All authors reviewed the manuscript and approved on its submission.

COMPETING INTEREST

The authors declare no competing interest

DATA AVAILABILITY

Demographic profiles (location and sex) and detection status from each individual Ae. aegypti adult mosquito used in the study are presented in the Supplementary. Accession numbers of Nucleotide sequences of PCR-amplified fragments of wsp and 16S have been deposited in the GENBANK nucleotide database under accession numbers ______to ______and _____to ______respectively.

SUPPLMENTAL MATERIAL

Table S1. Demographic profile (Sex, Sampling Site Code, Location), Detection status (wsp and 16S rDNA) of all individual adult Aedes aegypti mosquitoes used in the study. Positive Wolbachia infection in mosquito samples presents the supergroup classification and GENBANK accession number.

Figure S1. Complete wsp phylogeny of Wolbachia from Ae. aegypti (n=113). The alignment was analyzed in the program PHYML and Wolbachia host Dirofilaria immitis was selected as an outgroup. All sample sequences are indicated in red dots. The condensed version of this tree is presented as Figure 1.

Figure S2. Complete 16S rDNA phylogeny of Wolbachia from Ae. aegypti (n=85). The alignment was analyzed in the program PHYML and Rickettsia sp. was selected as an outgroup. All sample sequences are indicated in red dots. The condensed version of this tree is presented as Figure 2

ACKNOWLEDGEMENTS

We would like to thank M.J.L.B. Martinez, J.D.R. Capistrano, V.S.P. Tiopianco, B.M.C Orantia, C.R. Estrada, M.G. Cuenca, K.M. Viacrusis and L.F.T. Hernandez for their valuable work in the collection of the mosquitoes. Also we are grateful to the valuable and pertinent comments of the anonymous reviewers. This work is funded by the JSPS Grant-in-Aid for Scientific Research (16H05750, 17H01624, 17K18906), JSPS Bilateral Joint Research Projects, and Leading Academia in Marine and Environmental Pollution Research – Ehime University (Y29-1-8)

REFERENCES

  1. 1.↵
    Juliano, Steven A., and L. Philip Lounibos. Ecology of invasive mosquitoes: effects on resident species and on human health. Ecology letters 8.5: 558–574 (2005).
    OpenUrlCrossRefPubMedWeb of Science
  2. 2.↵
    Powell, Jeffrey R., and Walter J. Tabachnick. History of domestication and spread of Aedes aegypti-A Review. Memorias do Instituto Oswaldo Cruz 108: 11–17 (2013).
    OpenUrlCrossRefPubMedWeb of Science
  3. 3.↵
    Morrison, Amy C., et al. Defining challenges and proposing solutions for control of the virus vector Aedes aegypti. PLoS medicine 5.3: e68 (2008).
    OpenUrl
  4. 4.↵
    Stanaway JD, Shepard DS, Undurraga EA, Halasa YA, Coffeng LE, Brady OJ, Hay SI, Bedi N, Bensenor IM, Castañeda-Orjuela CA, Chuang TW. The global burden of dengue: an analysis from the Global Burden of Disease Study. The Lancet infectious diseases. 2016 Jun 30;16(6):712–23 (2013)
    OpenUrl
  5. 5.
    Achee NL, Gould F, Perkins TA, Reiner Jr RC, Morrison AC, Ritchie SA, Gubler DJ, Teyssou R, Scott TW. A critical assessment of vector control for dengue prevention. PLoS neglected tropical diseases. May 7;9(5):e0003655 (2015).
    OpenUrl
  6. 6.
    Reiner Jr RC, Achee N, Barrera R, Burkot TR, Chadee DD, Devine GJ, Endy T, Gubler D, Hombach J, Kleinschmidt I, Lenhart A. Quantifying the epidemiological impact of vector control on dengue. PLoS neglected tropical diseases. May 26;10(5):e0004588 (2016).
    OpenUrl
  7. 7.
    Kamtchum-Tatuene J, Makepeace BL, Benjamin L, Baylis M, Solomon T. The potential role of Wolbachia in controlling the transmission of emerging human arboviral infections. Current opinion in infectious diseases. Feb;30(1):108 (2017).
    OpenUrlPubMed
  8. 8.
    Murray JV, Jansen CC, De Barro P. Risk associated with the release of Wolbachia- infected Aedes aegypti mosquitoes into the environment in an effort to control dengue. Frontiers in public health.;4 (2016).
  9. 9.
    Bourtzis K, Dobson SL, Xi Z, Rasgon JL, Calvitti M, Moreira LA, Bossin HC, Moretti R, Baton LA, Hughes GL, Mavingui P. Harnessing mosquito–Wolbachia symbiosis for vector and disease control. Acta tropica. Apr 30;132:S150–63 (2014).
    OpenUrlCrossRefPubMedWeb of Science
  10. 10.↵
    Serbus LR, Casper-Lindley C, Landmann F, Sullivan W. The genetics and cell biology of Wolbachia-host interactions. Annual review of genetics. Dec 1;42:683–707 (2008).
  11. 11.↵
    Werren JH, Baldo L, Clark ME. Wolbachia: master manipulators of invertebrate biology. Nature reviews. Microbiology. Oct 1;6(10):741 (2008).
    OpenUrl
  12. 12.↵
    Saridaki A, Bourtzis K. Wolbachia: more than just a bug in insects genitals. Current opinion in microbiology. Feb 28;13(1):67–72 (2010).
    OpenUrlCrossRefPubMedWeb of Science
  13. 13.↵
    Rasgon JL. Using predictive models to optimize Wolbachia-based strategies for vector-borne disease control. Transgenesis and the Management of Vector-Borne Disease.:114–25 (2008).
  14. 14.↵
    Glowska, Eliza, Anna Dragun-Damian, Miroslawa Dabert, and Michael Gerth. “New Wolbachia supergroups detected in quill mites (Acari: Syringophilidae).” Infection, Genetics and Evolution 30: 140–146 (2015).
    OpenUrl
  15. 15.↵
    Ahmed MZ, Breinholt JW, Kawahara AY. Evidence for common horizontal transmission of Wolbachia among butterflies and moths. BMC evolutionary biology. 2016 May 27;16(1):118.
    OpenUrl
  16. 16.↵
    McMeniman CJ, Lane RV, Cass BN, Fong AW, Sidhu M, Wang YF, O’neill SL. atStable introduction of a life-shortening Wolbachia infection into the mosquito Aedes aegypti. Science. 2009 Jan 2;323(5910):141–4.
    OpenUrlAbstract/FREE Full Text
  17. 17.
    Xi Z, Khoo CC, Dobson SL. Wolbachia establishment and invasion in an Aedes aegypti laboratory population. Science. 2005 Oct 14;310(5746):326–8.
    OpenUrlAbstract/FREE Full Text
  18. 18.↵
    Min, Kyung-Tai, and Seymour Benzer. “Wolbachia, normally a symbiont of Drosophila, can be virulent, causing degeneration and early death.” Proceedings of the National Academy of Sciences 94, no. 20: 10792–10796 (1997).
    OpenUrlAbstract/FREE Full Text
  19. 19.↵
    Dutra HL, Rocha MN, Dias FB, Mansur SB, Caragata EP, Moreira LA. Wolbachia blocks currently circulating Zika virus isolates in Brazilian Aedes aegypti mosquitoes. Cell host & microbe. 2016 Jun 8;19(6):771–4.
    OpenUrlCrossRefPubMed
  20. 20.
    Aliota MT, Walker EC, Yepes AU, Velez ID, Christensen BM, Osorio JE. The wMel strain of Wolbachia reduces transmission of chikungunya virus in Aedes aegypti. PLoS neglected tropical diseases. 2016 Apr 28;10(4):e0004677.
    OpenUrl
  21. 21.
    Frentiu FD, Zakir T, Walker T, Popovici J, Pyke AT, van den Hurk A, McGraw EA, O’Neill SL. Limited dengue virus replication in field-collected Aedes aegypti mosquitoes infected with Wolbachia. PLoS neglected tropical diseases. 2014 Feb 20;8(2):e2688.
    OpenUrl
  22. 22.↵
    Walker TJ, Johnson PH, Moreira LA, Iturbe-Ormaetxe I, Frentiu FD, McMeniman CJ, Leong YS, Dong Y, Axford J, Kriesner P, Lloyd AL. The wMel Wolbachia strain blocks dengue and invades caged Aedes aegypti populations. Nature. 2011 Aug 25;476(7361):450–3.
    OpenUrlCrossRefPubMedWeb of Science
  23. 23.↵
    Popovici J, Moreira LA, Poinsignon A, Iturbe-Ormaetxe I, McNaughton D, O’Neill SL. Assessing key safety concerns of a Wolbachia-based strategy to control dengue transmission by Aedes mosquitoes. Memorias do Instituto Oswaldo Cruz. 2010 Dec;105(8):957–64.
    OpenUrlCrossRefPubMedWeb of Science
  24. 24.↵
    De Barro PJ, Murphy B, Jansen CC, Murray J. The proposed release of the yellow fever mosquito, Aedes aegypti containing a naturally occurring strain of Wolbachia pipientis, a question of regulatory responsibility. Journal für Verbraucherschutz und Lebensmittelsicherheit. 2011 May 1;6(1):33–40.
    OpenUrl
  25. 25.↵
    Iturbe-Ormaetxe I, Walker T, Lo’Neill S. Wolbachia and the biological control of mosquito-borne disease. EMBO reports. 2011 Jun 1;12(6):508–18.
    OpenUrlAbstract/FREE Full Text
  26. 26.↵
    Kittayapong P, Baisley KJ, Baimai V, O’Neill SL. Distribution and diversity of Wolbachia infections in Southeast Asian mosquitoes (Diptera: Culicidae). Journal of medical entomology. 2000 May;37(3):340–5.
    OpenUrlCrossRefPubMedWeb of Science
  27. 27.
    Ricci I, Cancrini G, Gabrielli S, D’amelio S, Favia G. Searching for Wolbachia (Rickettsiales: Rickettsiaceae) in mosquitoes (Diptera: Culicidae): large polymerase chain reaction survey and new identifications. Journal of medical entomology. 2002 Jul;39(4):562–7.
    OpenUrlCrossRefPubMed
  28. 28.
    Rasgon JL, Scott TW. Impact of population age structure on Wolbachia transgene driver efficacy: ecologically complex factors and release of genetically modified mosquitoes. Insect biochemistry and molecular biology. 2004 Jul 31;34(7):707–13.
    OpenUrlCrossRefPubMedWeb of Science
  29. 29.
    Hilgenboecker K, Hammerstein P, Schlattmann P, Telschow A, Werren JH. How many species are infected with Wolbachia?–a statistical analysis of current data. FEMS microbiology letters. 2008 Apr 1;281(2):215–20.
    OpenUrlCrossRefPubMedWeb of Science
  30. 30.↵
    Wiwatanaratanabutr I. Geographic distribution of wolbachial infections in mosquitoes from Thailand. Journal of invertebrate pathology. 2013 Nov 30;114(3):337–40.
    OpenUrlCrossRef
  31. 31.↵
    Nugapola NN, De Silva WP, Karunaratne SP. Distribution and phylogeny of Wolbachia strains in wild mosquito populations in Sri Lanka. Parasites & vectors. 2017 May 10;10(1):230.
    OpenUrl
  32. 32.↵
    Baldini F, Segata N, Pompon J, Marcenac P, Shaw WR, Dabiré RK, Diabaté A, Levashina EA, Catteruccia F. Evidence of natural Wolbachia infections in field populations of Anopheles gambiae. Nature communications. 2014 Jun 6;5.
  33. 33.↵
    Shaw WR, Marcenac P, Childs LM, Buckee CO, Baldini F, Sawadogo SP, Dabiré RK, Diabaté A, Catteruccia F. Wolbachia infections in natural Anopheles populations affect egg laying and negatively correlate with Plasmodium development. Nature communications. 2016;7.
  34. 34.↵
    Gomes FM, Hixson BL, Tyner MD, Ramirez JL, Canepa GE, Silva TL, Molina-Cruz A, Keita M, Kane F, Traoré B, Sogoba N. Effect of naturally occurring Wolbachia in Anopheles gambiae sl mosquitoes from Mali on Plasmodium falciparum malaria transmission. Proceedings of the National Academy of Sciences. 2017 Nov 7:201716181.
    OpenUrl
  35. 35.↵
    Teo CH, Lim PK, Voon K, Mak JW. Detection of dengue viruses and Wolbachia in Aedes aegypti and Aedes albopictus larvae from four urban localities in Kuala Lumpur, Malaysia. Tropical Biomedicine. 2017 Sep 1;34(3):583–97.
    OpenUrl
  36. 36.↵
    Coon KL, Brown MR, Strand MR. Mosquitoes host communities of bacteria that are essential for development but vary greatly between local habitats. Molecular ecology. 2016 Nov 1;25(22):5806–26.
    OpenUrlCrossRef
  37. 37.↵
    Thongsripong P, Chandler JA, Green AB, Kittayapong P, Wilcox BA, Kapan DD, Bennett SN. Mosquito vector-associated microbiota: Metabarcoding bacteria and eukaryotic symbionts across habitat types in Thailand endemic for dengue and other arthropod-borne diseases. Ecology and evolution. 2018 Jan 1;8(2):1352–68.
    OpenUrl
  38. 38.↵
    Carvajal TM, Viacrusis KM, Hernandez LF, Ho HT, Amalin DM, Watanabe K. Machine learning methods reveal the temporal pattern of dengue incidence using meteorological factors in metropolitan Manila, Philippines. BMC infectious diseases. 2018 Dec;18(1):183.
    OpenUrl
  39. 39.↵
    Rueda LM. Pictorial keys for the identification of mosquitoes (Diptera: Culicidae) associated with dengue virus transmission. Walter Reed Army Inst Of Research Washington Dc Department Of Entomology; 2004 Aug 3.
  40. 40.↵
    Crane S. DNA Extraction From Archival Museum Insect Specimens. 2011. Accessed Mar 2015. Available from: https://s3-eu-west-1.amazonaws.com/pfigshare-u-files/1114092/extractionmuseum.pdf
  41. 41.↵
    Zhou W, Rousset F, O’Neill S. Phylogeny and PCR–based classification of Wolbachia strains using wsp gene sequences. Proceedings of the Royal Society of London B: Biological Sciences. 1998 Mar 22;265(1395):509–15
    OpenUrlCrossRefPubMedWeb of Science
  42. 42.↵
    Simoes PM, Mialdea G, Reiss D, Sagot MF, Charlat S. Wolbachia detection: an assessment of standard PCR protocols. Molecular Ecology Resources. 2011 May 1;11(3):567–72.
    OpenUrl
  43. 43.↵
    Tamura K, Stecher G, Peterson D, Filipski A, Kumar S. MEGA6: molecular evolutionary genetics analysis version 6.0. Molecular biology and evolution. 2013 Dec 1;30(12):2725–9.
    OpenUrlCrossRefPubMedWeb of Science
  44. 44.↵
    Guindon S, Dufayard J, Lefort V, Anisimova M, Hordijk W, Gascuel O. New algorithms and methods to estimate maximum-likelihood phylogenies: assessing the performance of PhyML 3.0. Syst Biol. 2010;59(3):307–21.
    OpenUrlCrossRefPubMedWeb of Science
  45. 45.↵
    Lefort V, Longueville JE, Gascuel O. SMS: Smart Model Selection in PhyML. Molecular Biology and Evolution. 2017 May 11:msx149.
    OpenUrl
  46. 46.↵
    R Development Core Team. R: A language and environment for statistical computing. R Foundation for Statistical Computing, Vienna, Austria. 2016
  47. 47.
    Baddeley A, Turner R. Spatstat: an R package for analyzing spatial point patterns. Journal of statistical software. 2005 Jan 26;12(6):1–42.
    OpenUrl
  48. 48.↵
    Augustinos AA, Santos-Garcia D, Dionyssopoulou E, Moreira M, Papapanagiotou A, Scarvelakis M, Doudoumis V, Ramos S, Aguiar AF, Borges PA, Khadem M. Detection and characterization of Wolbachia infections in natural populations of aphids: is the hidden diversity fully unraveled?. PLoS one. 2011 Dec 13;6(12):e28695.
    OpenUrlCrossRefPubMed
  49. 49.↵
    Ahmed MZ, Breinholt JW, Kawahara AY. Evidence for common horizontal transmission of Wolbachia among butterflies and moths. BMC evolutionary biology. 2016 May 27;16(1):118.
    OpenUrl
  50. 50.↵
    Jeyaprakash A, Hoy MA. Long PCR improves Wolbachia DNA amplification: wsp sequences found in 76% of sixty-three arthropod species. Insect molecular biology. 2000 Aug 1;9(4):393–405.
    OpenUrlCrossRefPubMedWeb of Science
  51. 51.↵
    Zhou W, Rousset F, O’Neill S. Phylogeny and PCR–based classification of Wolbachia strains using wsp gene sequences. Proceedings of the Royal Society of London B: Biological Sciences. 1998 Mar 22;265(1395):509–15.
    OpenUrlCrossRefPubMedWeb of Science
  52. 52.↵
    Kitrayapong P, Baimai V, O’Neill SL. Field prevalence of Wolbachia in the mosquito vector Aedes albopictus. The American journal of tropical medicine and hygiene. 2002 Jan 1;66(1):108–11.
    OpenUrlAbstract
  53. 53.↵
    Ellegaard KM, Klasson L, Näslund K, Bourtzis K, Andersson SG. Comparative genomics of Wolbachia and the bacterial species concept. PLoS genetics. 2013 Apr 4;9(4):e1003381.
    OpenUrl
  54. 54.↵
    Kramer L, Grandi G, Leoni M, Passeri B, McCall J, Genchi C, Mortarino M, Bazzocchi C. Wolbachia and its influence on the pathology and immunology of Dirofilaria immitis infection. Veterinary parasitology. 2008 Dec 10;158(3):191–5.
    OpenUrlCrossRefPubMed
  55. 55.↵
    Kageyama D, Narita S, Imamura T, Miyanoshita A. Detection and identification of Wolbachia endosymbionts from laboratory stocks of stored-product insect pests and their parasitoids. Journal of stored products research. 2010 Jan 31;46(1):13–9.
    OpenUrlCrossRef
  56. 56.↵
    Perez-Losada M, Viscidi RP, Demma JC, Zenilman J, Crandall KA. Population genetics of Neisseria gonorrhoeae in a high-prevalence community using a hypervariable outer membrane porB and 13 slowly evolving housekeeping genes. Molecular biology and evolution. 2005 Sep 1;22(9):1887–902.
    OpenUrlCrossRefPubMedWeb of Science
  57. 57.↵
    Werren JH, Bartos JD. Recombination in Wolbachia. Current Biology. 2001 Mar 20;11(6):431–5.
    OpenUrlCrossRefPubMedWeb of Science
  58. 58.↵
    Jiggins FM. The rate of recombination in Wolbachia bacteria. Molecular Biology and Evolution. 2002 Sep 1;19(9):1640–3.
    OpenUrlPubMedWeb of Science
  59. 59.↵
    Hoffmann AA, Iturbe-Ormaetxe I, Callahan AG, Phillips BL, Billington K, Axford JK, Montgomery B, Turley AP, O’Neill SL. Stability of the wMel Wolbachia infection following invasion into Aedes aegypti populations. PLoS neglected tropical diseases. 2014 Sep 11;8(9):e3115.
    OpenUrl
  60. 60.
    Hoffmann AA, Montgomery BL, Popovici J, Iturbe-Ormaetxe I, Johnson PH, Muzzi F, et al. Successful establishment of Wolbachia in Aedes populations to suppress dengue transmission. Nature. 2011;476(7361):454–U107.
    OpenUrlCrossRefPubMedWeb of Science
  61. 61.↵
    Hedges LM, Brownlie JC, O’neill SL, Johnson KN. Wolbachia and virus protection in insects. Science. 2008 Oct 31;322(5902):702-.
    OpenUrlAbstract/FREE Full Text
  62. 62.↵
    Joubert DA, Walker T, Carrington LB, De Bruyne JT, Kien DH, Hoang NL, Chau NV, Iturbe-Ormaetxe I, Simmons CP, O’Neill SL. Establishment of a Wolbachia superinfection in Aedes aegypti mosquitoes as a potential approach for future resistance management. PLoS pathogens. 2016 Feb 18;12(2):e1005434.
    OpenUrlCrossRef
  63. 63.↵
    Dale C, Moran NA. Molecular interactions between bacterial symbionts and their hosts. Cell. 2006 Aug 11;126(3):453–65.
    OpenUrlCrossRefPubMedWeb of Science
  64. 64.↵
    Osborne SE, Iturbe-Ormaetxe I, Brownlie JC, O’Neill SL, Johnson KN. Antiviral protection and the importance of Wolbachia density and tissue tropism in Drosophila simulans. Applied and environmental microbiology. 2012 Oct 1;78(19):6922–9.
    OpenUrlAbstract/FREE Full Text
  65. 65.↵
    Martinez J, Longdon B, Bauer S, Chan YS, Miller WJ, Bourtzis K, Teixeira L, Jiggins FM. Symbionts commonly provide broad spectrum resistance to viruses in insects: a comparative analysis of Wolbachia strains. PLoS pathogens. 2014 Sep 18;10(9):e1004369.
    OpenUrlCrossRefPubMed
  66. 66.↵
    Teixeira L, Ferreira Á, Ashburner M. The bacterial symbiont Wolbachia induces resistance to RNA viral infections in Drosophila melanogaster. PLoS biology. 2008 Dec 23;6(12):e1000002.
    OpenUrlCrossRef
  67. 67.↵
    Hedges LM, Brownlie JC, O’neill SL, Johnson KN. Wolbachia and virus protection in insects. Science. 2008 Oct 31;322(5902):702-.
    OpenUrlAbstract/FREE Full Text
  68. 68.↵
    Frentiu FDZ, Walker T, Popovici T, Pyke J, Van den Hurk AT, McGraw A, et al. Limited dengue virus replication in field-collected Aedes aegypti mosquitoes infected with Wolbachia. PLoS Negl Trop Dis. 2014;8:e2688.
    OpenUrlCrossRefPubMed
  69. 69.
    1. Wright R,
    2. Pal R
    Laven H. In: in Genetics of Insect Vectors of Disease. Wright R, Pal R, editors. Amsterdam: Elsevier; 1967. pp. 251–275.
  70. 70.↵
    Blagrove MS, Arias-Goeta C, Failloux AB, Sinkins SP. Wolbachia strain wMel induces cytoplasmic incompatibility and blocks dengue transmission in Aedes albopictus. Proceedings of the National Academy of Sciences. 2012 Jan 3;109(1):255–60.
    OpenUrlAbstract/FREE Full Text
Back to top
Posted September 08, 2018.
Download PDF

Supplementary Material

Email
Detection of Wolbachia in Field-Collected Mosquito Vector, Aedes aegypti
Thaddeus M. Carvajal, Kazuki Hashimoto, Reza Kurniawan Harnandika, Divina M Amalin, Kozo Watanabe
bioRxiv 408856; doi: https://doi.org/10.1101/408856
Twitter logo Facebook logo LinkedIn logo Mendeley logo
Citation Tools

Subject Area