Skip to main content
  • Loading metrics

High DDT resistance without apparent association to kdr and Glutathione-S-transferase (GST) gene mutations in Aedes aegypti population at hotel compounds in Zanzibar

  • Ayubo Kampango ,

    Contributed equally to this work with: Ayubo Kampango, Emma F. Hocke

    Roles Conceptualization, Data curation, Formal analysis, Investigation, Methodology, Software, Supervision, Validation, Visualization, Writing – original draft, Writing – review & editing

    Affiliations Sector de Estudos de Vectores, Instituto Nacional de Saúde (INS), Vila de Marracuene, Província de Maputo, Mozambique, Department of Zoology and Entomology, University of Pretoria (UP), Hatfield, South Africa

  • Emma F. Hocke ,

    Contributed equally to this work with: Ayubo Kampango, Emma F. Hocke

    Roles Data curation, Formal analysis, Investigation, Methodology, Software, Validation, Visualization, Writing – review & editing

    Affiliations Center for Medical Parasitology, Department of Immunology and Microbiology, University of Copenhagen, Copenhagen, Denmark, Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), Copenhagen, Denmark

  • Helle Hansson,

    Roles Data curation, Investigation, Validation, Visualization, Writing – review & editing

    Affiliations Center for Medical Parasitology, Department of Immunology and Microbiology, University of Copenhagen, Copenhagen, Denmark, Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), Copenhagen, Denmark

  • Peter Furu,

    Roles Conceptualization, Funding acquisition, Project administration, Supervision, Writing – review & editing

    Affiliation Global Health Section, Department of Public Health, University of Copenhagen, Copenhagen, Denmark

  • Khamis A. Haji †,

    † Deceased.

    Roles Investigation, Supervision, Writing – review & editing

    Affiliation Zanzibar Malaria Elimination Programme (ZAMEP), Unguja Island, Zanzibar, Tanzania

  • Jean-Philippe David,

    Roles Investigation, Methodology, Writing – review & editing

    Affiliation Laboratoire d’Ecologie Alpine (LECA), UMR 5553, Centre National de la Recherche Scientifique (CNRS)—Université Grenoble-Alpes, Grenoble, France

  • Flemming Konradsen,

    Roles Investigation, Methodology, Resources, Writing – review & editing

    Affiliation Global Health Section, Department of Public Health, University of Copenhagen, Copenhagen, Denmark

  • Fatma Saleh,

    Roles Investigation, Methodology, Resources, Supervision, Writing – review & editing

    Affiliation Department of Allied Health Sciences, School of Health and Medical Sciences, The State University of Zanzibar, Unguja Island, Zanzibar, Tanzania

  • Christopher W. Weldon,

    Roles Conceptualization, Methodology, Resources, Supervision, Writing – review & editing

    Affiliation Department of Zoology and Entomology, University of Pretoria (UP), Hatfield, South Africa

  • Karin L. Schiøler,

    Roles Conceptualization, Methodology, Project administration, Resources, Supervision, Writing – review & editing

    Affiliation Global Health Section, Department of Public Health, University of Copenhagen, Copenhagen, Denmark

  • Michael Alifrangis

    Roles Conceptualization, Methodology, Project administration, Resources, Supervision, Writing – review & editing

    Affiliations Center for Medical Parasitology, Department of Immunology and Microbiology, University of Copenhagen, Copenhagen, Denmark, Department of Infectious Diseases, Copenhagen University Hospital (Rigshospitalet), Copenhagen, Denmark


Global efforts to control Aedes mosquito-transmitted pathogens still rely heavily on insecticides. However, available information on vector resistance is mainly restricted to mosquito populations located in residential and public areas, whereas commercial settings, such as hotels are overlooked. This may obscure the real magnitude of the insecticide resistance problem and lead to ineffective vector control and resistance management. We investigated the profile of insecticide susceptibility of Aedes aegypti mosquitoes occurring at selected hotel compounds on Zanzibar Island. At least 100 adults Ae. aegypti females from larvae collected at four hotel compounds were exposed to papers impregnated with discriminant concentrations of DDT (4%), permethrin (0.75%), 0.05 deltamethrin (0.05%), propoxur (0.1%) and bendiocarb (0.1%) to determine their susceptibility profile. Allele-specific qPCR and sequencing analysis were applied to determine the possible association between observed resistance and presence of single nucleotide polymorphisms (SNPs) in the voltage-gated sodium channel gene (VGSC) linked to DDT/pyrethroid cross-resistance. Additionally, we explored the possible involvement of Glutathione-S-Transferase gene (GSTe2) mutations for the observed resistance profile. In vivo resistance bioassay indicated that Ae. aegypti at studied sites were highly resistant to DDT, mortality rate ranged from 26.3% to 55.3% and, moderately resistant to deltamethrin with a mortality rate between 79% to and 100%. However, genotyping of kdr mutations affecting the voltage-gated sodium channel only showed a low frequency of the V1016G mutation (n = 5; 0.97%). Moreover, for GSTe2, seven non-synonymous SNPs were detected (L111S, C115F, P117S, E132A, I150V, E178A and A198E) across two distinct haplotypes, but none of these were significantly associated with the observed resistance to DDT. Our findings suggest that cross-resistance to DDT/deltamethrin at hotel compounds in Zanzibar is not primarily mediated by mutations in VGSC. Moreover, the role of identified GSTe2 mutations in the resistance against DDT remains inconclusive. We encourage further studies to investigate the role of other potential insecticide resistance markers.

Author summary

Available information on mosquito resistance to insecticides is mainly restricted to residential and public areas, whereas commercial settings, such as hotels are overlooked. This may hide the real size of an insecticide resistance problem and lead to ineffective mosquito control. We investigated insecticide susceptibility of Aedes aegypti mosquitoes occurring at selected hotel compounds on Zanzibar Island. We also looked at whether resistance occurred in mosquitoes with gene mutations for two proteins (voltage-gated sodium channels and glutathione-S-transferase) that are known to cause resistance to insecticides in other parts of the world. The Ae. aegypti mosquitoes collected from hotels were highly resistant to DDT, and moderately and possibly resistant to deltamethrin and propoxur, respectively. However, resistance to these insecticides was not linked to mutations in either of the studied genes. The presence of insecticide resistance in Ae. aegypti in hotel compounds on Zanzibar is concerning and shows that these areas can act as sources of resistant mosquitoes. More needs to be done to establish the underlying causes for insecticide resistance in hotel Ae. aegypti populations, and this information can then be used to design measures that prevent resistance from becoming more widespread on Zanzibar.


Aedes-borne arboviral diseases remain one of the most important public health threats, despite substantial investments made to reduce transmission over the last two decades [1]. Aedes mosquito species, Ae. aegypti and Ae. albopictus, are the most widely distributed vectors of the five most important arboviruses associated with explosive epidemics and severe infections, i.e. dengue virus (DENV), yellow fever virus (YFV), chikungunya virus (CHIKV), Zika virus (ZIKV) [2,3] and Rift Valley fever virus (RVF) [4]. An estimated 105 million dengue infections occur each year worldwide including 51 million febrile disease cases [5]. Although the global prevalence of yellow fever has been reduced by 47% since 1980 [6], new outbreaks have re-emerged in sub-Saharan Africa and South America [710]. A similar trend has been observed with ZIKV infections in Angola, Guinea-Bissau, Cape Verde and Ethiopia [11], and with CHIKV in several other countries within the African continent [12]. Notably, the mainland of Tanzania has experienced several DENV outbreaks within the past decade. The most recent outbreak in 2019 resulted in 6,670 confirmed cases and 13 deaths [13]. Moreover, evidence for possible active transmission of DENV in Zanzibar archipelago has increased [1416]. This highlights the need for preparedness amid an increasing risk of Aedes-borne arboviral disease epidemics in Tanzania, both on the mainland and Zanzibar Islands [17,18].

General efforts to control mosquito-borne diseases (MBDs) rely heavily on chemical insecticides, mainly delivered in the form of residual spraying and long lasting insecticide treated nets [19,20]. However, large scale and intensive use of chemical insecticides, both for vector and agricultural pest control, have facilitated the emergence and expansion of resistance in vector populations against virtually all classes of insecticides registered for use in public health [21,22]. This phenomenon has threatened the gains achieved and the long-term prospect for global elimination of important MBDs, such as malaria and dengue [21]. Moreover, the higher costs associated with the development of novel, safe and efficient chemical insecticides means that existing insecticides will continue to play a pivotal role in the fight against MBDs [20]. Therefore, permanent monitoring of insecticide susceptibility profiles of local vector populations is crucial to detect the emergence of resistance, before resistant allele frequencies reach a significant level. This will support the implementation of informed and cost-saving resistance management strategies aimed at reversing resistance or maintaining susceptibility of vector species to available insecticides.

In contrast to malaria vectors [2325], the insecticide resistance profiles of Aedes mosquito species occurring in sub-Saharan Africa, including Zanzibar, remains poorly documented. In addition, routine monitoring of insecticide resistance usually targets mosquitoes in and around human settlements. This may lead to the adoption of ineffective resistance management strategies that do not cover residual foci of resistant vector populations. In a recent study, Kampango et al. [26] showed that hotels on Zanzibar Island sustain abundant populations of vector mosquito species such as Ae. aegypti and Ae. bromeliae. Pyrethroids are a widely used class of insecticides, representing more than 30% of the global insecticide market [27]. They are the only class of insecticides approved for impregnation of bed nets [28] and are preferred for spraying due to their fast action, relatively low toxicity to humans and short-lived persistence in the environment [20]. In contrast, the use of DDT is restricted to indoor spraying and only recommended under particular circumstances [29]. Both pyrethroids and DDT target the transmembrane voltage-gated sodium channels (VGSC) of excitable cells leading to knockdown and death of insects on direct contact [27]. Several Single Nucleotide Polymorphisms (SNPs) in the gene coding for VGSC have been linked to pyrethroid and DDT cross resistance in different vector populations of global health concern [30,31]. Around thirteen non-synonymous SNPs in the VGSC gene, also known as knockdown resistance (kdr) mutations, have been discovered in pyrethroid resistant Ae. aegypti. However, only four of them (V410L, S989P, V1016G/I and F1534C) have been frequently associated with DDT/pyrethroid resistance [32,33]. In Africa, the SNPs resulting in F1534C and V1016I mutations have been frequently reported [3436], whereas the V1016G mutation was only recently reported in West Africa [37]. DDT and pyrethroid cross resistance in Ae. aegypti have also been linked to metabolic resistance, manifested by increased over-expression of genes coding for detoxification enzymes that metabolize or sequester an insecticide before it reaches the target site [32]. This includes several genes of the cytochrome monooxygenase P450 family as well as glutathione-S-transferases (GSTs) [32]. Widespread occurrence of cross-resistance mutations can dramatically prevent or delay the implementation of control measures with devastating consequences, not least in epidemic situations. Therefore, thorough knowledge of resistance profiles, and underpinning molecular mechanisms can reduce the frequency of ineffective and wasted insecticide applications and reduce the likelihood of adverse effects on humans and the environment.

Our main objective was to determine the insecticide susceptibility of Ae. aegypti populations identified at selected hotel compounds on Zanzibar Island. Secondly, we investigated the potential molecular mechanisms underlying observed phenotypic resistance to pyrethroids and DDT in the local Ae. aegypti populations.

Material and methods

Ethics statement

The study received ethical clearance from the Zanzibar Health Research Institute, Ministry of Health, Zanzibar. Ref: No. ZAHREC/03/PR/Oct/2019/001. Consent was obtained from hotel management to conduct the study on their properties.

Description of study sites

Mosquito surveys were carried out from October to November 2019 in four selected hotels located in the Southeast coastal region of Zanzibar Island (Fig 1). The study sites were previously described by Kampango et al. [26,38]. Briefly, hotels were selected according to compound size (total residential and non-residential area not less than one hectare), accessibility during low and high tourism seasons, willingness to share data and willingness to accept publication of findings. For privacy reasons, hotel names are anonymized, and hotels referred to as Hotel A, which occupied an area of 6.6 hectares, Hotel B (28.1 hectares), Hotel C (2.0 hectares) and Hotel D (3.6 hectares). The hotels were located at least two kilometres apart. The rainfall regime in Zanzibar archipelago is divided into two main rainy seasons; the long rainy season usually starts from mid-March to June, and the short rainy season from November to December. Average monthly precipitation ranges from 30 mm in July to 320 mm in December, and accumulated annual rainfall can reach 1600 mm [39]. Ae. aegypti is the most dominant mosquito species found at these hotel compounds and breeds mostly in discarded plastic containers, used tires and tree holes [26,38].

Fig 1. Location of study sites on Zanzibar Island.

Dark magenta lines depict main roads. Zanzibar administrative borders shapefile and elevation raster were obtained freely from Shapefile of roads was freely obtained from

Collection of mosquito larvae

At each hotel, immature mosquitoes were surveyed for six to seven days. Details of the adopted sampling strategy have been previous described in Kampango et al. [26,38]. For relatively small containers [water volume less than 1 litre up to 5 litres, e.g., plastic bottles and beer or soda cans (metal containers)], all larvae and pupae were sampled using pipettes. For large containers (water volume >5 litres, e.g., buckets, jerry cans and ceramic pottery) with relatively few specimens, all larvae and pupae were sampled using dippers, sweep nets or bowls. For the largest containers (water volume > 20 litres, e.g., water tanks, wells, and septic tanks), specimens were collected by 10–15 random dips/sweeps. If a container was too deep (e.g., wells) samples were collected using small, suspended buckets (approx. 5 litres), filling ten buckets from different sites of the water container.

Sample processing and identification

Collected mosquito larvae and pupae were maintained at insectary environmental conditions of 27 ± 2°C and 75 ± 10% relative humidity until they emerged as adults [40]. Larvae were fed with ground adult cat food biscuits and kept at 12h light: dark regime. Adult Ae. aegypti were separated from other mosquito species using distinctive morphological features according to taxonomic keys proposed by Huang [41].

Insecticide resistance phenotypes

Insecticide susceptibility bioassays were performed according to WHO cylinder bioassay guidelines [40]. Three to five days old adult female Ae. aegypti were exposed to a diagnostic dose of organochlorine DDT (4%), pyrethroids permethrin (0.75%) and deltamethrin (0.05%) and carbamates propoxur (0.1%) and bendiocarb (0.1%). There is currently no consensus on insecticide diagnostic doses for Ae. aegypti. Therefore, we used diagnostic doses accepted for malaria vectors, as indicated by WHO insecticide bioassay guidelines [40]. The assessed insecticides were selected to represent insecticide classes in use at the hotels during the survey (S1 Table), as well as those generally used for public health, mainly deltamethrin and permethrin. DDT was intensively used for malaria control in Zanzibar in the past, but was discontinued in 1989 due to the evolution of resistance in Anopheles populations [42,43]. However, we decided to test DDT because the insecticide is still recommended by the WHO for vector control when resistance to other insecticides is high [29]. At least 100 female mosquitoes, divided into replicates of 20–25 specimens, were exposed to each type of insecticide per site. Simultaneously, 50 mosquitoes (25 per cylinder) exposed to papers impregnated with silicone oil acted as negative controls. The knockdown rate of mosquitoes exposed to insecticides was recorded every 10 minutes over an exposure period of one hour, after which mosquitoes were transferred into recovery cylinders and provided with cotton wool soaked in a 10% sucrose solution. The final mortality rate was estimated 24 hours after the end of exposure, and adjusted with Abbott´s correction formula when the mortality in the controls was between 5% and 20% [40]. When the mortality rate in the controls was more than 20%, the bioassay was repeated. Mosquitoes were considered susceptible to an insecticide if the final mortality rate was ≥ 98%, possibly resistant when the mortality rate was between 90% and 97%, and resistant if mortality was ≤ 90% [40]. All specimens exposed to DDT and pyrethroids (permethrin and deltamethrin) that were alive, and a random subsample of twenty specimens from all four hotels that were dead 24 hours after exposure were stored in silica gel at -20°C for further molecular analysis.

DNA extraction for downstream Aedes resistance genes analysis

For DNA extraction, whole mosquitoes were submerged into liquid nitrogen and ground with a pestle. The mosquito DNA was then extracted using the E-Z 96 Tissue DNA kit (Omega Bio-tek, Norcross, United States) with an adjusted initial step for arthropods as advised by the manufacturer, as follows. Initially, 180 μL of TL buffer and 20 μL of proteinase K were added and then vortexed. The samples were incubated at 56°C overnight until the mosquito was digested (exoskeleton remains excluded). Thereafter, 200 μL of BL buffer was added, and vortexed thoroughly. Finally, 200 μL of 96% ethanol was added, and once again vortexed. After these steps, the rest of the standard extraction protocol was used according to extraction kit manufacturer instructions.

Genotyping of polymorphisms in the VGSC gene

A subsample of 514 Ae. aegypti mosquitoes, comprising specimens found alive (n = 333) after being exposed to DDT (n = 296) or deltamethrin (n = 37), and dead (n = 181) after being exposed to DDT (n = 96), deltamethrin (n = 51) or permethrin (n = 34, full susceptibility), were initially analysed for the presence of kdr mutations by allele-specific qPCR (AS-qPCR). For the genotyping assay for polymorphisms V1016G/I, F1534C, and V410L, mosquito extracted DNA was investigated by AS-qPCR. Published primer mixes were used consisting of a common reverse primer and two specific primers targeting each polymorphic site [4446]. The specificity of the primers was attained for the 3’-end. Additionally, a GC-tail of different length was added to the 5’-end of either the wild type or mutant primer, making them distinguishable (S2 Table). Real-time PCR reactions were carried out using PerfeCTa FastMix SYBR green (QuantaBio, Beverly, United States). The final reaction had a total volume of 20 μL consisting of 10 μL SYBR green, 2 μL primer mix of 0.2 μM (except for the V1016G mutant primer with a concentration of 0.3 μM) of each common reverse, forward mutant primer, and a forward wild-type primer, 6 μL dH2O, and 2 μL DNA template. Cycling conditions for SNP F1534C were based on [47], but with a minor change in elongation temperature (from 60°C to 72°C), resulting in the following programme: first denaturing step of 95°C for 10 min, then denaturing 95°C for 15 seconds, annealing time of 54°C for 15 seconds, and elongation time of 72°C for 30 seconds for 40 cycles [47]. Cycling conditions for the SNP causing the V1016I change consisted of a denaturing step of 95°C for 10 min, then denaturing 94°C for 30 seconds, annealing time of 62°C for 1 min, and elongation time of 72°C for 45 seconds for 40 cycles [44]. Cycling conditions for V1016G consisted of a first denaturing step of 94°C for 10 min, then denaturing 94°C for 30 seconds, annealing time of 55°C for 30 sec, and elongation time of 72°C for 30 seconds for 40 cycles [45]. Cycling conditions for V410L consisted of a first denaturing step of 95°C for 10 min, then denaturing 95°C for 1 min, annealing time of 56°C for 20 sec, and elongation time of 72°C for 20 seconds for 40 cycles [46].

Sequencing of domains in the VGSC gene

Genetic diversity in partial sections of segment S6 in domains I-IV of the VGSC gene were investigated by Sanger sequencing. A total subset of 85 mosquitoes was used to partially sequence segment 6 of domain I-IV of the VGSC to verify the qPCR results and to explore the VGSC gene for other known (S989P, A1007G, I1011M/V, and T1520I) and possibly unknown SNPs. The subset of mosquitoes included 20 sequenced samples in domain I (10 dead, 10 alive), 85 sequenced samples in domain II (25 dead, 60 alive), 28 sequenced samples in domain III (12 dead, 16 alive), and 19 sequenced in domain IV (10 dead, 9 alive). The sections were amplified by conventional PCR, in a reaction volume of 20 μL consisting of 11 μL dH2O, 1 μL of each forward and reverse primer with a concentration of 0.20 μM, 5 μL master mix, and 2 μL of DNA. Each domain was amplified with domain-specific primers (S3 Table). Briefly, for domain I: initial denaturation of 95°C for 10 min, followed by 35 cycles of denaturation at 94°C for 30 sec, annealing at 60°C for 30 sec, and elongation at 72°C for 1 min with a final elongation of 5 min. For domains II, III, and IV: initial denaturation at 95°C for 10 min, followed by denaturation at 95°C for 30 sec, annealing at 55°C for 1 min and elongation at 72°C for 1 min repeated for 40 cycles followed by a final elongation for 1 min and 30 seconds. Amplicons were verified by electrophoresis (1% agarose gel) and purified using MicroElute Cycle-Pure Kit (Omega Bio-Tek, Norcross, United States) (S3 Table). Concentrations in ng/μL of purified amplicons were measured three times for better precision on the NanoDrop 2000/2000c Spectrophotometer (Thermo Fisher Scientific, Waltham, United States), adjusted prior to commercial sequencing (Eurofins, Luxembourg). All sequences were compared to reference sequences in Genbank using the MEGA-BLAST algorithm [48]. Consensus sequences were constructed using BioEdit 7.2 [49] with a minimum of 20 bp overlap and 85% similarity. All sequences were then aligned by the MUSCLE algorithm [50] using the Molecular Evolutionary Genetics Analysis version 10 (MEGA X) [51].

DNA sequencing of the Ae. aegypti GSTe2 gene

For amplification and sequencing of the GSTe2 gene, specific primers were constructed according to the reference gene LOC110676856 in the Ae. aegypti AaegL5.0 assembly ( Two primers were constructed to amplify the whole gene sequence and an additional four primers were constructed for Sanger sequencing of the whole gene (S1 Fig and S3 Table). The PCR reaction was carried out with a total volume of 50 μL consisting of 2 μL of each amplification primer (0.20 μM) (S2 Table), 17 μL of TEMPase Hot Start DNA Polymerase (VWR, Radnor, United States), 24 μL dH2O, and 5μL of DNA. The amplification included initial denaturation at 94°C for 15 minutes, cycling consisted of denaturation at 94°C for 1 min, annealing at 55°C for 1 min, elongation at 72°C for 2 min for 35 cycles followed by a final elongation at 72°C for 10 min. Amplicons were verified by electrophoreses (1% agarose gel) and purified using MicroElute Cycle-Pure Kit (Omega Bio-Tek, Norcross, United States). The concentration of purified amplicons was measured three times on a NanoDrop 2000/2000c Spectrophotometer (Thermo Fisher Scientific, Waltham, United States), and the average concentration used for further calculations. Concentrations in ng/μL of purified amplicons were measured on the NanoDrop 2000/2000c Spectrophotometer (Thermo Fisher Scientific, Waltham, United States), adjusted prior to commercial sequencing (Eurofins, Luxembourg). The procedure for analysis of GSTe2 sequences was similar to analysis of those VGSC as mentioned above.

Quantification of GSTe2 gene variants

For detection and quantification of copy number variations (CNV) of the GSTe2 gene, the concentration of 5 μL of dsDNA was measured on a Qubit 4 fluorometer (Thermo Fischer Scientific, Waltham, United States) using the Qubit dsDNA HS Assay Kit (Invitrogen, Waltham, United States). Since no CNV controls were available, the concentration of the samples was equalized to compare differences in signal from resistant and susceptible mosquito samples. The concentration of the dsDNA was adjusted to be 3 ng/μL. A CNV specific qPCR for GSTe2 was then performed using specific primers (S2 Table) at a concentration of 0.32 μM. The qPCR assay used a final volume of 20 μL consisting of 10 μL PerfeCTa FastMix SYBR green (QuantaBio, Beverly, United States), 6 μL of dH2O, 1μL of each primer, and 2μL of adjusted DNA. The following qPCR conditions were used: an initial denaturation at 95°C for 10 minutes, cycling consisted of 95°C denaturation for 15 seconds, annealing at 55°C for 30 seconds repeated for 40 cycles.

Statistical analysis

Log-probit regression analyses were applied to predict the probability of Ae. aegypti being knocked down by each insecticide during exposure, and to estimate the average time necessary to knockdown 50% (KDT50) and 95% (KDT95) of Ae. aegypti populations when in contact with each type of insecticide. Regression models were fitted using the R software package drc v. 3.0–1 [52]. Fisher exact tests were applied to determine whether the difference between the frequencies of insecticide resistance polymorphisms was significant. A simple t-test was applied for CNV to investigate any significant difference in qPCR Ct-values in the GSTe2 gene between susceptible and resistant Ae. aegypti samples.


Insecticide susceptibility

A total of 2,561 Ae. aegypti females were tested (Hotel A: n = 400 specimens; Hotel B: n = 836; Hotel C: n = 775; Hotel D: n = 550). The results of probit regression analysis are depicted in Fig 2 and summarized in Table 1. Mosquitoes from Hotel A were tested with only four insecticides due to too few specimens being available. In general, Ae. aegypti specimens from all four hotels showed the slowest knockdown rate when exposed to DDT (Fig 2 and Table 1). The estimated average knockdown time KDT50 of DDT exposed mosquitoes varied from 61.1 (95% CI, 56.6–65.5) among specimens from Hotel D, to a maximum of 152.7 (95% CI, 84.7–220.7) minutes for specimens from Hotel B. Mortality rates estimated twenty-four hours post-exposure ranged from 26.3% (Hotel B) to 55.3% (Hotel D) indicating presence of resistance to DDT in Ae. aegypti populations from all four hotels (Table 1). Results also suggest moderate resistance to deltamethrin, with mortality rates ranging from 79% (Hotel B) to 100% (Hotel D), and possible low levels of resistance to propoxur in the population from Hotel D, where the mortality rate was 96%. Full susceptibility was observed with permethrin and bendiocarb across all study sites (Table 1).

Fig 2. Knockdown probability of Aedes aegypti exposed to discriminant doses of insecticides.

Consistent delayed knockdown of Ae. aegypti exposed to DDT is visible at all sites, which indicates resistance.

Table 1. Estimates of mortality rates and average exposure time necessary to knockdown Aedes aegypti exposed to insecticides.

Shaded rows indicate resistance or suspected resistance requiring confirmation.

Frequency of SNPs in the VGSC gene

Molecular analysis targeting mutations in the VGSC identified only five mosquitoes that were heterozygous at V1016G. These five mosquitoes were all DDT resistant as determined by the insecticide susceptibility bioassay, and all were from the same site (Hotel B). Results of nucleotide sequences of partially sequenced segment 6 of domain I-IV of the VGSC were identical to the results obtained by qPCR. No previously described or new SNPs were detected in the VGSC gene (Fig 3).

Fig 3. Alignment of eleven representative nucleotide sequences from DDT and deltamethrin resistant Ae. aegypti samples.

The reference sequences are constructed to contain mutant codons and the sequence obtained from each of the VGSC structure domains (I—IV) are also shown. The green box highlights the five V1016G mutants detected through allele-specific qPCR in segment six (S6) of domain II that were verified by Sanger sequencing.

Polymorphisms and CNV in the GSTe2 gene

A total of 57 mosquitoes were PCR positive for presence of the GSTe2 gene. 47 confirmed positive samples were chosen for purification and sequencing, and 21 mosquitoes were successfully sequenced in the whole GSTe2 gene (Five susceptible and 16 resistant). Seven non-synonymous SNPs resulting in the following changes: L111S, C115F, P117S, E132A, I150V, E178A and A198E. These substitutions were consistently distributed across two distinct haplotypes. Thirteen of the 21 sequenced haplotypes belonged to haplotype 1 (4 susceptible and 7 resistant) and 8 belonged to haplotype 2 (1 susceptible and 7 resistant) (S2 Fig). Haplotype 2 showed low similarity (~90% homology) to available sequences in Genbank, whereas haplotype 1 showed ~96% homology. However, no statistically significant association between haplotype variant and insecticide resistance was found (p = 0.65). Regarding CNV, no statistically significant difference was found between the qPCR Ct values measured for 14 DDT susceptible (mean Ct value = 20.70 (range 19.08–22.34) and 15 DDT resistant (mean Ct value = 20.56 (range 19.43–21.76) Ae. aegypti samples (p = 0.61).


Accurate knowledge of mosquito susceptibility to available insecticidal control measures and the underlying environmental and biological mechanisms conferring resistance is crucial for selection of appropriate vector control strategies and implementation of cost-saving resistance management practices. To the best of our knowledge, this is the first attempt to investigate the status of insecticide resistance and possible association to molecular alterations in Ae. aegypti populations of the Zanzibar archipelago. In vivo susceptibility bioassays indicated that Ae. aegypti populations at four hotel sites were highly resistant to DDT, moderately resistant to deltamethrin and possibly resistant to propoxur, while fully susceptible to permethrin and bendiocarb. The insecticide susceptibility status of arboviral vectors, not least Ae. aegypti, remains poorly investigated in Tanzania, as indeed is most of the East African region (, with the few published reports limited to phenotypic characterisation of resistance. This includes a recent survey of Ae. aegypti in the Ifakara area of south-eastern Tanzania, showing full susceptibility to deltamethrin, permethrin, bendiocarb and pirimiphos-methyl [53] and a study from Dar es Salaam, in which Aedes aegypti displayed resistance to deltamethrin and lambda-cyhalothrin, but varying susceptibility to permethrin [54].

The lowest mortality rate of mosquitoes exposed to DDT was observed at Hotel B, whereas mortality rates from the remaining three hotels were statistically comparable. DDT was for many years an integral part of anti-malaria house-spraying campaigns in Zanzibar. However, it was fully withdrawn in 1988 due to high levels of DDT resistance in An. gambiae s.l., the dominant malaria vector complex in the region [42]. An exploratory survey of mosquito control practices at hotels, carried out prior to this study, did not find any evidence of recent use of DDT or analogue insecticides at the study hotels (S1 Table). However, two hotels (Hotel A and Hotel B) had implemented periodic outdoor spraying with an emulsified concentrate of the organophosphate dichlorvos and the pyrethroids deltamethrin, D-trans-allethrin (S1 Table). This may somehow explain the high level of deltamethrin resistance observed at hotel B, in particular. Space spraying with deltamethrin was also performed periodically at Hotel B. The four hotels performed aerosol (indoor) spraying using locally available commercial canned mixtures of the pyrethroids imiprothrin and cypermethrin, or with a formulation of the carbamate, propoxur. All of the above-mentioned insecticides are currently registered for use in Tanzania [55]. With 40 years having elapsed since the last widespread use of DDT to control Anopheles mosquitoes it is notable that Ae. aegypti in Zanzibar are resistant to this insecticide. However, mosquitoes can retain resistance to DDT over long periods. For instance, field studies in India on the malaria vectors An. stephensi and An. culicifacies showed that these species had retained resistance to DDT thirty years after the use of this insecticide was officially terminated [56]. Contrarily, other published evidence shows that slow reversion toward DDT susceptibility can occur in wild populations of An. stephensi and Ae. aegypti when the DDT pressure is removed [57,58]. As such, we propose that the past intense use of dieldrin (1958–1961), and thereafter DDT (until 1988), as part of malaria eradication campaigns in Zanzibar [42,59], may have led to unintended selection of DDT resistance in local Ae. aegypti populations.

Allele-specific qPCR-based assays detected the presence of heterozygous V1016G mutant alleles in the VGSC gene of five specimens of Ae. aegypti (found at Hotel B). No other investigated point mutations commonly found in Ae. aegypti were detected. This finding contrasts with the outcome of our phenotypic resistance bioassay, which indicated higher resistance to DDT and moderate resistance to deltamethrin. The geographical occurrence of the V1016G mutation is expanding in the Middle East [60], southeast Asia [45,6164] and Latin America [65]. Notably, it was recently reported in Cameroon, West Africa [37]. The origin of V1016G/I mutations in African populations of Ae. aegypti remains poorly understood. Fan et al. [66] proposed that the variant may have arisen from genetic recombination in the evolution of kdr alleles or by introduction from another continent. We did not observe a clear association between the presence of the V1016G alleles and reduced susceptibility to DDT and deltamethrin in Ae. aegypti from Zanzibar. The lower frequency of V1016G alleles suggests recent introduction or occurrence of this type of kdr mutation in Ae. aegypti populations at the studied sites. It may also suggest reduced fitness of V1016G alleles and might be unfavourable to maintain. As noted earlier, there is intensive pesticide use at the studied hotels, which may have applied selection pressure for resistance in the Ae. aegypti populations. Therefore, further investigations are encouraged to understand factors driving the emergence and propagation of kdr mutations in order to prevent their successful establishment and expansion throughout the Zanzibar archipelago.

Several genes of the family cytochrome P450 and glutathione-S-transferases (GSTes) have been associated with pyrethroid and DDT resistance in Ae. aegypti populations [35,67,68]. For instance, a study by Helvecio et al. [69] found that several SNPs in GSTe2 correlated with temephos resistance in Ae. aegypti. However, in this study, sequencing analyses indicated no significant correlation between Ae. aegypti resistance to DDT and presence of GSTe2 SNPs observed (L111S, C115F, P117S, E132A, I150V, E178A, and A198E). In Senegal, Sene et al. [68] also reported higher phenotypic resistance to DDT and permethrin in Ae. aegypti but, similar to this study, no kdr mutations were detected. The authors found overexpression of genes of the P450 family namely CYP6BB2, CYP9J26 and CYP9J32, associated with resistance to pyrethroids and DDT, and also high overexpression of the GSTD4 glutathione-S-transferase gene, associated with resistance to DDT. This study did not investigate the potential role of P450 family genes since, in general, Ae. aegypti showed susceptibility to the two types of pyrethroids tested, excepting at Hotel B where resistance to deltamethrin was detected. Using lab-based resistance selection experiments, Smith et al. [67] showed that cytochrome P450 genes can also mediate cross resistance to pyrethroids and organophosphates in Ae. aegypti either in the presence or absence of kdr mutations, underscoring the complexity of molecular mechanisms of insecticide resistance in Ae. aegypti populations. No significant association was detected between GSTe2 polymorphisms and phenotypic resistance to DDT, suggesting that a potentially different resistance mechanism may be involved. There are many other poorly investigated resistance mechanisms to pyrethroids and DDT, such as the recently described cuticular-based resistance to pyrethroids and DDT in Ae. aegypti from Cameroon [70]. Finally, previous studies with An. arabiensis and An. gambiae s.s. reported an eight-fold higher increase of DDT-dehydrochlorinase activity of glutathione S-transferases in resistant populations compared to susceptible ones [71]. As such, we encourage further investigations into the probable involvement of other, less common, mechanisms that might underpin pyrethroid/DDT resistance in Ae. aegypti and similar vectors of public health concern.

Concluding remarks

Our study shows that resistance to pyrethroids and DDT is present in Ae. aegypti of Zanzibar. It also shows, for the first time, the presence of the kdr mutation V1016G in the region. However, any association between the V1016G mutation and resistance to pyrethroids and DDT in Ae. aegypti populations remains inconclusive due to low detection frequency of the resistant allele. Emergence of V1016G kdr mutations in Zanzibar should be of great concern given the intense use of pyrethroids in and around hotels. Therefore, efforts should be made to reduce selection pressure by replacing the use of chemical control by environmental mosquito management practices. Successful expansion of V1016G throughout the archipelago would compromise the effectiveness of vector control by both type 1 and type 2 pyrethroids with devastating consequences in the event of an Aedes-borne disease epidemic. Our findings also highlight the need for continued and comprehensive investigations on other potential mechanisms for pyrethroid resistance in local Ae. aegypti populations. Moreover, the risk of large-scale use of insecticides for mosquito control at private facilities, outside direct government control, should be thoroughly investigated.

Supporting information

S1 Fig. Overview of the GSTe2 transcript and constructed primers used for amplification and sequencing.

Arrows indicate direction of synthesis.


S2 Fig. Alignment starting from position 110 to 200 of two representative haplotype protein sequences with SNPs L111S, C115F, P117S, E132A, I150V, E178A, A198E present.


S1 Table. Mosquito control practices, type, quantity and frequency of chemical insecticides utilization at hotel compounds.


S2 Table.

Primers used for allele-specific qPCR of the VGSC (A) and CNV assay of the GSTe2 gene (B).


S3 Table.

Primers used to amplify and sequence partial segment 6 in domain I-IV (A) and whole GSTe2 gene (B).


S1 Data. Excel format dataset containing records of in vivo susceptibility bioassays.



The authors would like to thank all members of the EnSuZa project for constructive discussions and support during the study. We would also like to thank the hotel managers and the Zanzibar Association of Tourism Investors for their unconditional support and permanent collaboration.


  1. 1. WHO: Global vector control response 2017–2030. Geneva, Switerland: World Health Organization; 2017: 53.
  2. 2. Powell JR: Mosquitoes on the move. Science 2016, 354(6315):971. pmid:27884993
  3. 3. Paixão ES, Teixeira MG, Rodrigues LC: Zika, chikungunya and dengue: the causes and threats of new and re-emerging arboviral diseases. BMJ global health 2018, 3(Suppl 1):e000530–e000530. pmid:29435366
  4. 4. Tantely LM, Boyer S, Fontenille D: A review of mosquitoes associated with Rift Valley fever virus in Madagascar. The American Journal of Tropical Medicine and Hygiene 2015, 92(4):722–729. pmid:25732680
  5. 5. Cattarino L, Rodriguez-Barraquer I, Imai N, Cummings DAT, Ferguson NM: Mapping global variation in dengue transmission intensity. Scince Translational Medicine 2020, 12(eaax4144). pmid:31996463
  6. 6. Gaythorpe KAM, Hamlet A, Jean K, Garkauskas Ramos D, Cibrelus L, Garske T, Ferguson N: The global burden of yellow fever. eLife 2021, 10:e64670. pmid:33722340
  7. 7. Grobbelaar AA, Weyer J, Moolla N, Jansen van Vuren P, Moises F, Paweska JT: Resurgence of Yellow Fever in Angola, 2015–2016. Emerging Infectious Diseases 2016, 22(10):1854–1855. pmid:27536787
  8. 8. Ingelbeen B, Weregemere NA, Noel H, Tshapenda GP, Mossoko M, Nsio J, Ronsse A, Ahuka-Mundeke S, Cohuet S, Kebela BI: Urban yellow fever outbreak-Democratic Republic of the Congo, 2016: Towards more rapid case detection. PLoS Neglected Tropical Diseases 2018, 12(12):e0007029–e0007029. pmid:30532188
  9. 9. Cunha M, Duarte-Neto AN, Pour SZ, Ortiz-Baez AS, Černý J, Pereira B, Braconi CT, Ho YL, Perondi B, Sztajnbok J et al: Origin of the São Paulo Yellow Fever epidemic of 2017–2018 revealed through molecular epidemiological analysis of fatal cases. Scientific Reports 2019, 9(1):20418. pmid:31892699
  10. 10. Gaythorpe KA, Hamlet A, Cibrelus L, Garske T, Ferguson NM: The effect of climate change on yellow fever disease burden in Africa. eLife 2020, 9:e55619. pmid:32718436
  11. 11. Zika Epidemiology Report: Global overview []
  12. 12. Russo G, Subissi L, Rezza G: Chikungunya fever in Africa: a systematic review. Pathogens and Global Health 2020, 114(3):111–119. pmid:32191166
  13. 13. Weekly bulletin on outbreaks and other emergencies []
  14. 14. Gautret P, Simon F, Hervius Askling H, Bouchaud O, Leparc-Goffart I, Ninove L, Parola P, EuroTravNet: Dengue type 3 virus infections in European travellers returning from the Comoros and Zanzibar, February-April 2010. Euro surveillance: bulletin Europeen sur les maladies transmissibles = European communicable disease bulletin 2010, 15(15):19541. pmid:20429996
  15. 15. Ali MA, James OC, Mohamed AA, Joachim A, Mubi M, Omodior O: Etiologic Agents of Fever of Unknown Origin Among Patients Attending Mnazi Mmoja Hospital, Zanzibar. Journal of Community Health 2020, 45(5):1073–1080. pmid:32399732
  16. 16. Shauri HS, Ngadaya E, Senkoro M, Buza JJ, Mfinanga S: Seroprevalence of Dengue and Chikungunya antibodies among blood donors in Dar es Salaam and Zanzibar, Tanzania: a cross-sectional study. BMC Infectious Diseases 2021, 21(1):911. pmid:34488666
  17. 17. Mboera LE, Mweya CN, Rumisha SF, Tungu PK, Stanley G, Makange MR, Misinzo G, De Nardo P, Vairo F, Oriyo NM: The Risk of Dengue Virus Transmission in Dar es Salaam, Tanzania during an Epidemic Period of 2014. PLoS Neglected Tropical Diseases 2016, 10(1):e0004313. pmid:26812489
  18. 18. Saleh F, Kitau J, Konradsen F, Kampango A, Abassi R, Schiøler KL: Epidemic risk of arboviral diseases: Determining the habitats, spatial-temporal distribution, and abundance of immature Aedes aegypti in the Urban and Rural areas of Zanzibar, Tanzania. PLoS Neglected Tropical Diseases 2020, 14(12):e0008949. pmid:33284806
  19. 19. Rozendaal JA: Vector Control: Methods for use by individuals and communities. Geneva, Switzerland: World Health Organization; 1997.
  20. 20. van Emden HF, Service MW: Pest and Vector Control. United Kingdom: Cambridge University Press; 2004. pmid:15603586
  21. 21. Hemingway J, Ranson H, Magill A, Kolaczinski J, Fornadel C, Gimnig J, Coetzee M, Simard F, Roch DK, Hinzoumbe CK et al: Averting a malaria disaster: will insecticide resistance derail malaria control? Lancet 2016, 387(10029):1785–1788. pmid:26880124
  22. 22. The Arthropod Pesticide Resistance Database. []
  23. 23. Haji KA, Khatib BO, Smith S, Ali AS, Devine GJ, Coetzee M, Majambere S: Challenges for malaria elimination in Zanzibar: pyrethroid resistance in malaria vectors and poor performance of long-lasting insecticide nets. Parasites & Vectors 2013, 6:82. pmid:23537463
  24. 24. Haji KA, Thawer NG, Khatib BO, Mcha JH, Rashid A, Ali AS, Jones C, Bagi J, Magesa SM, Ramsan MM et al: Efficacy, persistence and vector susceptibility to pirimiphos-methyl (Actellic 300CS) insecticide for indoor residual spraying in Zanzibar. Parasites & Vectors 2015, 8(1):628.
  25. 25. Jones CM, Haji KA, Khatib BO, Bagi J, Mcha J, Devine GJ, Daley M, Kabula B, Ali AS, Majambere S et al: The dynamics of pyrethroid resistance in Anopheles arabiensis from Zanzibar and an assessment of the underlying genetic basis. Parasites & Vectors 2013, 6(1):343. pmid:24314005
  26. 26. Kampango A, Furu P, Sarath DL, Haji KA, Konradsen F, Schioler KL, Alifrangis M, Weldon C, Saleh F: Targeted elimination of species-rich larval habitats can rapidly collapse mosquito species arbovirus vector populations in Zanzibar. Medical and Veterinary Entomology 2021, 35(4):523–533 pmid:33970496
  27. 27. Zlotikin E: Insecticides Affecting Voltage-Gated Ion Channels. In: Biochemical Sites of Insecticide Action and Resistance. Edited by Ishaaya I: Springer; 2001: 43–69.
  28. 28. WHO: Pesticides and their application for the control of vectors and pests of public heath importance, 6 edn. Geneva: WHO; 2006.
  29. 29. Rehwagen C: WHO recommends DDT to control malaria. BMJ 2006, 333(7569):622–622. pmid:16990319
  30. 30. Du Y, Nomura Y, Zhorov BS, Dong K: Sodium Channel Mutations and Pyrethroid Resistance in Aedes aegypti. Insects 2016, 7(4):60. pmid:27809228
  31. 31. Ndiath MO: Insecticides and Insecticide Resistance. Methods in Molecular Biology 2019, 2013:287–304. pmid:31267507
  32. 32. Moyes CL, Vontas J, Martins AJ, Ng LC, Koou SY, Dusfour I, Raghavendra K, Pinto J, Corbel V, David J-P et al: Contemporary status of insecticide resistance in the major Aedes vectors of arboviruses infecting humans. PLoS Neglected Tropical Diseases 2017, 11(7):e0005625. pmid:28727779
  33. 33. Zuharah WF, Sufian M: The discovery of a novel knockdown resistance (kdr) mutation A1007G on Aedes aegypti (Diptera: Culicidae) from Malaysia. Scientific Reports 2021, 11(1):5180. pmid:33664425
  34. 34. Kawada H, Higa Y, Futami K, Muranami Y, Kawashima E, Osei JHN, Sakyi KY, Dadzie S, de Souza DK, Appawu M et al: Discovery of Point Mutations in the Voltage-Gated Sodium Channel from African Aedes aegypti Populations: Potential Phylogenetic Reasons for Gene Introgression. PLoS Neglected Tropical Diseases 2016, 10(6):e0004780. pmid:27304430
  35. 35. Sombie A, Saiki E, Yameogo F, Sakurai T, Shirozu T, Fukumoto S, Sanon A, Weetman D, McCall PJ, Kanuka H et al: High frequencies of F1534C and V1016I kdr mutations and association with pyrethroid resistance in Aedes aegypti from Somgande (Ouagadougou), Burkina Faso. Tropical Medicine and Health 2019, 47:2. pmid:30787670
  36. 36. Yougang AP, Kamgang B, Bahun TAW, Tedjou AN, Nguiffo-Nguete D, Njiokou F, Wondji CS: First detection of F1534C knockdown resistance mutation in Aedes aegypti (Diptera: Culicidae) from Cameroon. Infectious Diseases of Poverty 2020, 9(1):152. pmid:33138860
  37. 37. Djiappi-Tchamen B, Nana-Ndjangwo MS, Mavridis K, Talipouo A, Nchoutpouen E, Makoudjou I, Bamou R, Mayi AM, Awono-Ambene P, Tchuinkam T et al: Analyses of Insecticide Resistance Genes in Aedes aegypti and Aedes albopictus Mosquito Populations from Cameroon. Genes 2021, 12(6). pmid:34071214
  38. 38. Kampango A, Furu P, Sarath DL, Haji KA, Konradsen F, Schiøler KL, Alifrangis M, Saleh F, Weldon CW: Risk factors for occurrence and abundance of Aedes aegypti and Aedes bromeliae at hotel compounds in Zanzibar. Parasites & Vectors 2021, 14(1):544. pmid:34686195
  39. 39. Climate & Soils []
  40. 40. WHO: Test procedures for insecticide resistance monitoring in malaria vectors, bio-efficacy and persistence of insecticide on treated surfaces. Geneva, Switzerland: WHO; 2016.
  41. 41. Huang Y-M: The subgenus Stegomyia of Aedes in the Afrotropical Region with keys to the species (Diptera: Culicidae). Zootaxa 2004, 700:1–120.
  42. 42. Schwartz E, Pener H, Issa SM, Golenser J: An Overview of the Malaria Situation in Zanzibar. Journal of Community Health 1997, 22(1):33–44. pmid:9120045
  43. 43. Prapanthadara L-A, Hemingway J, Ketterman AJ: DDT-resistance in Anopheles gambiae (Diptera: Culicidae) from Zanzibar, Tanzania, based on increased DDT-dehydrochlorinase activity of glutathione S-transferases. Bulletin of Entomological Research 2009, 85(2):267–274.
  44. 44. Martins AJ, Lima JBP, Peixoto AA, Valle D: Frequency of Val1016Ile mutation in the voltage-gated sodium channel gene of Aedes aegypti Brazilian populations. Tropical Medicine & International Health 2009, 14(11):1351–1355. pmid:19735371
  45. 45. Stenhouse SA, Plernsub S, Yanola J, Lumjuan N, Dantrakool A, Choochote W, Somboon P: Detection of the V1016G mutation in the voltage-gated sodium channel gene of Aedes aegypti (Diptera: Culicidae) by allele-specific PCR assay, and its distribution and effect on deltamethrin resistance in Thailand. Parasites & Vectors 2013, 6(1):253.
  46. 46. Villanueva-Segura K, Ponce-Garcia G, Lopez-Monroy B, Mora-Jasso E, Perales L, Gonzalez-Santillan FJ, Ontiveros-Zapata K, Davila-Barboza JA, Flores AE: Multiplex PCR for simultaneous genotyping of kdr mutations V410L, V1016I and F1534C in Aedes aegypti (L.). Parasites & Vectors 2020, 13(1):325. pmid:32586378
  47. 47. Linss JGB, Brito LP, Garcia GA, Araki AS, Bruno RV, Lima JBP, Valle D, Martins AJ: Distribution and dissemination of the Val1016Ile and Phe1534Cys Kdr mutations in Aedes aegypti Brazilian natural populations. Parasites & Vectors 2014, 7(1):25. pmid:24428880
  48. 48. Morgulis A, Coulouris G, Raytselis Y, Madden TL, Agarwala R, Schäffer AA: Database indexing for production MegaBLAST searches. Bioinformatics 2008, 24(16):1757–1764. pmid:18567917
  49. 49. Hall TA: BioEdit: A User-Friendly Biological Sequence Alignment Editor and Analysis Program for Windows 95/98/NT. Nucleic Acids Symposium Series 1999, 41:95–98.
  50. 50. Edgar RC: MUSCLE: multiple sequence alignment with high accuracy and high throughput. Nucleic Acids Research 2004, 32(5):1792–1797. pmid:15034147
  51. 51. Kumar S, Stecher G, Li M, Knyaz C, Tamura K: MEGA X: Molecular Evolutionary Genetics Analysis across Computing Platforms. Molecular Biology and Evolution 2018, 35(6):1547–1549. pmid:29722887
  52. 52. Ritz C, Streibig JC: Bioassay Analysis usong R. Journal of Statistical Software 2005, 12(5).
  53. 53. Kahamba NF, Limwagu AJ, Mapua SA, Msugupakulya BJ, Msaky DS, Kaindoa EW, Ngowo HS, Okumu FO: Habitat characteristics and insecticide susceptibility of Aedes aegypti in the Ifakara area, south-eastern Tanzania. Parasites & Vectors 2020, 13(1):53. pmid:32033619
  54. 54. Mathias L, Baraka V, Philbert A, Innocent E, Francis F, Nkwengulila G, Kweka EJ: Habitat productivity and pyrethroid susceptibility status of Aedes aegypti mosquitoes in Dar es Salaam, Tanzania. Infectious Diseases of Poverty 2017, 6(1):102. pmid:28595653
  55. 55. List of Registered Pesticides to be used in the United Republic of Tanzania 2020 []
  56. 56. Raghavendra K VV, Srivastava HC, Gunasekaran K, Sreehari U, Dash AP.: Persistence of DDT, malathion & deltamethrin resistance in Anopheles culicifacies after their sequential withdrawal from indoor residual spraying in Surat district, India. Indian Journal of Medical Research 2010, 132:260–264.
  57. 57. Abedi ZH, Brown AWA: Development and reversion of DDT resistance in Aedes aegypti. Canadian Journal of Genentic and Cytology 1960, 2(3):252–261.
  58. 58. Bang YH, Jatanasen S, Tonn RJ: Development and reversion of DDT resistance in an Aedes aegypti population in Bangkok, Thailand. Bulletin of the World Health Organization 1971, 45(3):404–410. pmid:5316918
  59. 59. Lines JD, Nassor NS: DDT resistance in Anopheles gambiae declines with mosquito age. Medical and Veterinary Entomolgy 1991, 5:261–265. pmid:1768918
  60. 60. Al Nazawi AM, Aqili J, Alzahrani M, McCall PJ, Weetman D: Combined target site (kdr) mutations play a primary role in highly pyrethroid resistant phenotypes of Aedes aegypti from Saudi Arabia. Parasites & Vectors 2017, 10(1):161. pmid:28347352
  61. 61. Kawada H, Higa Y, Komagata O, Kasai S, Tomita T, Thi Yen N, Loan LL, Sánchez RAP, Takagi M: Widespread Distribution of a Newly Found Point Mutation in Voltage-Gated Sodium Channel in Pyrethroid-Resistant Aedes aegypti Populations in Vietnam. PLoS Neglected Tropical Diseases 2009, 3(10):e527. pmid:19806205
  62. 62. Kawada H, Oo SZM, Thaung S, Kawashima E, Maung YNM, Thu HM, Thant KZ, Minakawa N: Co-occurrence of Point Mutations in the Voltage-Gated Sodium Channel of Pyrethroid-Resistant Aedes aegypti Populations in Myanmar. PLoS Neglected Tropical Diseases 2014, 8(7):e3032. pmid:25077956
  63. 63. Srisawat R, Komalamisra N, Eshita Y, Zheng M, Ono K, Itoh TQ, Matsumoto A, Petmitr S, Rongsriyam Y: Point mutations in domain II of the voltage-gated sodium channel gene in deltamethrin-resistant Aedes aegypti (Diptera: Culicidae). Applied Entomology and Zoology 2010, 45(2):275–282.
  64. 64. Li C-X, Kaufman PE, Xue RD, Zhao MH, Wang G, Yan T, Guo XX, Zhang YM, Dong YD, Xing D et al: Relationship between insecticide resistance and kdr mutations in the dengue vector Aedes aegypti in Southern China. Parasites & Vectors 2015, 8(325). pmid:26068925
  65. 65. Murcia O, Henríquez B, Castro A, Koo S, Young J, Márquez R, Pérez D, Cáceres L, Valderrama A: Presence of the point mutations Val1016Gly in the voltage-gated sodium channel detected in a single mosquito from Panama. Parasites & Vectors 2019, 12(1):62.
  66. 66. Fan Y, O’Grady P, Yoshimizu M, Ponlawat A, Kaufman PE, Scott JG: Evidence for both sequential mutations and recombination in the evolution of kdr alleles in Aedes aegypti. PLoS Neglected Tropical Diseases 2020, 14(4):e0008154. pmid:32302303
  67. 67. Smith LB, Sears C, Sun H, Mertz RW, Kasai S, Scott JG: CYP-mediated resistance and cross-resistance to pyrethroids and organophosphates in Aedes aegypti in the presence and absence of kdr. Pesticide Biochemistry and Physiology 2019, 160:119–126. pmid:31519246
  68. 68. Sene NM, Mavridis K, Ndiaye EH, Diagne CT, Gaye A, Ngom EHM, Ba Y, Diallo D, Vontas J, Dia I et al: Insecticide resistance status and mechanisms in Aedes aegypti populations from Senegal. PLoS Neglected Tropical Diseases 2021, 15(5):e0009393. pmid:33970904
  69. 69. Helvecio E, Romão TP, de Carvalho-Leandro D, de Oliveira IF, Cavalcanti AEHD, Reimer L, de Paiva Cavalcanti M, de Oliveira APS, Paiva PMG, Napoleão THet al: Polymorphisms in GSTE2 is associated with temephos resistance in Aedes aegypti. Pesticide Biochemistry and Physiology 2020, 165:104464. pmid:32359546
  70. 70. Borel DT, Nana-Ndjangwo Stella M, Vasileia B, Konstantinos M, Makoudjou I, Awono-Ambene P, Charles W, Tchuinkam T, John V, Antonio-Nkondjio C: First Evidence of the Implication of Cuticular Based Mechanisms in Aedes Aegypti Populations Resistant To Pyrethroids and DDT in Cameroon. Research Square 2021.
  71. 71. Prapanthadara L, Hemingway J, Ketterman A: DDT-resistance in Anopheles gambiae (Diptera: Culicidae) from Zanzibar, Tanzania, based on increased DDT-dehydrochlorinase activity of glutathione S-transferases. Bulletin of Entomological Research 1995, 85(2):267–274.