Next Article in Journal
Appressoria-Producing Sordariomycetes Taxa Associated with Jasminum Species
Previous Article in Journal
Dynamic and Seasonal Distribution of Enteric Viruses in Surface and Well Water in Riyadh (Saudi Arabia)
Previous Article in Special Issue
A Participatory Approach in Assessing the Knowledge, Attitude, and Practices (KAP) of Stakeholders and Livestock Owners about Ticks and Tick-Borne Diseases from Sindh, Pakistan
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Systematic Review

Mosquitoes, Lymphatic Filariasis, and Public Health: A Systematic Review of Anopheles and Aedes Surveillance Strategies

Arumugam Bhuvaneswari
Ananganallur Nagarajan Shriram
Kishan Hari K. Raju
1 and
Ashwani Kumar
Indian Council of Medical Research—Vector Control Research Centre, Puducherry 605006, India
Center for Global Health Research, Saveetha Medical College and Hospital, Saveetha Institute of Medical and Technical Sciences, Saveetha University, Chennai 605102, India
Author to whom correspondence should be addressed.
Pathogens 2023, 12(12), 1406;
Submission received: 21 September 2023 / Revised: 6 November 2023 / Accepted: 15 November 2023 / Published: 29 November 2023


Lymphatic Filariasis (LF) affects over 120 million people in 72 countries, with sub-periodic filariasis common in the Pacific. Wuchereria bancrofti has three physiological races, each with a unique microfilarial periodicity, and each race is isolated to a specific geographical region. Sub-periodic W. bancrofti is transmitted by various Aedes mosquito species, with Aedes polynesiensis and Aedes samoanus being the primary vectors in Samoa. The Aedes scutellaris and Aedes kochi groups are also important vectors in the South Pacific Islands. Anopheles species are important vectors of filariasis in rural areas of Asia and Africa. The Anopheles gambiae complex, Anopheles funestus, and the Anopheles punctulatus group are the most important vectors of W. bancrofti. These vectors exhibit indoor nocturnal biting behaviour and breed in a variety of habitats, including freshwater, saltwater, and temporary water bodies. Effective vector surveillance is central to LF control and elimination programs. However, the traditional Human Landing Collection (HLC) method, while valuable, poses ethical concerns and risks to collectors. Therefore, this review critically analyses alternative trapping tools for Aedes and Anopheles vectors in LF-endemic regions. We looked at 14 research publications that discussed W. bancrofti vector trapping methods. Pyrethrum Spray Catches (PSC), one of the seven traps studied for Anopheles LF vectors, was revealed to be the second most effective strategy after HLC, successfully catching Anopheles vectors in Nigeria, Ghana, Togo, and Burkina Faso. The PSC method has several drawbacks, such as the likelihood of overlooking exophilic mosquitoes or underestimating Anopheles populations. However, exit traps offered hope for capturing exophilic mosquitoes. Anopheles populations could also be sampled using the Anopheles Gravid Trap (AGT). In contrast, the effectiveness of the Double Net Traps (DNT) and the CDC Light Trap (CDC LT) varied. Gravid mosquito traps like the OviArt Gravid Trap (AGT) were shown to be useful tools for identifying endophilic and exophilic vectors during the exploration of novel collection techniques. The Stealth trap (ST) was suggested for sampling Anopheles mosquitoes, although specimen damage may make it difficult to identify the species. Although it needs more confirmation, the Ifakara Tent Trap C design (ITT-C) showed potential for outdoor mosquito sampling in Tanzania. Furvela tent traps successfully captured a variety of Anopheles species and are appropriate for use in a variety of eco-epidemiological settings. By contrast, for Aedes LF vectors, no specific sampling tool was identified for Aedes niveus, necessitating further research and development. However, traps like the Duplex cone trap, Resting Bucket Trap (RB), and Sticky Resting Bucket trap (SRB) proved effective for sampling Aedes albopictus, offering potential alternatives to HLC. This review emphasises the value of looking into alternative trapping methods for Aedes and Anopheles vectors in the LF-endemic region. Further research is required to determine the efficacy of novel collection techniques in various contexts, even if PSC and AGT show promise for sampling Anopheles vectors. The identified traps, along with ongoing research, provide valuable contributions to vector surveillance efforts in LF-endemic regions, enabling LF control and elimination strategies to advance.

1. Introduction

The elimination of Lymphatic Filariasis (LF) and malaria, two mosquito-borne diseases, is planned for 2030 through preventive chemotherapy and vector control measures [1,2]. Sensitive, quick, and species-specific diagnostic methods for parasite detection in humans and vectors are required to confirm the cessation of transmission, and they are crucial for determining disease prevalence [3,4,5,6,7,8]. These technologies define intervention endpoints and confirm the efficacy of mass drug administration (MDA) programmes [9].
Medical entomologists have been utilizing molecular xenomonitoring (Mx) for decades to assess the risk of transmission of vector-borne illnesses (VBDs) by detecting human infections in arthropod vectors. Mx, a powerful tool for tracking disease transmission, as it can detect microfilarial DNA in mosquito samples even in trace amounts, served as a proxy for human infection surveillance related to VBDs [10,11,12,13,14,15,16,17]. Creating xenomonitoring systems for programmatic use that are accurate, reliable, and cost-effective can be challenging due to the diverse vector–parasite combinations.
In 2022, the World Health Organization (WHO) estimated that there were 120 million people infected with W. bancrofti and 12 million people infected with Brugia malayi. The disease is most common in tropical and subtropical regions, and it is estimated that 80% of all cases are found in Africa (World Health Organization, Global Programme to Eliminate Lymphatic Filariasis. Progress Report 2022. Geneva: World Health Organization; 2023). Research on mosquito sampling techniques for Anopheles and Aedes vectors of diurnally sub-periodic (DspWB) and nocturnally periodic (NpWb) W. bancrofti (Wb) is lacking, despite the abundance of literature emphasizing the use of Mx for LF elimination with a Culex quinquefasciatus (C.q.)—W. bancrofti (Wb) vector–parasite combination. Choosing the best Mx tool for public health programmes and surveillance is critical, even though Anopheles and Aedes vectors are less responsible for the LF burden than Culex-transmitted filariasis.
The epidemiological significance of parasite DNA prevalence in local vector mosquitoes can be demonstrated by improving mosquito sampling techniques for local mosquito vector species. This will lead to sufficient sample sizes and more accurate prevalence estimates. Consequently, these enhancements have increased the operational value of LF and malaria elimination programmes.
The utilisation of different mosquito sampling techniques by vector control programmes in diverse eco-epidemiological settings may lead to biased assessments of species diversity and abundance. This systematic review critically examines the prevalence, geographic distribution, and bio-ecology of sub-periodic filariasis, specifically focusing on Anopheles spp. and Aedes spp. The review also investigates and compares the effectiveness of various Mx traps. The primary objective is to identify the most suitable Mx tool for various tasks, considering the available information on sampling techniques while taking into account the biology of the mosquito species acting as vectors.

1.1. Prevalence and Distribution of Lymphatic Filariasis

Over 120 million people in 72 countries across Asia, Africa, the Western Pacific, and parts of the Caribbean and South America are affected by LF. Sub-periodic filariasis caused by W. bancrofti is particularly prevalent in the Pacific region, including the islands of Tahiti, Samoa [18], Tonga, and Fiji, Australia, New Guinea, and the nearby Melanesia, Micronesia, and Polynesia islands are all part of the Pacific region [19].
The South Pacific islands exhibit a similar pattern of patchy filariasis distribution as the rest of the world [20,21]. Early data from Fiji [22] revealed a low frequency of 6.4% among residents of the Labasa, compared to a high prevalence of 25.2% in Taveuni. The prevalence rates in various communities appeared to be influenced by the behaviours of the population and the proximity of densely populated vector areas to human settlements, increasing the likelihood of infection transmission. Particularly in the Nancowry group of islands, Nicobar district, the day-biting Aedes (Downsiomyia) niveus transmit the diurnally sub-periodic W. bancrofti disease in India [23,24,25,26,27,28].

1.1.1. Physiological Races of W. bancrofti

There are three physiological races of W. bancrofti, each having a unique microfilarial periodicity. Apart from the Polynesian sub-region, the nocturnally periodic race is widely distributed in tropical and sub-tropical environments worldwide. A nocturnally sub-periodic race is found in the jungle regions of West Thailand, while the diurnally sub-periodic race is isolated to the Polynesian sub-region [29,30,31]. Each race has unique intermediate hosts, and the microfilarial periodicity of each race coincides with the biting rhythm of the principal vector mosquitoes.

1.1.2. Types of W. bancrofti Infection Identified, Based on Their Ecological Distribution

  • The Culex fatigans type, transmitted by the Culex pipiens complex, including races like Culex fatigans, is known as Culex quinquefasciatus and Culex molestus. This is the most widely distributed ecological type [29].
  • The Anopheles type, in tropical Africa, Anopheles gambiae, Anopheles funestus, and related species are the principal vectors of W. bancrofti in rural areas, while other regions have vectors such as Anopheles maculatus, Anopheles whartoni, Anopheles flavirostris, and Anopheles punctulatus [29].
  • The Aedes (Finlaya) poecilus type is responsible for transmitting nocturnally periodic W. bancrofti in the Phillipines. Additionally, the Aedes (Finlaya) kochi group serves as efficient vectors for the diurnally sub-periodic race in the Polynesian region.
  • The Aedes (Ochlerotatus) vigilax type is the primary vector of the diurnally sub-periodic race of W. bancrofti endemic in the New Caledonian region.
  • Aedes (Stegomyia) polynesiensis-type mosquitoes are the principal vectors of the diurnally sub-periodic form W. bancrofti in the Polynesian region.

Breeding Ecology, Biology and Implication in the Transmission

In Samoa, sub-periodic W. bancrofti is primarily transmitted by two vectors: A. polynesiensis and A. samoanus [12,32,33,34]. A. polynesiensis, a container breeder, and A. samoanus, a leaf axil breeder, are nocturnal species restricted to the Samoa islands [18,35,36,37,38]. Another member of the kochi group, A. tutuilae, breeds exclusively in pandanus leaf axils and is also a nocturnal species found only in Samoa [18].
In the South Pacific Islands, the sub-periodic W. bancrofti is primarily transmitted by A. pseudoscutellaris, and along with A. polynesiensis, they were efficient transmitters, and the night-biting A. fijiensis of the kochi group was equally efficient [22,39,40]. C. quinquefasciatus, the vector of periodic W. bancrofti, could also transmit sub-periodic W. bancrofti to a limited extent in the Society Islands and Fiji [22,40]. Further studies by Burnett [41], Rossen [42], and Symes [22,40] confirmed these findings, emphasizing the need for re-examination potential vectors, particularly in areas where members of the A. scutellaris and kochi groups have been reported [43].
In India’s Andaman and Nicobar Islands, DspWb is transmitted by the day-biting Aedes (Downsiomyia) niveus, a tree-hole breeder, with diurnal biting behaviour. A. polynesiensis in American Samoa and Western Samoa exhibits similar biting behaviour, primarily diurnal with a small proportion biting at night [27,44]. Sampling A. niveus presents a significant challenge for researchers and LF control programmes conducting surveillance [45].
In Samoa, efficient vectors of sub-periodic W. bancrofti include A. polynesiensis, A. upolensis, and in the Tongo region A. tabu. A. polynesiensis prefers resting in dry coconut husks, tree holes, the undersides of partially detached bark on dead trees, and similar sheltered sites. A. upolensis is a true forest dweller, diurnally active and breeds in tree hollows or cavities of dead trees. A. tabu is predominantly found in plantations but also occurs in shady areas in villages, limited to the Tonga islands. Their breeding habitats include tree holes, artificial containers, coconut shells, and leaf axils of taro [35] Other important Aedes vectors are A. poecilus, the A. scutellaris group, and O. togoi (earlier known as A. togoi) [46].

The Anopheles Vector and Its Ecology

Anopheles species are significant vectors of malaria and filariasis in rural regions of Asia and Africa. Some examples include the A. punctulatus group in Papua New Guinea (PNG) [47], A. gambiae s.s., A. arabiensis, and A. funestus on the Kenyan coast [48]; A. subpictus in the Indonesian islands of Flores and Timor [49]; and A. gambiae s.s. in Ghana [50], which transmits both P. falciparum and W. bancrofti parasites.
There are about 26 Anopheles species vectoring Bancroftian and B. filariasis. Of these, eighteen species transmit W. bancrofti, three species transmit B. malayi, and five species transmit both parasites. A. barbirostris is the only known vector of B. timori. Among them, the A. funestus group and members of the A. gambiae complex including A. gambiae s.s., A. arabiensis, A. melas, and A. merus are the most important vectors of W. bancrofti. A. merus breeds in saltwater, while the other three species breed in freshwater [49] with A. melas often associated with mangroves [51].
In Africa, the A. gambiae complex and A. funestus are the most important vectors of W. bancrofti [52]. These vectors exhibit indoor nocturnal biting behaviour. A. gambiae is highly anthropophilic and employs a “patrolling” or “ranging” flight strategy to encounter host cues. A. funestus is highly endophilic and anthropophilic but can display moderate to high zoophagy in areas with large livestock populations. They breed throughout the year, with A. funestus preferring permanent water bodies and certain stagnant water bodies, while the A. gambiae complex breeds in temporary or man-made water bodies like pools, puddles, brick pits, fields, construction sites, hoof prints, or tire tracks. This adaptability allows them to maintain population numbers even during dry months, promoting year-round malaria transmission [53].
In Asia, W. bancrofti is transmitted by A. jeyporiensis candidiensis and A. minimus in China, by A. flavirostris in the Philippines, and by A. balabacensis, A. maculatus, A. letifer, and A. whartoni in Malaysia [49]. The A. punctulatus group includes A. punctulatus, A. farauti, and A. koliensis, which are the principal vectors of the periodic W. bancrofti in Papua New Guinea (PNG), West Papua (Indonesia), Solomon Islands, and Vanuatu [54,55].
The A. punctulatus group prefers to breed in small, shallow, exposed pools devoid of other flora and fauna [56]. In an inland village in PNG, 99.9% of A. punctulatus were recorded. A. farauti is a coastal species that can breed in brackish water but is also found at altitudes over 1000 m above sea level in PNG. A. koliensis is a nocturnal mosquito with a preference for indoor feeding and breeds in streams at the forest margins. In East Sepik Province, PNG, both A. punctulatus and A. koliensis were found to be potential vectors. A. punctulatus breeds along river edges during the rainy season and, during the dry season, along sections of the dried-up river, forming numerous sun-lit puddles that serve as additional breeding sites [57]. Aedes and Anopheles species which are responsible for transmitting W. bancrofti in various regions of the world is listed below (Table 1)

2. Methodology

2.1. Database Search and Systematic Review

We systematically analysed published research articles using specific databases including Google Scholar, ResearchGate, PubMed, and ScienceDirect. Our search focused on topics such as molecular xenomonitoring, W. bancrofti (Wb), Lymphatic Filariasis, Mosquito sampling methods, mosquito trapping techniques, Aedes- and Anopheles-transmitted filariasis, and sub-periodic filariasis. Through this search, we identified 41 relevant articles.
In our analysis, we specifically examined the efficiency of different mosquito trapping techniques, with a focus on sampling Aedes and Anopheles mosquito vectors of W. bancrofti. This information is crucial for conducting molecular xenomonitoring, VBD surveillance, and implementing effective public health programmes. After careful evaluation, we selected 14 records that met our inclusion criteria, making them eligible for data collection and comprehensive analysis.

2.2. Exclusion Criteria

Brugian filariasis studies involved the Anopheles species in transmission. LF was transmitted by Culex mosquitoes.
Studies focused on human blood surveys for microfilarial infection detection for Transmission Assessment Surveys (TAS).

2.3. Inclusion Criteria

  • Aedes and Anopheles transmitted W. bancrofti.
  • Co-endemicity of LF and malaria transmitted by the Anopheles vector.
  • Molecular xenomonitoring of Aedes and Anopheles LF vectors.
  • Mosquito trapping techniques employed in diverse LF-endemic regions for Transmission Assessment Surveys
  • Efficiency assessment of various mosquito traps.
The flow chart below depicts the search strategies for sampling techniques related to Anopheles- and Aedes-mediated W. bancrofti. Full-text records meeting the inclusion criteria were selected for review. The study design was described using all five steps of the PRISMA (Preferred Reporting Items for Systematic Reviews) (Figure 1) checklist to ensure review quality.

3. Sampling Strategies for LF-Endemic Aedes and Anopheles Vectors

The 14 included publications focused on assessing trapping techniques for Aedes and Anopheles vectors transmitting W. bancrofti in endemic regions where an Mx of LF has been conducted. While the HLC method has been essential, ethical concerns and risks to collectors necessitate exploring alternative trapping tools [51].
This review critically analysed the efficiency of seven traps for sampling Anopheles LF vectors such as PSC, GT, BGS, CDC LT, ET, DNT, and AGT (Table 2) and discussed novel collection methods including the Stealth trap, Ifakara Tent Trap, Mbita trap, Furvela tent trap as alternatives to HLC [51]. For Aedes LF vectors, four different traps were analysed (BGS, GT, CDC LT, DNT) along with the Resting Bucket Trap (RB), Sticky Resting Bucket trap (SRB), Duplex cone trap, and novel sticky trap [51].
PSC was the second most efficient method after HLC, effectively capturing Anopheles vectors in Nigeria, Ghana, Togo, and Burkina Faso [7,51,69,79]. However, PSC has drawbacks, as it may miss exophilic mosquitoes or underestimate Anopheles populations [72,78,80,81]. The Exit trap was useful for trapping exophilic mosquitoes [71]. BGS showed varying results in different settings, being more effective in some regions but less so in others [17,34,50]. The AGT appeared to be an appropriate trap for sampling Anopheles populations [51]. CDC LT and DNT also had variable efficiency [71]. In Mali, HLC was observed to be more productive than the PSC [82].
The review identifies traps with potential for xenomonitoring LF vectors, but further studies are needed to assess their effectiveness in different settings [51,73]. The CDC Light Trap captured more Culex and fewer Anopheles vectors [51]. A comparison of various vector sampling techniques in Anopheles- and Aedes-mediated LF-endemic regions is provided in Table 3.

4. Methods for Sampling Mosquitoes to Conduct Disease Surveillance and Research

It is critical to comprehend the behaviour, distribution, and function of mosquitoes as disease vectors in order to effectively combat illnesses such as LF, malaria, dengue, Zika, and others. Scientists are able to advance disease management by gaining insights into mosquito populations and their interactions with the environment through the use of a variety of collection techniques.
Human Landing Catch (HLC): Employing mosquitoes’ biological propensity to seek a blood meal for reproduction, this technique captures mosquitoes as they descend onto human or animal hosts. They are captured using an oral aspirator prior to biting the host [83].
Pyrethrum Spray Collection (PSC): PSC entails the application of an insecticide aerosol containing pyrethrum within confined spaces. Mosquitoes are rendered immobile and disoriented, causing them to descend onto a white cloth, from which they can be collected [84].
CDC Light Trap: These devices attract and capture adult mosquitoes using artificial light sources. By simulating natural light sources, the trap attracts mosquitoes for further study [85].
CDC Gravid Trap: Specialized equipment built to capture female mosquitoes in search of sites where they can lay their eggs. These devices leverage the inherent behaviour of mosquitoes by employing attractants such as organic matter to establish an optimal environment for oviposition [86].
Biogents Sentinel (BGS) Trap: These traps imitate human or animal hosts by combining chemical attractants, visual signals, and heat. The chemical lures and design of the trap increase its attraction to blood-seeking mosquitoes [86].
Window Exit Trap: As mosquitoes enter or leave designated areas, this trapping system captures them. It is comprised of two chambers, one of which is baited with heat and carbon dioxide to simulate human presence and entice mosquitoes to enter; these mosquitoes are then collected in the exit chamber [87].
DNT: This consists of two box nets, one protecting the collector and a second larger net, which is placed directly over the inner net. The outer net is raised off the ground so that mosquitoes attracted to the human bait are collected between the two nets [88].

4.1. Novel Tools for Sampling Anopheles Vectors: W. bancrofti and P. falciparum Transmission

Gravid mosquito traps, such as the OviArt Gravid Trap (AGT), offer valuable sampling tools for both endophilic and exophilic vectors, enhancing the detection of parasite-infected mosquitoes for VBD surveillance and Transmission Assessment Surveys. AGT, specifically targeting gravid Anopheles mosquitoes, has shown promising results with improved catch size compared to other traps like the Box gravid trap. However, further improvements in battery protection and transportation convenience are needed. AGT is made of a rectangular basin measuring 45 cm × 33 cm × 11.5 cm (length × width × height), with a 4 cm hole on the side and a 6 L rectangular basin. An open plastic tube (collection chamber) was inserted into the hole and the other opening of the tube was sealed with fibreglass netting to prevent trapped mosquitoes from escaping. The tube was placed and secured halfway into an aluminium collapsible pipe. The flexible tube was connected to a 12 V fan that provided suction on the water surface [89].
The Stealth trap (ST) has been recommended as a valuable tool for capturing Anopheles mosquitoes, particularly A. gambiae s.l., in West Africa. While ST shows high capture rates, it may cause damage to specimens, making species identification challenging [82]. The Ifakara Tent Trap C design (ITT-C) has been considered promising for outdoor mosquito sampling in Tanzania and has been evaluated for routine malaria vector surveillance. Modifications and validation in different settings are needed for optimal performance [90]. The ITT has been reported to be comparatively superior in terms of capture rates compared to HLC [91] and CDC LT [92]. However, operators’ exposure to mosquito bites necessitates modifications for improved performance [93].
Furvela tent traps provide an efficient way to capture Anopheles mosquitoes, especially in situations with diverse mosquito fauna. Combining CDC-LT or window-exit traps (to sample endophagic mosquitoes) with Furvela tent traps (to sample exophagic ones) allows robust sampling of diverse mosquito species [94]. Both CDC-LT and Furvela tent traps are portable and suitable for surveillance. These tools hold promise for effective vector monitoring in various eco-epidemiological settings [82,90,94,95].

4.2. Aedes Mosquito Sampling Techniques: W. bancrofti Transmission

In a typical endemic setting for DspWb mediated by A. niveus, BGS, DNT, GT, and HLC were deployed for vector mosquito sampling to assess vector infection. However, none of these trapping methods were suitable for sampling A. niveus, with BGS and DNT capturing more A. albopictus and A. aegypti, and GT capturing C. quinquefasciatus [61]. Thus, there is currently no specific sampling tool identified for A. niveus apart from HLC.
In contrast, BGS was found to sample adequate numbers of A. polynesiensis and A. samoanus vectors of LF in Samoa. However, the sampling method suitable for A. polynesiensis in Samoa may not be applicable for A. niveus in Nancowry Islands, and BGS showed limited efficiency in capturing A. niveus [16]. For A. albopictus, some traps like the Duplex cone trap, Resting Bucket Trap (RB), and Sticky Resting Bucket (SRB) have been reported to be efficient in sampling.
The Duplex cone trap was found to be the most productive trap for sampling A. albopictus, offering a promising alternative to HLC [96]. RB and SRB traps were also effective in capturing A. albopictus in various habitats, with SRB showing higher capture rates [97]. Additionally, the novel sticky trap was reported to be more precise than the ovitrap for sampling A. albopictus in urban settings [98].
These findings suggest that, for specific mosquito species like A. niveus, further research and development of the suitable sampling tools are needed. Meanwhile, traps like the Duplex cone trap, RB, SRB, and novel sticky trap show promise for efficient sampling of A. albopictus.

5. Conclusions

In conclusion, this review has focused on the assessment of trapping techniques for Aedes and Anopheles vectors responsible for transmitting W. bancrofti in LF regions endemic to LF. While the Human Landing Collection (HLC) method has been instrumental, it raises ethical concerns and poses risks to collectors, underscoring the need for alternative trapping methods. Among the traps examined, the Pyrethrum Spray Catches (PSC) method demonstrated high efficiency in capturing Anopheles vectors across multiple countries. However, it may not effectively capture exophilic mosquitoes and could underestimate Anopheles populations. Other traps like the Anopheles Gravid Trap (AGT) and the Exit Trap, also showed potential, although their effectiveness varied in different settings.
In the case of Aedes vectors, traps like BGS and CDC LT proved useful in specific regions, yet no dedicated sampling tool was identified for A. niveus apart from HLC. Novel traps like the Stealth trap, the Ifakara Tent Trap C design (ITT-C), and Furvela tent traps offer promising alternatives for capturing Anopheles mosquitoes in diverse eco-epidemiological settings. Regarding A. albopictus, traps like the Duplex cone trap, Resting Bucket Trap (RB), and Sticky Resting Bucket (SRB) exhibited high efficiency in various habitats. Nonetheless, further research is imperative to develop suitable sampling tools for specific mosquito species like A. niveus. In summary, these findings provide valuable insights into efficient sampling strategies for LF-endemic Aedes and Anopheles vectors, thereby facilitating vector surveillance and enhancing disease control efforts.

Author Contributions

Conceptualization—A.N.S.; Writing—(original draft) A.B. and A.N.S.; Mapping—K.H.K.R.; Review and editing—A.N.S., A.B. and A.K. All authors have read and agreed to the published version of the manuscript.


This research did not receive any external funding.

Institutional Review Board Statement

Not applicable.

Informed Consent Statement

Not applicable.

Data Availability Statement

Data is presented in tables within the manuscript.


We extend our heartfelt gratitude to A. Elango, Principal Technical Officer in the Division of Vector Biology and Control, as well as to Divya Teja, Scientist-C (Project) in the same division, for their invaluable technical assistance and support.

Conflicts of Interest

The authors declare no conflict of interest.


  1. World Health Organization. WHO Position Statement on Integrated Vector Management to Control Malaria and Lymphatic Filariasis. Available online: (accessed on 23 November 2022).
  2. WHO. WHO Releases New Guidance on Insecticide-Treated Mosquito Nets. 2007. Available online: (accessed on 23 November 2022).
  3. World Health Organization. WHO Validates 3 More Countries for Eliminating Lymphatic Filariasis. Available online: (accessed on 23 November 2022).
  4. Mishra, K.; Raj, D.K.; Hazra, R.K.; Dash, A.P.; Supakar, P.C. The Development and Evaluation of a Single Step Multiplex PCR Method for Simultaneous Detection of Brugia malayi and Wuchereria bancrofti. Mol. Cell. Probes 2007, 21, 355–362. [Google Scholar] [CrossRef] [PubMed]
  5. Nuchprayoon, S. DNA-Based Diagnosis of Lymphatic Filariasis. Southeast. Asian J. Trop. Med. Public. Health 2009, 40, 904–913. [Google Scholar] [PubMed]
  6. Thanchomnang, T.; Intapan, P.M.; Tantrawatpan, C.; Lulitanond, V.; Chungpivat, S.; Taweethavonsawat, P.; Kaewkong, W.; Sanpool, O.; Janwan, P.; Choochote, W.; et al. Rapid Detection and Identification of Wuchereria bancrofti, Brugia malayi, B. pahangi, and Dirofilaria immitis in Mosquito Vectors and Blood Samples by High-Resolution Melting Real-Time PCR. Korean J. Parasitol. 2013, 51, 645–650. [Google Scholar] [CrossRef] [PubMed]
  7. Owusu, I.O.; de Souza, D.K.; Anto, F.; Wilson, M.D.; Boakye, D.A.; Bockarie, M.J.; Gyapong, J.O. Evaluation of Human and Mosquito Based Diagnostic Tools for Defining Endpoints for Elimination of Anopheles Transmitted Lymphatic Filariasis in Ghana. Trans. R. Soc. Trop. Med. Hyg. 2015, 109, 628–635. [Google Scholar] [CrossRef] [PubMed]
  8. Poole, C.B.; Li, Z.; Alhassan, A.; Guelig, D.; Diesburg, S.; Tanner, N.A.; Zhang, Y.; Jr, T.C.E.; LaBarre, P.; Wanji, S.; et al. Colorimetric Tests for Diagnosis of Filarial Infection and Vector Surveillance Using Non-Instrumented Nucleic Acid Loop-Mediated Isothermal Amplification (NINA-LAMP). PLoS ONE 2017, 12, e0169011. [Google Scholar] [CrossRef] [PubMed]
  9. Fischer, P.; Erickson, S.M.; Fischer, K.; Fuchs, J.F.; Rao, R.U.; Christensen, B.M.; Weil, G.J. Persistence of Brugia Malayi DNA in Vector and Non-Vector Mosquitoes: Implications for Xenomonitoring and Transmission Monitoring of Lymphatic Filariasis. Am. J. Trop. Med. Hyg. 2007, 76, 502–507. [Google Scholar] [CrossRef]
  10. Beng, T.S.; Ahmad, R.; Hisam, R.S.R.; Heng, S.K.; Leaburi, J.; Ismail, Z.; Sulaiman, L.; Soyoti, R.F.H.M.; Hanlim, L. Molecular Xenomonitoring of Filarial Infection in Malaysian Mosquitoes under the National Program for Elimination of Lymphatic Filariasis. Southeast. Asian J. Trop. Med. Public. Health 2016, 47, 617–624. [Google Scholar]
  11. Chu, B.K.; Deming, M.; Biritwum, N.-K.; Bougma, W.R.; Dorkenoo, A.M.; El-Setouhy, M.; Fischer, P.U.; Gass, K.; de Peña, M.G.; Mercado-Hernandez, L.; et al. Transmission Assessment Surveys (TAS) to Define Endpoints for Lymphatic Filariasis Mass Drug Administration: A Multicenter Evaluation. PLOS Negl. Trop. Dis. 2013, 7, e2584. [Google Scholar] [CrossRef]
  12. Schmaedick, M.A.; Koppel, A.L.; Pilotte, N.; Torres, M.; Williams, S.A.; Dobson, S.L.; Lammie, P.J.; Won, K.Y. Molecular Xenomonitoring Using Mosquitoes to Map Lymphatic Filariasis after Mass Drug Administration in American Samoa. PLoS Negl. Trop. Dis. 2014, 8, e3087. [Google Scholar] [CrossRef]
  13. Rao, R.U.; Samarasekera, S.D.; Nagodavithana, K.C.; Punchihewa, M.W.; Dassanayaka, T.D.M.; Gamini, P.K.D.; Ford, E.; Ranasinghe, U.S.B.; Henderson, R.H.; Weil, G.J. Programmatic Use of Molecular Xenomonitoring at the Level of Evaluation Units to Assess Persistence of Lymphatic Filariasis in Sri Lanka. PLOS Negl. Trop. Dis. 2016, 10, e0004722. [Google Scholar] [CrossRef]
  14. Subramanian, S.; Jambulingam, P.; Chu, B.K.; Sadanandane, C.; Vasuki, V.; Srividya, A.; AbdulKader, M.S.M.; Krishnamoorthy, K.; Raju, H.K.; Laney, S.J.; et al. Application of a Household-Based Molecular Xenomonitoring Strategy to Evaluate the Lymphatic Filariasis Elimination Program in Tamil Nadu, India. PLOS Negl. Trop. Dis. 2017, 11, e0005519. [Google Scholar] [CrossRef]
  15. Subramanian, S.; Jambulingam, P.; Krishnamoorthy, K.; Sivagnaname, N.; Sadanandane, C.; Vasuki, V.; Palaniswamy, C.; Vijayakumar, B.; Srividya, A.; Raju, H.K.K. Molecular Xenomonitoring as a Post-MDA Surveillance Tool for Global Programme to Eliminate Lymphatic Filariasis: Field Validation in an Evaluation Unit in India. PLOS Negl. Trop. Dis. 2020, 14, e0007862. [Google Scholar] [CrossRef] [PubMed]
  16. Premkumar, A.; Shriram, A.N.; Krishnamoorthy, K.; Subramanian, S.; Vasuki, V.; Vijayachari, P.; Jambulingam, P. Molecular Xenomonitoring of Diurnally Subperiodic Wuchereria bancrofti Infection in Aedes (Downsiomyia) Niveus (Ludlow, 1903) after Nine Rounds of Mass Drug Administration in Nancowry Islands, Andaman and Nicobar Islands, India. PLOS Negl. Trop. Dis. 2020, 14, e0008763. [Google Scholar] [CrossRef]
  17. McPherson, B.; Mayfield, H.J.; McLure, A.; Gass, K.; Naseri, T.; Thomsen, R.; Williams, S.A.; Pilotte, N.; Kearns, T.; Graves, P.M.; et al. Evaluating Molecular Xenomonitoring as a Tool for Lymphatic Filariasis Surveillance in Samoa, 2018–2019. Trop. Med. Infect. Dis. 2022, 7, 203. [Google Scholar] [CrossRef] [PubMed]
  18. Samarawickrema, W.A.; Kimura, E.; Spears, G.F.; Penaia, L.; Sone, F.; Paulson, G.S.; Cummings, R.F. Distribution of Vectors, Transmission Indices and Microfilaria Rates of Subperiodic Wuchereria bancrofti in Relation to Village Ecotypes in Samoa. Trans. R. Soc. Trop. Med. Hyg. 1987, 81, 129–135. [Google Scholar] [CrossRef]
  19. Ottesen, E.A.; Duke, B.O.; Karam, M.; Behbehani, K. Strategies and Tools for the Control/Elimination of Lymphatic Filariasis. Bull. World Health Organ. 1997, 75, 491–503. [Google Scholar]
  20. Wilson, J. A Missionary Voyage to the Southern Pacific Ocean, Performed in the Years 1796, 1797, 1798, in the Ship Duff, Commanded by Captain James Wilson. Compiled from Journals of the Officers and the Missionaries. Available online: (accessed on 25 November 2022).
  21. Koniger Beobachtungen Über Elephantiasis Auf Samoa. Von Dr. Königer, Marine Assistenzarzt 1. Klasse (Pp.413–422, 3 Holzschnitte, 1 Lith. Taf. 4 Abb.). by Königer: (1879)|Antiq. F.-D. Söhn—Medicusbooks.Com. Available online: (accessed on 25 November 2022).
  22. Symes, C.B. Observations on the Epidemiology of Filariasis in Fiji. J. Trop. Med. Hyg. 1960, 63, 1–14. [Google Scholar]
  23. Kalra, N.L. Filariasis among Aborigines of Andaman and Nicobar Islands. 1. Detection of Non-Periodic Bancroftian Filariasis among Nicobarese of Nanncowry Group of Nicobar Islands. J. Commun. Dis. 1974, 6, 40–56. [Google Scholar]
  24. Russel, S.; Das, M.; Rao, C.K. Filariasis in Andaman and Nicobar Islands. Part I. Survey Findings-Nancowry, Terressa, Chowra, Carnicobar and Port Blair. J. Commun. Dis. 1975, 7, 15–30. [Google Scholar]
  25. Basu, P.C. A note on malaria and filariasis in Andaman and Nicobar. Bull. Natl. Soc. India Malar. Other Mosq. Borne Dis. 1958, 6, 193–206. [Google Scholar]
  26. Das, M.; Russel, S.; Rao, C.K. Filariasis in Andaman and Nicobar Islands. Part II. Periodicity of Microfilaria of Wuchereria bancrofti. J. Commun. Dis. 1975, 7, 251–256. [Google Scholar]
  27. Tewari, S.C.; Hiriyan, J. Description of Aedes (Finlaya) Niveus (Diptera: Culicidae) from Andaman and Nicobar, India. Mosq. Syst. 1995, 27, 167–176. [Google Scholar]
  28. Shriram, A.N.; Murhekar, M.V.; Ramaiah, K.D.; Sehgal, S.C. Prevalence of Diurnally Subperiodic Bancroftian Filariasis among the Nicobarese in Andaman and Nicobar Islands, India: Effect of Age and Gender. Trop. Med. Int. Health 2002, 7, 949–954. [Google Scholar] [CrossRef] [PubMed]
  29. Sasa, M. Human Filariasis: A Global Survey of Epidemiology and Control; University Park Press: University Park, PN, USA, 1976. [Google Scholar]
  30. Harinasuta, C.; Sucharit, S.; Deesin, T.; Surathin, K.; Vutikes, S. Bancroftian Filariasis in Thailand, a New Endemic Area. Southeast. Asian J. Trop. Med. Public. Health 1970, 1, 233–245. [Google Scholar]
  31. Khan, A.M.; Dutta, P.; Das, S.; Pathak, A.K.; Sarmah, P.; Hussain, M.E.; Mahanta, J. Microfilarial Periodicity of Wuchereria Bancrofti in Assam, Northeast India. J. Vector Borne Dis. 2015, 52, 208–212. [Google Scholar] [PubMed]
  32. Ramalingam, S.; Belkin, J.N. Vectors of Sub-Periodic Bancroftian Filariasis in the Samoa–Tonga Area. Nature 1964, 201, 105–106. [Google Scholar] [CrossRef]
  33. Ichimori, K. Entomology of the Filariasis Control Programme in Samoa, Aedes Polynesiensis and Ae. Samoanus. Med. Entomol. Zool. 2001, 52, 11–21. [Google Scholar] [CrossRef]
  34. Hapairai, L.K.; Plichart, C.; Naseri, T.; Silva, U.; Tesimale, L.; Pemita, P.; Bossin, H.C.; Burkot, T.R.; Ritchie, S.A.; Graves, P.M.; et al. Evaluation of Traps and Lures for Mosquito Vectors and Xenomonitoring of Wuchereria bancrofti Infection in a High Prevalence Samoan Village. Parasites Vectors 2015, 8, 1–9. [Google Scholar] [CrossRef]
  35. Ramalingam, S. The Epidemiology of Filarial Transmission in Samoa and Tonga. Ann. Trop. Med. Parasitol. 1968, 62, 305–324. [Google Scholar] [CrossRef]
  36. Suzuki, T.; Sone, F. The Bionomics of Filariasis Vectors in Western Samoa. Jpn. J. Sanit. Zool. 1974, 25, 251–257. [Google Scholar] [CrossRef]
  37. Suzuki, T.; Sone, F. Breeding Habits of Vector Mosquitoes of Filariasis and Dengue Fever in Western Samoa. Eisei dobutsu. Jpn. J. Sanit. Zool. 1978, 29, 279–286. [Google Scholar] [CrossRef]
  38. Bahr, P.H. Filariasis and Elephantiasis in Fiji. J. Am. Med. Assoc. 1912, LVIII, 2055. [Google Scholar] [CrossRef]
  39. Marks, E.N. The Vector of Filariasis in Polynesia: A Change in Nomenclature. Ann. Trop. Med. Parasitol. 1951, 45, 137–140. [Google Scholar] [CrossRef] [PubMed]
  40. Symes, C.B. Filarial Infections in Mosquitoes in Fiji. Trans. R. Soc. Trop. Med. Hyg. 1955, 49, 280–284. [Google Scholar] [CrossRef] [PubMed]
  41. Burnett, G.F. Filariasis Research in Fiji 1957-1959. Part 1. Epidemiology. J. Trop. Med. Hyg. 1960, 63, 153–162. [Google Scholar]
  42. Rosen, L. Observations on the Epidemiology of Human Filariasis in French Oceania. Am. J. Hyg. 1955, 61, 219–248. [Google Scholar]
  43. Belkin, J.N. The Mosquitoes of the South Pacific (Diptera, Culicidae); Cambridge University Press: London, UK, 1962; Volume 2. [Google Scholar]
  44. Jachowski, L.A. Filariasis in American Samoa. V. Bionomics of the Principal Vector, Aedes Polynesiensis Marks. Am. J. Hyg. 1954, 60, 186–203. [Google Scholar]
  45. Shriram, A.N.; Ramaiah, K.D.; Krishnamoorthy, K.; Sehgal, S.C. Diurnal Pattern of Human-Biting Activity and Transmission of Subperiodic Wuchereria bancrofti (Filariidea: Dipetalonematidae) by Ochlerotatus niveus (Diptera: Culicidae) on the Andaman and Nicobar Islands of India. Am. J. Trop. Med. Hyg. 2005, 72, 273–277. [Google Scholar] [CrossRef]
  46. Bockarie, M.J.; Pedersen, E.M.; White, G.B.; Michael, E. Role of Vector Control in the Global Program to Eliminate Lymphatic Filariasis. Annu. Rev. Entomol. 2009, 54, 469–487. [Google Scholar] [CrossRef]
  47. Bockarie, M.J.; Tavul, L.; Kastens, W.; Michael, E.; Kazura, J.W. Impact of Untreated Bednets on Prevalence of Wuchereria Bancrofti Transmitted by Anopheles Farauti in Papua New Guinea. Med. Vet. Entomol. 2002, 16, 116–119. [Google Scholar] [CrossRef]
  48. Muturi, E.J.; Mbogo, C.M.; Ng’ang’a, Z.W.; Kabiru, E.W.; Mwandawiro, C.; Novak, R.J.; Beier, J.C. Relationship between Malaria and Filariasis Transmission Indices in an Endemic Area along the Kenyan Coast. J. Vector Borne Dis. 2006, 43, 77–83. [Google Scholar] [PubMed]
  49. World Health Organization. Geographical Distribution of Arthropod-Borne Diseases and Their Principal Vectors. Available online: (accessed on 1 December 2022).
  50. De Souza, D.; Kelly-Hope, L.; Lawson, B.; Wilson, M.; Boakye, D. Environmental Factors Associated with the Distribution of Anopheles Gambiae s.s in Ghana; an Important Vector of Lymphatic Filariasis and Malaria. PLoS ONE 2010, 5, e9927. [Google Scholar] [CrossRef] [PubMed]
  51. Opoku, M.; Minetti, C.; Kartey-Attipoe, W.D.; Otoo, S.; Otchere, J.; Gomes, B.; de Souza, D.K.; Reimer, L.J. An Assessment of Mosquito Collection Techniques for Xenomonitoring of Anopheline-Transmitted Lymphatic Filariasis in Ghana. Parasitology 2018, 145, 1783–1791. [Google Scholar] [CrossRef]
  52. De Souza, D.K.; Koudou, B.; Kelly-Hope, L.A.; Wilson, M.D.; Bockarie, M.J.; Boakye, D.A. Diversity and Transmission Competence in Lymphatic Filariasis Vectors in West Africa, and the Implications for Accelerated Elimination of Anopheles-Transmitted Filariasis. Parasites Vectors 2012, 5, 259. [Google Scholar] [CrossRef]
  53. Kahamba, N.F.; Finda, M.; Ngowo, H.S.; Msugupakulya, B.J.; Baldini, F.; Koekemoer, L.L.; Ferguson, H.M.; Okumu, F.O. Using Ecological Observations to Improve Malaria Control in Areas Where Anopheles Funestus Is the Dominant Vector. Malar. J. 2022, 21, 158. [Google Scholar] [CrossRef]
  54. Webber, R.H. The Natural Decline of Wuchereria Bancrofti Infection in a Vector Control Situation in the Solomon Islands. Trans. R. Soc. Trop. Med. Hyg. 1977, 71, 396–400. [Google Scholar] [CrossRef] [PubMed]
  55. Webber, R.H. Eradication of Wuchereria Bancrofti Infection through Vector Control. Trans. R. Soc. Trop. Med. Hyg. 1979, 73, 722–724. [Google Scholar] [CrossRef]
  56. Beebe, N.W.; Russell, T.; Burkot, T.R.; Cooper, R.D. Anopheles Punctulatus Group: Evolution, Distribution, and Control. Annu. Rev. Entomol. 2015, 60, 335–350. [Google Scholar] [CrossRef]
  57. Bockarie, M.; Kazura, J.; Alexander, N.; Dagoro, H.; Bockarie, F.; Perry, R.; Alpers, M. Transmission Dynamics of Wuchereria Bancrofti in East Sepik Province, Papua New Guinea. Am. J. Trop. Med. Hyg. 1996, 54, 577–581. [Google Scholar] [CrossRef]
  58. Bartilol, B.; Omedo, I.; Mbogo, C.; Mwangangi, J.; Rono, M.K. Bionomics and Ecology of Anopheles Merus along the East and Southern Africa Coast. Parasites Vectors 2021, 14, 84. [Google Scholar] [CrossRef]
  59. Strickland, G.T. Hunter’s Tropical Medicine and Emerging Infectious Diseases. Rev. Inst. Med. Trop. 2001, 43, 112. [Google Scholar] [CrossRef]
  60. Jitpakdi, A.; Choochote, W.; Panart, P.; Tookyang, B.; Panart, K.; Prajakwong, S. Possible Transmission of Two Types of Wuchereria bancrofti in Muang District, Chiang Mai, Northern Thailand. Southeast. Asian J. Trop. Med. Public Health 1998, 29, 141–143. [Google Scholar] [PubMed]
  61. Gould, D.J.; Bailey, C.L.; Vongpradist, S. Implication of Forest Mosquitoes in the Transmission of Wuchereria bancrofti in Thailand. Mosq. News 1982, 42, 560–564. [Google Scholar]
  62. Muturi, E.J.; Jacob, B.G.; Kim, C.-H.; Mbogo, C.M.; Novak, R.J. Are Coinfections of Malaria and Filariasis of Any Epidemiological Significance? Parasitol. Res. 2008, 102, 175–181. [Google Scholar] [CrossRef] [PubMed]
  63. Hii, J.L.; Kan, S.; Vun, Y.S.; Chin, K.F.; Lye, M.S.; Mak, J.W.; Cheong, W.H. Anopheles Flavirostris Incriminated as a Vector of Malaria and Bancroftian Filariasis in Banggi Island, Sabah, Malaysia. Trans. R. Soc. Trop. Med. Hyg. 1985, 79, 677–680. [Google Scholar] [CrossRef]
  64. Chen, B.; Harbach, R.E.; Butlin, R.K. Molecular and Morphological Studies on the Anopheles Minimus Group of Mosquitoes in Southern China: Taxonomic Review, Distribution and Malaria Vector Status. Med. Vet. Entomol. 2002, 16, 253–265. [Google Scholar] [CrossRef]
  65. Manguin, S.; Bangs, M.J.; Pothikasikorn, J.; Chareonviriyaphap, T. Review on Global Co-Transmission of Human Plasmodium Species and Wuchereria Bancrofti by Anopheles Mosquitoes. Infect. Genet. Evol. 2010, 10, 159–177. [Google Scholar] [CrossRef]
  66. Sallum, M.; Peyton, E.L.; Wilkerson, R. Six New Species of the Anopheles Leucosphyrus Group, Reinterpretation of An. Elegans and Vector Implications. Med. Vet. Entomol. 2005, 19, 158–199. [Google Scholar] [CrossRef] [PubMed]
  67. White, G.B. Anopheles Bwambae Sp.n., a Malaria Vector in the Semliki Valley, Uganda, and Its Relationships with Other Sibling Species of the An.Gambiae Complex (Diptera: Culicidae). Syst. Entomol. 1985, 10, 501–522. [Google Scholar] [CrossRef]
  68. Abeyewickreme, W.; Wanniarachchi, P. Anopheles (Cellia) Jamesii: A Potential Natural Vector of Bancroftian Filariasis in Sri Lanka. Trans. R. Soc. Trop. Med. Hyg. 1991, 85, 644. [Google Scholar] [CrossRef]
  69. Richards, F.O.; Eigege, A.; Miri, E.S.; Kal, A.; Umaru, J.; Pam, D.; Rakers, L.J.; Sambo, Y.; Danboyi, J.; Ibrahim, B.; et al. Epidemiological and Entomological Evaluations after Six Years or More of Mass Drug Administration for Lymphatic Filariasis Elimination in Nigeria. PLOS Negl. Trop. Dis. 2011, 5, e1346. [Google Scholar] [CrossRef]
  70. Dorkenoo, M.A.; de Souza, D.K.; Apetogbo, Y.; Oboussoumi, K.; Yehadji, D.; Tchalim, M.; Etassoli, S.; Koudou, B.; Ketoh, G.K.; Sodahlon, Y.; et al. Molecular Xenomonitoring for Post-Validation Surveillance of Lymphatic Filariasis in Togo: No Evidence for Active Transmission. Parasites Vectors 2018, 11, 52. [Google Scholar] [CrossRef] [PubMed]
  71. Coulibaly, S.; Sawadogo, S.P.; Nikièma, A.S.; Hien, A.S.; Bamogo, R.; Koala, L.; Sangaré, I.; Bougma, R.W.; Koudou, B.; Fournet, F.; et al. Assessment of Culicidae Collection Methods for Xenomonitoring Lymphatic Filariasis in Malaria Co-Infection Context in Burkina Faso. bioRxiv 2022. bioRxiv 2022.04.26.489492. [Google Scholar]
  72. Coulibaly, Y.I.; Dembele, B.; Diallo, A.A.; Konaté, S.; Dolo, H.; Coulibaly, S.Y.; Doumbia, S.S.; Soumaoro, L.; Coulibaly, M.E.; Bockarie, M.J.; et al. The Impact of Six Annual Rounds of Mass Drug Administration on Wuchereria bancrofti Infections in Humans and in Mosquitoes in Mali. Am. J. Trop. Med. Hyg. 2015, 93, 356–360. [Google Scholar] [CrossRef]
  73. Jones, C.; Ngasala, B.; Derua, Y.; Tarimo, D.; Reimer, L.; Bockarie, M.; Malecela, M. Lymphatic Filariasis Transmission in Rufiji District, Southeastern Tanzania: Infection Status of the Human Population and Mosquito Vectors after Twelve Rounds of Mass Drug Administration. Parasites Vectors 2018, 11, 588. [Google Scholar] [CrossRef] [PubMed]
  74. Mathenge, E.M.; Misiani, G.O.; Oulo, D.O.; Irungu, L.W.; Ndegwa, P.N.; Smith, T.A.; Killeen, G.F.; Knols, B.G. Comparative Performance of the Mbita Trap, CDC Light Trap and the Human Landing Catch in the Sampling of Anopheles Arabiensis, An. Funestus and Culicine Species in a Rice Irrigation in Western Kenya. Malar. J. 2005, 4, 7. [Google Scholar] [CrossRef]
  75. Atmosoedjono, S.; Dennis, D.T. Anopheles aconitus and An. subpictus naturally infected with Wuchereria bancrofti in Flores, Indonesia. Mosq. News 1977, 37, 529. [Google Scholar]
  76. Reimer, L.J.; Thomsen, E.K.; Tisch, D.J.; Henry-Halldin, C.N.; Zimmerman, P.A.; Baea, M.E.; Dagoro, H.; Susapu, M.; Hetzel, M.W.; Bockarie, M.J.; et al. Insecticidal Bed Nets and Filariasis Transmission in Papua New Guinea. N. Engl. J. Med. 2013, 369, 745–753. [Google Scholar] [CrossRef]
  77. Coulibaly, Y.I.; Coulibaly, S.Y.; Dolo, H.; Konate, S.; Diallo, A.A.; Doumbia, S.S.; Soumaoro, L.; Coulibaly, M.E.; Dicko, I.; Sangare, M.B.; et al. Dynamics of Antigenemia and Transmission Intensity of Wuchereria bancrofti Following Cessation of Mass Drug Administration in a Formerly Highly Endemic Region of Mali. Parasit. Vectors 2016, 9, 628. [Google Scholar] [CrossRef]
  78. Coulibaly, S.; Sawadogo, S.P.; Hien, A.S.; Nikièma, A.S.; Sangaré, I.; Rabila, B.; Koala, L.; Bougouma, C.; Bougma, R.W.; Ouedraogo, G.A.; et al. Malaria and Lymphatic Filariasis Co-Transmission in Endemic Health Districts in Burkina Faso. Adv. Entomol. 2021, 9, 155–175. [Google Scholar] [CrossRef]
  79. Dorkenoo, M.A.; Bronzan, R.; Yehadji, D.; Tchalim, M.; Yakpa, K.; Etassoli, S.; Adjeloh, P.; Maman, I.; Sodahlon, Y. Surveillance for Lymphatic Filariasis after Stopping Mass Drug Administration in Endemic Districts of Togo, 2010–2015. Parasit. Vectors 2018, 11, 244. [Google Scholar] [CrossRef]
  80. Schmaedick, M.A.; Ball, T.S.; Burkot, T.R.; Gurr, N.E. Evaluation of Three Traps for Sampling Aedes Polynesiensis and Other Mosquito Species in American Samoa. J. Am. Mosq. Control Assoc. 2008, 24, 319–322. [Google Scholar] [CrossRef] [PubMed]
  81. Ndiath, M.O.; Mazenot, C.; Gaye, A.; Konate, L.; Bouganali, C.; Faye, O.; Sokhna, C.; Trape, J.-F. Methods to Collect Anopheles Mosquitoes and Evaluate Malaria Transmission: A Comparative Study in Two Villages in Senegal. Malar. J. 2011, 10, 270. [Google Scholar] [CrossRef]
  82. Cansado-Utrilla, C.; Jeffries, C.L.; Kristan, M.; Brugman, V.A.; Heard, P.; Camara, G.; Sylla, M.; Beavogui, A.H.; Messenger, L.A.; Irish, S.R.; et al. An Assessment of Adult Mosquito Collection Techniques for Studying Species Abundance and Diversity in Maferinyah, Guinea. Parasit. Vectors 2020, 13, 150. [Google Scholar] [CrossRef] [PubMed]
  83. Gimnig, J.E.; Walker, E.D.; Otieno, P.; Kosgei, J.; Olang, G.; Ombok, M.; Williamson, J.; Marwanga, D.; Abong’o, D.; Desai, M.; et al. Incidence of Malaria among Mosquito Collectors Conducting Human Landing Catches in Western Kenya. Am. J. Trop. Med. Hyg. 2013, 88, 301–308. [Google Scholar] [CrossRef]
  84. Russell, T.L.; Staunton, K.; Burkot, T. Standard Operating Procedure for Collecting Resting Mosquitoes with Pyrethrum Spray Catch. 2022. Available online: (accessed on 23 November 2022).
  85. Sriwichai, P.; Karl, S.; Samung, Y.; Sumruayphol, S.; Kiattibutr, K.; Payakkapol, A.; Mueller, I.; Yan, G.; Cui, L.; Sattabongkot, J. Evaluation of CDC Light Traps for Mosquito Surveillance in a Malaria Endemic Area on the Thai-Myanmar Border. Parasites Vectors 2015, 8, 636. [Google Scholar] [CrossRef]
  86. Cilek, J.E.; Knapp, J.A.; Richardson, A.G. Comparative Efficiency of Biogents Gravid Aedes Trap, Cdc Autocidal Gravid Ovitrap, and CDC Gravid Trap in Northeastern Florida. J. Am. Mosq. Control. Assoc. 2017, 33, 103–107. [Google Scholar] [CrossRef] [PubMed]
  87. Githinji, E.K.; Irungu, L.W.; Ndegwa, P.N.; Machani, M.G.; Amito, R.O.; Kemei, B.J.; Murima, P.N.; Ombui, G.M.; Wanjoya, A.K.; Mbogo, C.M.; et al. Impact of Insecticide Resistance on P. Falciparum Vectors’ Biting, Feeding, and Resting Behaviour in Selected Clusters in Teso North and South Subcounties in Busia County, Western Kenya. J. Parasitol. Res. 2020, 2020, 9423682. [Google Scholar] [CrossRef]
  88. Tangena, J.-A.A.; Thammavong, P.; Hiscox, A.; Lindsay, S.W.; Brey, P.T. The Human-Baited Double Net Trap: An Alternative to Human Landing Catches for Collecting Outdoor Biting Mosquitoes in Lao PDR. PLoS ONE 2015, 10, e0138735. [Google Scholar] [CrossRef]
  89. Dugassa, S.; Lindh, J.M.; Oyieke, F.; Mukabana, W.R.; Lindsay, S.W.; Fillinger, U. Development of a Gravid Trap for Collecting Live Malaria Vectors Anopheles Gambiae s.l. PLoS ONE 2013, 8, e68948. [Google Scholar] [CrossRef]
  90. Opondo, K. Efficacy of the D-Design Ifakara Tent Trap for Sampling Malaria Vectors in An Area of Mass Long Lasting Insecticidal Bed Nets Use. Ph.D. Thesis, University of Nairobi, Nairobi, Kenya, 2012. [Google Scholar]
  91. Sikulu, M.; Govella, N.J.; Ogoma, S.B.; Mpangile, J.; Kambi, S.H.; Kannady, K.; Chaki, P.C.; Mukabana, W.R.; Killeen, G.F. Comparative Evaluation of the Ifakara Tent Trap-B, the Standardized Resting Boxes and the Human Landing Catch for Sampling Malaria Vectors and Other Mosquitoes in Urban Dar Es Salaam, Tanzania. Malar. J. 2009, 8, 197. [Google Scholar] [CrossRef] [PubMed]
  92. Govella, N.J.; Chaki, P.P.; Mpangile, J.M.; Killeen, G.F. Monitoring Mosquitoes in Urban Dar Es Salaam: Evaluation of Resting Boxes, Window Exit Traps, CDC Light Traps, Ifakara Tent Traps and Human Landing Catches. Parasites Vectors 2011, 4, 40. [Google Scholar] [CrossRef] [PubMed]
  93. Govella, N.J.; Chaki, P.P.; Geissbuhler, Y.; Kannady, K.; Okumu, F.; Charlwood, J.D.; Anderson, R.A.; Killeen, G.F. A New Tent Trap for Sampling Exophagic and Endophagic Members of the Anopheles Gambiae Complex. Malar. J. 2009, 8, 157. [Google Scholar] [CrossRef] [PubMed]
  94. Charlwood, J.D.; Rowland, M.; Protopopoff, N.; Le Clair, C. The Furvela Tent-Trap Mk 1.1 for the Collection of Outdoor Biting Mosquitoes. PeerJ. 2017, 5, e3848. [Google Scholar] [CrossRef]
  95. Dugassa, S.; Lindh, J.M.; Lindsay, S.W.; Fillinger, U. Field Evaluation of Two Novel Sampling Devices for Collecting Wild Oviposition Site Seeking Malaria Vector Mosquitoes: OviART Gravid Traps and Squares of Electrocuting Nets. Parasit. Vectors 2016, 9, 272. [Google Scholar] [CrossRef]
  96. Freier, J.E.; Francy, D.B. A Duplex Cone Trap for the Collection of Adult Aedes Albopictus. J. Am. Mosq. Control Assoc. 1991, 7, 73–79. [Google Scholar]
  97. Brown, R.; Hing, C.T.; Fornace, K.; Ferguson, H.M. Evaluation of Resting Traps to Examine the Behaviour and Ecology of Mosquito Vectors in an Area of Rapidly Changing Land Use in Sabah, Malaysian Borneo. Parasites Vectors 2018, 11, 346. [Google Scholar] [CrossRef]
  98. Facchinelli, L.; Valerio, L.; Pombi, M.; Reiter, P.; Costantini, C.; Della Torre, A. Development of a Novel Sticky Trap for Container-Breeding Mosquitoes and Evaluation of Its Sampling Properties to Monitor Urban Populations of Aedes albopictus. Med. Vet. Entomol. 2007, 21, 183–195. [Google Scholar] [CrossRef]
Figure 1. Identification of Sampling techniques for Anopheles- and Aedes-mediated W. bancrofti through databases. PRISMA flow diagram of study selection and inclusion.
Figure 1. Identification of Sampling techniques for Anopheles- and Aedes-mediated W. bancrofti through databases. PRISMA flow diagram of study selection and inclusion.
Pathogens 12 01406 g001
Table 1. Aedes and Anopheles LF vectors.
Table 1. Aedes and Anopheles LF vectors.
SL.NO.RegionVector SpeciesReference
1Flores and Timor (Indonesian Islands)An. subpictusWHO-2022 [3]
2ChinaAn. jeyporiensis candidiensis
3An. minimus
4PhilippinesAn. flavirostrisWHO-2022 [3]
5GhanaAn. gambiae complex,Owusu et al., 2015 [7]
6An. funestus,
7An. arabiensis
8An. melas
9American SamoaAe. polynesiensisSchmaedick et al., 2014 [12]
10Ae. samoanus
11Ae. aegypti,
12Ae. (Finlaya) group
13India (A &N Islands)Ae. (Downsiomyia) niveusPremkumar et al., 2020 [16]
14ThailandAe. niveus group Harinasuta et al., 1970 [30]
15SamoaAe. polynesiensisHapairai et al., 2015 [34]
16Ae. samoanus
17Ae. (Finlaya) spp.
18Ae. aegypti
19Ae. upolensisRamalingam et al., 1968 [35]
20Polynesian regionAe. kochi group Burnett et al., 1960 [41]
21PhilippinesAe. poecilusBockarie et al., 2009 [46]
22Papua New Guinea, West Papua (Indonesia), Solomon isalnds, VanautuAn. punctulatusWebber et al., 1977, 1979, 1991 [54]
23An. farauti
24An. koliensis
25TanzaniaAn. merusBartilol et al., 2021 [58]
26Polynesia, New CaledoniaAe. polynesiensisStrickland Hunter’s Tropical Medicine and emerging Infectious diseases [59]
27Ae. tabu
28Ae. vigilax
29ThailandAe. annandaleiJitpakdi et al., 1998 [60]
30Ae. desmotesGould et al., 1982 [61]
31Ae. harinasutai
32MalaysiaAn. leucosphyrusMuturi et al., 2008 [62]
33An. barbirostris
34An. balabacensis
35An. maculatus
36An. letifer
37An. whartoni
38An. donaldi
39An. campestris
40IndonesiaAn. balabacensis
42Solomons islandAn. koriensis
43China and KoreaAn. sinensis
44PhilippinesAn. minimus
45Banggi Island, Sabah, MalaysiaAn. flavirostrisHii et al., 1975 [63]
46Hainan island, ChinaAn. minimusChen et al., 2002 [64]
47Papua New GuineaAn. koliensisManguin et al., 2010 [65]
48An. bancroftii
49An. farauti s.l.
50An. punctulatus
51Brazil, Dominican republic, Guyana, Haiti, Costa Rica, Suriname, Trinidad, Tobago and BrazilAn. darlingi
52An. aquasalis
53An. albimanus
54An. bellator
55BorneoAn. balabacensisSallum et al., 2005 [66]
56An. latens
57UgandaAn. bwambaeGB White et al., 1985 [67]
58Sri LankaAn. (Cellia) jamesiiAbeyewickreme et al., 1991 [68]
59NigeriaAn. gambiae s.l.Richards et al., 2011 [69]
60An. funestus
61TogoAn. gambiaeDorkinoo et al., 2018 [70]
62Burkina Faso (West africa)An. gambiae s.l.Sanata Coulibaly et al., 2022 [71]
63An. funestus s.l.,
64An. coluzzii
65An. gambiae
66An. nili
67MaliAn. gambiae complex,Coulibaly et al., 2015 [72]
68An. funestus complex
69TanzaniaAn. gambiae complex Jones et al., 2018 [73]
70Kenya CoastAn. gambiae s.s.Mathenge et al., 2005 [74]
71An. arabiensis
72An. funestus
73IndonesiaAn. aconitusAtmosoedjono et al., 1977 [75]
Table 2. Studies carried out in different LF-endemic regions where the principal vectors are Aedes and Anopheles.
Table 2. Studies carried out in different LF-endemic regions where the principal vectors are Aedes and Anopheles.
S. No.Filariasis Endemic CountriesMain VectorContextStudy DateStudy DesignVector Sampling MethodsSample SizeAnalysis MethodReference
1NigeriaA. gambiae s.l.,
A. funestus,
Anopheles spp.,
Culex spp.
Post-MDA surveillance2009LongitudinalPSC4398DissectionRichards et al., 2011 [69]
2Papua New Guinea A. punctulatus
A. koliensis *
A. hinesorum *
A. farauti 4 *
A. farauti sensu stricto.*
Post-MDA surveillance2007–2008LongitudinalHLC21,899PCRReimer et al., 2013 [76]
A. polynesiensis
A. samoanus
A. aegypti
A. (Finlaya) group
A. oceanicus *
A. samoanus *
A. tutuilae *
A. nocturnus *
C. annulirostris *
C. sitiens *
C. quinquefasciatus
Post-MDA surveillance2011Cross-sectional BGS + lure22,014PCRSchmaedick et al., 2014 [12]
4MaliA. gambiae complex,
A. funestus complex
Post-MDA surveillance2007LongitudinalHLC4680DissectionCoulibaly et al., 2015 [72]
5GhanaAnopheles spp.
Culex spp.
Post-MDA surveillance2008Cross-sectionalPSC and GT4500PCROwusu et al., 2015 [7]
6SamoaA. polynesiensis
A. (Finlaya) sp.
A. aegypti *
A. upolensis *
C. annulirostris *
C. quinquefasciatus *
Post-MDA surveillance2012Cross-sectional BGS + Lure, HBC and CDC LT 5360PCRHapairai et al., 2015 [34]
7MaliA. gambiae complex
A. funestus complex
A. pharaoensis *
A. rufipes *
Post-MDA surveillance2009–2013LongitudinalHLC and PSC14,539Dissection and PCRCoulibaly et al., 2016 [77]
8TogoA. gambiae,
Culex spp.,
A. aegypti *,
Mansonia spp. *
Post-MDA surveillance2015Cross-sectional PSC, HLC, and ET 10,872PCRDorkinoo et al., 2018 [70]
9TanzaniaA. gambiae complex,
C. quinquefasciatus
Post-MDA surveillance2015Cross-sectional CDC LT and GT 1650Dissection and PCRJones et al., 2018 [73]
10GhanaA. gambiae complex,
A. funestus,
A. arabiensis
A. melas,
A. rufipes *
A. coustani *
Aedes spp *
Culex spp.*
Mansonia spp.*
Post-MDA surveillance2016–2017Cross-sectional AGT, Box GT, CDC GT, LT, ET, BGS, IRC, PSC2188PCROpoku et al., 2018 [51]
11India (A and N Islands)A. (Downsiomyia) niveus,
C. quinquefasciatus
A. albopictus *
A. aegypti *
A. edwardsi *
A. malayensis *
A. subalbatus *
Post-MDA surveillance2014–2015Cross-sectional BGS, GT, DNT, and HLC2170RT-PCRPremkumar et al., 2020 [16]
12Burkina Faso (West Africa)A. gambiae s.l.,
A. funestus s.l.,
A. coluzzii,
A. gambiae,
A. nili,
A. arabiensis *
Monitoring of LF and malaria prevalence2014 and 2015Cross-sectionalHLC, PSC29,183Conventional PCR and LAMPSanata Coulibaly et al., 2021 [78]
13SamoaA. polynesiensis,
A. samoanus,
A. (finlaya) spp.
A. aegypti,
A. albopictus *
A. upolensis *
C. quinquefasciatus *
Post-MDA Surveillance2018 and 2019LongitudinalBGS + Lure13,700PCRMcPherson et al., 2022 [17]
14Burkina FasoA. gambiae,
A. coluzzi *
A. arabiensis *
A. funestus group,
A. nili
Assessment of VSM2018Cross-SectionalCross-sectionalHLC, Window ET, DNT, PSC3322PCRSanata Coulibaly et al., 2022 [71]
* Non—LF vectors collected during sampling. HLC—Human Landing Catch, PSC—Pyrethrum Spray Catch, HBC—Human Bait Catch, BGS—Biogents Sentinel trap, CDC LT—Centres for Disease Control Light trap, ET—Exit Trap, GT—Gravid Trap, AGT—Anopheles gravid trap, DNT—Human baited double bed net-traps, BOX—box gravid trap, IRC—Indoor Resting Collection.
Table 3. Applicability of different vector sampling techniques to sample Anopheles and Aedes vectors in LF-endemic regions apart from the Human Landing Collection.
Table 3. Applicability of different vector sampling techniques to sample Anopheles and Aedes vectors in LF-endemic regions apart from the Human Landing Collection.
Nigeria Anopheles sp.X-----
Ghana XXX-
Togo -----
Burkina Faso ----
Mali X------
SamoaAedes sp.-----
Nancowry Islands, India -XX--X-
PSC—Pyrethrum Spray Catch, HBC—Human Bait Catch, GT—Gravid Trap, BGS—Biogents Sentinel trap, CDC LT—Centres for Disease Control Light trap, ET—Exit Trap, DNT—Human baited Double Net Traps, AGT—Anopheles gravid trap. : effective; X: invalid.
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Bhuvaneswari, A.; Shriram, A.N.; Raju, K.H.K.; Kumar, A. Mosquitoes, Lymphatic Filariasis, and Public Health: A Systematic Review of Anopheles and Aedes Surveillance Strategies. Pathogens 2023, 12, 1406.

AMA Style

Bhuvaneswari A, Shriram AN, Raju KHK, Kumar A. Mosquitoes, Lymphatic Filariasis, and Public Health: A Systematic Review of Anopheles and Aedes Surveillance Strategies. Pathogens. 2023; 12(12):1406.

Chicago/Turabian Style

Bhuvaneswari, Arumugam, Ananganallur Nagarajan Shriram, Kishan Hari K. Raju, and Ashwani Kumar. 2023. "Mosquitoes, Lymphatic Filariasis, and Public Health: A Systematic Review of Anopheles and Aedes Surveillance Strategies" Pathogens 12, no. 12: 1406.

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop