Mechanisms of insecticide resistance in mosquitoes and their implications for disease control

Introduction

Mosquito-borne diseases pose a serious public health challenge in tropical and sub-tropical populations (WHO, 2024). More than 700,000 deaths annually have been reported due to vector-led transmission of pathogens such as Plasmodium, Dengue virus, Chikungunya virus, encephalitis virus, and nematodes (WHO, 2024). These majorly include malaria, which accounted for an estimated 282 million cases and 610,000 deaths globally according to the latest data (WHO, 2025), Furthermore, the global dengue burden has expanded dramatically, reaching a historic peak of over 14.6 million reported cases and exceeding 12,000 deaths (WHO, 2025) yellow fever (estimated 67,000–173,000 severe infections and 31,000–82,000 deaths occur each year in Africa and the Americas) (Institut Pasteur, 2022), chikungunya among other diseases.

Mosquito-borne diseases (malaria, dengue, chikungunya, etc) are transmitted by disease-specific vectors, and the associated pathogen requires usually two host for completion of the life cycle (as shown in Table 1). As adult female mosquitoes require blood meals primarily for egg development, they act as efficient transmitters or carriers of various pathogens within their bodies, whereas adult males feed exclusively on plant nectar and do not blood-feed. Different mosquito species, either singly or in complex, play a critical role in the spread of infectious agents at the local or regional level. A multitude of factors, including abiotic (temperature, humidity, topography, climatic conditions, etc.) and biotic (host type, vector species, parasite interactions), affect the dynamics of mosquito-borne disease spread in a particular region. Also, past anthropogenic activities, such as land-use patterns and urbanization, have profoundly affected the bionomics and behavior of mosquitoes in newer environments.

Table 1. Mosquito-borne diseases along with transmitting vector and pathogen.

Vector control measures primarily focus on reducing the number of disease vectors or averting the interactions between host and vectors. These include physical, biological, and chemical control, which are applied against mosquitoes. Of them, the chemical strategy has been deployed on a large scale in government-led programs across many countries in the past. Different classes of compounds as arsenicals, pyrethroids, organophosphates, chlorinated-hydrocarbons, have been used for mosquito control (Meier et al., 2022) (Figure 1). While pyrethroids are deployed for indoor residual spraying (IRS) in certain settings, their primary and most impactful public health application globally has been as the chemical foundation for insecticide-treated nets (ITNs) and long-lasting insecticidal nets (LLINs). This intensive and prolonged insecticide selection pressure drives the rapid evolution of vector populations; while it selects for resistance alleles already present at low frequencies, resistance also frequently emerges through de novo mutations, structural variations such as gene duplications and copy-number adjustments, and altered regulatory pathways, which subsequently spread across regions via gene flow. Although vector control is recognized as a useful strategy for preventing the illnesses, its full potential is not being realized (WHO, 2014).

Methodology

Review design

This manuscript is structured as a comprehensive narrative review, aimed at synthesizing established paradigms, emerging pathways, and operational vector control policies surrounding insecticide resistance in mosquitoes. Because this topic spans molecular biology, biochemistry, entomological field surveillance, and macro-level global health policies, a narrative review format was selected to provide an interpretive thematic synthesis.

Search strategy and data sources

A systematic search of electronic literature was conducted across primary biomedical and scientific databases, including PubMed, Scopus, and Google Scholar, covering literature published up to 2026. To ensure historical depth and contemporary relevance, the search combined controlled vocabulary such as Medical Subject Headings (MeSH) terms and specialized keywords using Boolean operators. The primary search strings utilized were as follows:

• ▪ (“insecticide resistance” OR “resistance mechanisms”) AND (“mosquito” OR “Anopheles” OR “Aedes” OR “Culex”).
• ▪ (“knockdown resistance” OR “kdr” OR “acetylcholinesterase” OR “ace-1” OR “Rdl”) AND (“target-site insensitivity”).
• ▪ (“cytochrome P450” OR “monooxygenase” OR “glutathione S-transferase” OR “esterase”) AND (“metabolic detoxification”).
• ▪ (“clothianidin” OR “neonicotinoid” OR “chlorfenapyr” OR “pyrrole” OR “broflanilide” OR “meta-diamide” OR “pyriproxyfen” OR “insect growth regulator”) AND (“vector control” OR “LLIN” OR “IRS”).
• ▪ (“insecticide resistance management” OR “surveillance” OR “Integrated Vector Management”) AND (“India”).

The database search was supplemented by manual hand-searches of reference lists from retrieved seminal papers, as well as official technical reports, policy frameworks, and guidelines published by the World Health Organization (WHO) and the National Center for Vector Borne Diseases Control (NCVBDC) of India.

Study selection: inclusion and exclusion criteria

The retrieved literature underwent a multi-stage screening process, beginning with title and abstract evaluation, followed by full-text review. To be included in this synthesis, studies were required to meet the following predefined parameters:

• Relevance: Articles directly assessing physiological, biochemical, metabolic, or cuticular mechanisms of resistance in primary disease-carrying mosquito vectors (Anopheles, Aedes, and Culex).
• Study Design: Peer-reviewed primary research articles, comparative genomic/mRNA expression studies, field evaluations of vector interventions, and official international public health policy documents.
• Language: Formally published literature available in the English language.

Studies focusing strictly on non-vector agricultural pests, non-mosquito insects, or compounds not utilized or trialed in public health vector management were explicitly excluded.

Data synthesis and analysis

Data from the selected studies were analyzed using an iterative thematic approach. The literature was organized conceptually into cohesive domains: first, a timeline mapping the history of legacy chemical vector tools; second, a structural breakdown of target-site insensitivity mutations and metabolic detoxification gene groups; third, an investigation into emerging novel pathways (such as cuticular variations and gut microbiome effects); and finally, an analysis of macro-level resistance-mitigation policies globally and within the specific epidemiological landscape of India.

Mosquitoes and chemical control

The history of chemical control in mosquito vector management extends over a century (as shown in Figure 2), commencing with the application of Paris Green in 1921, one of the initial compounds employed to target mosquito larvae in aquatic environments (Choi and Wilson, 2017; Floore, 2006). DDT (dichlorodiphenyltrichloroethane) was discovered by Paul Hermann Müller in 1939, resulting in its extensive application in the 1940s during World War II for the vector suppression to prevent transmission of vector-borne diseases such as malaria and typhus (Casida and Quistad, 1998; Davies et al., 2007; Nájera et al., 2011). DDT rapidly emerged as the foundation of worldwide mosquito control initiatives owing to its enduring residual properties and cost-effectiveness (Casida and Quistad, 1998). By the 1950s, resistance to DDT began to develop in mosquito populations, particularly Anopheles mosquitoes, undermining control initiatives (N. Liu, 2015). During the 1960s, organophosphates (e.g., malathion) and carbamates (e.g., propoxur) were developed as substitutes for DDT, aiming at distinct enzymatic pathways in mosquitoes (Hemingway et al., 2002). The 1970s witnessed the emergence and utilization of pyrethroids, synthetic derivatives of natural pyrethrins, recognized for their superior efficacy and reduced toxicity to humans, leading to their extensive application in indoor residual spraying and insecticide-treated nets (Ranson and Lissenden, 2016). By the 1990s, mosquitoes had developed resistance to pyrethroids, posing new obstacles for vector control efforts (N. Liu, 2015). In 2004, the World Health Organization (WHO) (2004) advocated for Integrated Vector Management (IVM), a comprehensive strategy that integrated chemical, biological, and environmental methods to diminish dependence on insecticides exclusively. In the 2010s, innovative insecticides with unique mechanisms of action were launched, and resistance management measures, including pesticide rotation, were adopted to combat the escalating resistance issue (WHO, 2020; van den Berg et al., 2021).

The primary classes of public health insecticides, encompassing both the classical neurotoxins and modern resistance-management interventions (Figure 1), are described below with regard to their global deployment timelines, target vectors, and physiological modes of action.

Insecticide classes, resistance patterns, and regional applications in global mosquito control programs

Organochlorines

The organochlorines (OC), notably DDT, were introduced in the 1950s as a key tool for indoor residual spraying (IRS), with India being a primary user, accounting for approximately 82% of global consumption (van den Berg, 2009). These compounds primarily act by disrupting the function of the nervous system in insects, mainly through interference with ion transport across nerve cell membranes by modifying the sodium and potassium channels, leading to hyperexcitation and eventual paralysis (Aktar et al., 2009). OC pesticides can be classified into three main categories: cyclodienes (e.g., chlordane), hexachlorocyclohexanes (e.g., lindane), and dichlorodiphenylethanes (e.g., DDT). However, resistance in Anopheles culicifacies emerged early in India around 1959 (Sharma, 1999), and has since been reported across multiple African countries including Ethiopia, Mozambique, Namibia, South Africa, Zimbabwe, Zambia, Madagascar, Eritrea, Eswatini, Uganda, and Mauritius (van den Berg, 2009; van den Berg et al., 2017; WHO, 2021). Although effectively employed in the past to reduce typhus and malaria, they are now prohibited in the majority of developed nations due to their persistence and potential health risks (Aktar et al., 2009).

Organophosphates

Organophosphates are esters of phosphoric acid and alcohol introduced in the late 1960s (malathion) and 1970s (temephos) respectively. These compounds function by impairing the neurological systems of insects, primarily through the irreversible inhibition of the acetylcholinesterase (AChE) enzyme, resulting in the accumulation of acetylcholine at synapses and neuromuscular junctions. This leads to overstimulation of nerve impulses and ultimately causes paralysis and death of the insect (Costa et al). These compounds have been widely applied for IRS and larviciding across South and Southeast Asia- including India, Bangladesh, Cambodia, Nepal, Sri Lanka, Vietnam- as well as in parts of Africa (South Africa, Madagascar) and the Americas (Brazil, Colombia, Mexico, Peru) (WHO, 2017; van den Berg et al., 2012)). However, malathion resistance was documented in Anopheles populations in India as early as 1973 (Raghavendra et al., 2011), and temephos resistance was reported in Aedes aegypti larvae in Brazil (Belinato et al., 2012). Due to their high toxicity to non-target organisms, their usage is tightly regulated in many countries.

Carbamates

Carbamates are organic compounds derived from carbamic acid that were introduced between the late 1960s and 1970s as structural alternatives to counter early organochlorine (DDT) resistance. Similar to organophosphates, carbamates inhibit the acetylcholinesterase enzyme but do so reversibly, disrupting nerve signal transmission and causing neuromuscular paralysis (Colovic et al., 2013; Fukuto, 1990; Goel and Aggarwal, 2007). Carbamates are organic compounds derived from carbamic acid that were introduced between the late 1960s and 1970s as structural alternatives to counter early organochlorine (DDT) resistance. Similar to organophosphates, carbamates inhibit the acetylcholinesterase enzyme but do so reversibly, disrupting nerve signal transmission and causing neuromuscular paralysis in insects (Colovic et al., 2013; Fukuto, 1990; Goel and Aggarwal, 2007). In modern integrated vector management, they are used primarily as rotational alternatives in regions facing severe pyrethroid resistance. However, because carbamates share a common physiological target enzyme (AChE) with organophosphates, target-site insensitivity via mutations such as G119S routinely triggers broad cross-resistance between these two classes. Consequently, their long-term operational efficacy is increasingly challenged as has been reported in Anopheles gambiae complex mosquitoes in West African countries such as Benin and Nigeria (Djogbénou et al., 2011; Yadouleton et al., 2010) Carbamates, are known for their relatively short environmental persistence, typically breaking down within weeks or months, which makes them a preferable option in certain integrated pest management systems in Africa and Asia including India (Goel and Aggarwal, 2007; Malhotra et al., 2021).

Pyrethroids

The 1970s witnessed the emergence of pyrethroids, synthetic insecticides modeled after pyrethrum, an extract from the Chrysanthemum cinerariaefolium flower. These compounds are classified into two types: Type I pyrethroids, which lack a cyano group at the alpha position (e.g., allethrin, tetramethrin, permethrin), and Type II pyrethroids, which contain an alpha-cyano group (e.g., deltamethrin, cypermethrin, cyfluthrin) (Soderlund et al., 2002). Pyrethroids act by binding to voltage-gated sodium channels in the nerve membranes, delaying their closure and thereby prolonging nerve excitation, resulting in hyperexcitation, paralysis, and ultimately death of the insect (Soderlund et al., 2002). Introduced in India in 1996 (Rahi et al., 2024), they became the cornerstone of IRS and long-lasting insecticide-treated nets (LLINs) due to superior efficacy and reduced mammalian toxicity. However, resistance has emerged over the past decades due to knockdown resistance mutations in sodium channel genes, and detoxification by insect microsomal oxidases. Pyrethroid resistance in Anopheles gambiae and Anopheles funestus has been documented extensively across Africa, Asia (particularly noted in India since 2002, including Gujarat), and the Americas (IR Mapper, 2023; WHO, 2022). To counteract this, inhibitors like piperonyl butoxide (PBO) are often co-applied to block detoxification pathways and enhance insecticidal activity (Davies et al., 2007).

The emergence of widespread target-site and metabolic resistance to these four traditional neurotoxic classes necessitated a paradigm shift in global vector control. To sustain disease suppression, modern vector control frameworks have integrated novel chemical interventions targeting alternative physiological pathways, mitochondrial systems, or allosteric receptors, as detailed below.

Pyrroles

Pyrroles represent a distinctive shift in vector control away from classic neurotoxic pathways (Raghavendra et al., 2011). Chlorfenapyr acts as a pro-insecticide that possesses no intrinsic toxicity until it enters the insect host. Upon ingestion or contact, it undergoes metabolic activation by insect cytochrome P450 monooxygenases (Huang et al., 2023), which cleave its structural pyrrole ring to generate the highly active metabolite AC 303,268, also known as tralopyril (Yunta et al., 2023). This active molecule specifically targets the inner mitochondrial membrane, where it functions as a potent uncoupler of oxidative phosphorylation (Kweyamba et al., 2023). By disrupting the proton electrochemical gradient across the mitochondrial membrane, it halts the synthesis of adenosine triphosphate (ATP), leading to cellular energy depletion, systemic organ failure, and delayed mortality (Huang et al., 2023). Because this cellular pathway is completely independent of the nervous system, pyrroles exhibit no cross-resistance with pyrethroids, organophosphates, or carbamates (Raghavendra et al., 2011). This makes them a critical tool for neutralizing metabolic resistance, leading to their formal WHO recommendation for rollout in next-generation, dual-active long-lasting insecticidal nets (LLINs) to protect regions plagued by highly resistant Anopheles populations (Yunta et al., 2023). Chlorfenapyr-based dual-action LLINs have been recommended for rollout in several African countries, including Benin, Nigeria, Tanzania, Uganda, Rwanda, Mozambique, Malawi, Zambia, and Zimbabwe (Ngufor et al., 2020; WHO, 2023).

Phenyl pyrazoles

Phenyl pyrazoles are potent synthetic neuroactive compounds that disrupt the insect central nervous system via a distinct biochemical mechanism (Gupta, 2018). Fipronil operates as a highly specific, non-competitive antagonist of the insect gamma-aminobutyric acid (GABA)-gated chloride channels as well as insect-specific glutamate-activated chloride channels (GluCls), which are notably absent in mammalian physiology (Narahashi et al., 2010; Zhao et al., 2003). Under baseline conditions, endogenous GABA binding opens these ligand-gated channels, encouraging a stabilizing influx of chloride ions (Cl^-) that hyperpolarizes post-synaptic membranes and dampens over-excitation (Casida and Durkin, 2015). This targeted blockade shifts the neurochemical balance by severing essential inhibitory feedback loops, causing continuous axonal firing, systemic hyper-excitation, muscular convulsions, and lethal neurotoxicity (Gupta, 2018; Narahashi et al., 2010). However, because it binds near the shared cyclodiene binding site, its target-site efficacy can be significantly compromised by cross-resistance in field populations carrying Resistant-to-Dieldrin (rdl) gene locus mutations, specifically the highly conserved Alanine-to-Serine/Glycine substitutions (A296S/G or A302S according to species notations) that allosterically reshape the receptor pore configuration (Ffrench-Constant et al., 2000; Taylor‐Wells et al., 2015).

Neonicotinoids

Neonicotinoids are systemic, neuroactive compounds designed to replicate natural neurotransmitters while resisting immediate enzymatic degradation within the insect host (Uragayala et al., 2015). Clothianidin operates as a highly selective and potent agonist of the insect nicotinic acetylcholine receptors (nAChR), which govern fast excitatory synaptic transmission across the central nervous system (Thany, 2023). Under baseline physiological conditions, endogenous acetylcholine binds briefly to the receptor to propagate electrical signals before being rapidly broken down by the enzyme acetylcholinesterase (AChE). In contrast, clothianidin binds irreversibly to the nAChR binding pocket, resisting hydrolysis and permanently locking the ion channel pore into an open state (Uragayala et al., 2015). This mechanism facilitates a continuous, uncontrolled influx of sodium ions (Na⁺) across the postsynaptic membrane, culminating in sustained axonal depolarization, severe muscular tremors, systemic hyper-excitation, paralysis, and death (Thany, 2023; Uragayala et al., 2015). Because this molecular target site is structurally independent of the voltage-gated sodium channels and the acetylcholinesterase target proteins exploited by legacy chemistries, neonicotinoids express a complete absence of cross-resistance with pyrethroids, organophosphates, carbamates, or DDT. Operationally, clothianidin has been integrated into World Health Organization prequalified Indoor Residual Spraying (IRS) formulations, displaying prolonged residual toxicity on traditional wall substrates like mud and concrete (Agossa et al., 2018). Extensive community trials and pilot networks deployed across high-resistance settings- including Côte d’Ivoire, Benin, and regions in India such as Odisha and Chattisgarh- have successfully bypassed existing kdr-mediated resistance to drastically drop local malaria transmission and clinical case indices (Agossa et al., 2018; Hilton et al., 2024;WHO, 2023).

Insect growth regulators

Insect Growth Regulators (IGRs) represent a highly targeted, biorational strategy in vector management that shifts the focus away from immediate, acute neurotoxicity and toward selective physiological and developmental disruption (Graf, 1993). Pyriproxyfen operates as a highly potent juvenile hormone mimic (or analogue) that targets and interferes with critical endocrine signaling pathways within the insect host (Ishaaya and Horowitz, 1995). Under normal physiological conditions, endogenous juvenile hormone titers must fall sharply to allow the successful transformation of larvae into pupae, and subsequently into functional adult mosquitoes. Pyriproxyfen artificially overrides this drop by maintaining high juvenile hormone-like signaling, thereby causing severe endocrine imbalances during critical molting phases (Mulla, 1995). This disruption permanently arrests development, preventing successful larval-to-adult emergence and leaving immature vectors to die in their aquatic habitats. Furthermore, when adult female mosquitoes pick up sub-lethal doses via contact, pyriproxyfen disrupts oogenesis and follicular development, effectively sterilizing the females or vastly reducing their overall reproductive fitness (Harburguer et al., 2014; Parthasarathy and Palli, 2021). This dual developmental and reproductive impact makes pyriproxyfen an invaluable tool for resistance management. It is widely used in global larviciding programs and has been paired alongside pyrethroids in next-generation long-lasting insecticidal nets (LLINs) (He et al., 2025). Any adult mosquito that survives pyrethroid exposure due to metabolic enzyme upregulation is sterilized by the pyriproxyfen component, safely truncating transmission cycles (He et al., 2025).

Broflanilides

Broflanilide represents a pioneering addition to the chemical arsenal of vector control, establishing an entirely separate structural footprint designed to bypass deep-seated multi-insecticide resistance networks (Govoetchan et al., 2022). Belonging to the novel meta-diamide chemical family- classified under Group 30 by the Insecticide Resistance Action Committee (IRAC)- broflanilide operates primarily as a pro-insecticide (Nakao et al., 2013). Once absorbed or ingested by the mosquito host, it undergoes in vivo dealkylation to yield its highly active metabolite, desmethyl-broflanilide (Nakao et al., 2013). This active form functions as a non-competitive, allosteric modulator that binds directly to a unique hydrophobic pocket within the transmembrane segments of the insect gamma-aminobutyric acid (GABA)-gated chloride ion channels, structurally composed of the Resistant-to-Dieldrin (RDL) subunit locus (Nakao and Banba, 2016; Nakao et al., 2013). Crucially, the desmethyl-broflanilide binding cavity is distinct from the sites utilized by legacy cyclodiene organochlorines (e.g., dieldrin) or phenyl pyrazoles (e.g., fipronil), meaning it remains entirely unaffected by historical cross-resistance mutations (Casida and Durkin, 2015; Nakao and Banba, 2016). By physically keeping the channel pore in a locked-closed conformation, broflanilide blocks the essential inhibitory influx of chloride ions (Cl^-) across the postsynaptic membrane (Nakao et al., 2013). This unmitigated block severs core central nervous system braking loops, precipitating continuous axonal firing, muscular spasms, and systemic exhaustion. Operationally formulated as VECTRON™ T500 for Indoor Residual Spraying (IRS), this chemical class provides exceptionally long-lasting residual bio-efficacy on porous architectural substrates such as mud and concrete (Govoetchan et al., 2022). Multi-centre multi-country trials and experimental hut evaluations conducted in high-resistance settings- such as Covè in Benin and Moshi in Tanzania- have verified that wild populations of Anopheles gambiae s.l. remain fully susceptible to broflanilide (Govoetchan et al., 2022; Snetselaar et al., 2023). It reliably causes high, delayed mortality even against polyfactorial vector strains displaying robust combination resistance markers, including target-site kdr (L1014F/S) mutations, altered acetylcholinesterase targets, and elevated metabolic cytochrome P450 monooxygenase expression (Snetselaar et al., 2023).

Insecticide resistance in mosquitoes

The first report on insecticide resistance to chlorinated hydrocarbon paves the way for deeper insight into the development of resistance in mosquitoes. Widespread insecticide resistance, specifically pyrethroid resistance (accounted for 25% of the insecticide market) is one of the critical issues while devising strategies for controlling mosquitoes (van den Berg et al., 2021). Under intensive selection pressure, the development of a specific resistance phenotype can potentially confer cross-resistance to other chemical compounds, provided they share a common physiological target site or an overlapping metabolic detoxification pathway. However, cross-resistance is not a universal rule or an automatic outcome; its manifestation depends heavily on the specific biochemical mutations or enzyme families upregulated within the localized vector population. In more than 60 countries, there has been evidence of increased mosquito resistance to insecticides, which has an impact on all main vector species and insecticide classes (N. Liu, 2015).

Insecticide resistance is a pre-adaptive evolutionary phenomenon that develops through complex processes that depend on many factors such as genetic, ecological, physiological, and frequency, as well as the amount of insecticide applied (N. Liu, 2015; Shidrawi, 1990; Zhu et al., 2016). It has been proposed that prior to exposure of insecticide or stressor, some rare individuals do have resistance alleles and these alleles should increase after the stressor or insecticide being selected. Because of this, upon the insecticide selection, the frequency of occurrence of resistance alleles has surged in the mosquito population (N. Liu, 2015).

Therefore, it is of paramount importance to decipher how mosquitoes develop resistance and the underlying mechanisms to control the vector population and the spread of resistance. The insecticide acts on different target sites for its function, such as carbamates and organophosphates, and the target site is acetylcholinesterase. Pyrethroids and some organochlorines, including DDT and its analog, have sodium channels as target sites, and GABA is the target site for the rest of the organochlorines (P. Karunaratne et al., 2018; N. Liu, 2015; Taylor-Wells et al., 2015). Many resistance mechanisms have been reported and are categorized into four major groups: 1) Target insensitivity (mutation or alteration of conformation of target site which prevents insecticide binding), 2) Metabolic detoxification of insecticide (increase in the enzyme causing detoxification or inactivation of insecticide) 3) Cuticular resistance (Cuticular thickening and change in the cuticle composition prevent entry of insecticide), 4) Behavioral resistance (Hemingway et al., 2002; Karunaratne et al., 1995).

A detailed schematic is provided in Supplementary Data Sheet 1.

Mechanism of insecticide resistance

Target site insensitivity

Mutation in the structural genes, specifically non-synonymous point mutations leading to amino acid substitutions to target site insensitivity in insects. Although many mutations are possible, only a few mutations will retain the physiological function of that target site and, at the same time, will give insensitivity to insecticides. This mutation in the structural genes will cause a conformational change in the target site, affecting the binding affinities of insecticide. This affinity varies with respect to the molecular structure of insecticide, even if it belongs to the same group of insecticides.

Insensitive GABA receptors

Gamma-aminobutyric acid (GABA) is one of the main inhibitory transmitters that are present in both mammals and insects and binds to a specific class of receptors known as GABA receptors, which are classified into ionotropic (GABA_A) and metabotropic (GABA_B) GABA receptors. These receptors are structurally hetero-multimeric gated chloride-ion channels found in the insect’s central nervous system (CNS) and at neuromuscular junctions. GABA receptor is an oligomer of five subunits, and each subunit comprises a characteristic CYS loop (formed by a disulfide bond between two cysteine residues) and four transmembrane domains (M1-M4). Cyclodines are a sub-category of organochlorines that specifically target these channels. M2 transmembrane domain, which borders the ion channel, has a conserved alanine residue (Bloomquist, 2003) Upon substitutions of this conserved residue in the protein’s chloride ion channel pore provide resistance in many insects (as shown in Table 2). In addition to directly affecting the drug binding site, conserved alanine replacements also allosterically disturb the drug-preferred conformation of the receptor. Alanine to serine mutation(A302S) was initially identified in the Drosophila rdl locus gene coding for the second transmembrane domain of the GABA receptor subunit. Later, this mutation Alanine to Serine (A296S) was observed in insects, including Ae. aegypti and An. Gambiae (Taylor-Wells et al., 2015; Thompson et al., 1993) giving resistance against various organochlorines. In some resistant insects, the resistance occurred by alanine to glycine mutation rather than serine. An. gambiae has been found to have the A296G substitution, while dieldrin resistance in Anopheles arabiensis, Anopheles stephensi, Anopheles funestus, and Ae. aegypti is linked to the A296S substitution (as shown in Figure 3 -common mutation providing resistance to range of mosquito’s species). Anopheles gambiae, a malaria mosquito that exhibits significant resistance to dieldrin, was used to molecularly clone the rdl cDNA, revealing both this mutation (A296G) and another in the third transmembrane domain (T345M) are involved in resistance in insects. Using the Xenopus laevis oocytes expression system, the sensitivity to insecticide (imidacloprid, deltamethrin, and fipronil) is reduced because of the A296G and A296G + T324M mutation while there is no effect of T324 alone. In conclusion, this study identifies another mutation in the mosquito rdl that is associated with insecticide resistance. While T345M itself does not affect insecticide sensitivity, it may serve to offset the structural impact of A296G (Taylor-Wells et al., 2015).

Table 2. The table comprises of genetic mutations and alteration gene expression in insecticide target which confer to insecticide resistance.

Sodium channels

Voltage-gated sodium channels are crucial in action potential initiation and propagation in the nervous system (Du et al., 2016).Upon opening the voltage-gated sodium channels, a rapid influx of sodium ions depolarizes the axonal membrane and gets inactivated after reaching the sodium equilibrium potential. This inactivation has a crucial role in the termination of action potential propagation by decreasing the membrane potential. Because of this critical role in propagating impulses in excitable cells, the voltage-gated sodium channel is a potent target for various natural and synthetic neurotoxins such as DDT and pyrethroid insecticides (Dong et al., 2014). These insecticides bind to sodium channels and confer conformational change, leading to a change in gating kinetics that result in prolonged activation of the channel. Prolonged opening of this channel will continuously excite the nerve and cell, leading to paralysis and ultimately death of insects (schematic shown in Figure 4).

Like their mammalian counterparts, insect sodium channels have four homologous repeat domains (I–IV), each possessing six α-helical transmembrane segments (S1–S6). In each domain, the S1–S4 segments constitute the voltage-sensing module. The segments S5 and S6 from the four domains, in addition to the four membrane-reentrant P-loops that connect the S5 and S6 segments, form the pore module (N. Liu, 2015). The four voltage-sensing modules are arranged quasi-symmetrically around the pore module’s outer rim. Each S4 segment, which serves as a voltage sensor of the channel, contains repeated motifs (a positively charged amino acid residue followed by two hydrophobic residues). Structural modification in this native structure can impair the function.

In the past three decades, many mutations have been identified in sodium channels, which alter the usual function. Sodium channels become insensitive to DDT and pyrethroids due to structural modifications in the form of point mutations or substitutions brought about by single nucleotide polymorphisms. This is because the binding affinity decreased or was eliminated, generating resistance against both DDT and pyrethroids in insects. The earliest evidence of sodium channel insensitivity to pesticide inhibition was found in Musca domestica, where it manifested as knock-down resistance (KDR) (Farnham and Sawicki, 1976). Numerous reports have surfaced of possible KDR-like resistance in mosquitoes, based on cross-resistance between DDT and pyrethroids acting on the same sodium channel location (Du et al., 2016).

The association of many distinct mutations in sodium channel genes with decreased sensitivity to pyrethroid and cross-resistance to DDT is being identified in many insects, including mosquitoes. The most common pyrethroid-associated resistance mechanism is the mutation from leucine to phenylalanine at codon 104 caused by an A to T transversion in the S6 hydrophobic section of domain II (L. Zhou et al., 2009) (Figure 5). This L1014F mutation was the first pyrethroid resistance-associated mutation identified in houseflies and German cockroaches. Later it has been reported in many species of Anopheles and some species of Culex. Leucine to Phenylalanine (L1014F mutation) is reported in An. gambiae in West Africa region (Martinez‐Torres et al., 1998), An. stephensi (Enayati et al., 2003) in Dubai region, An. gambie (Diabate et al., 2004) in Burkina Faso (West Africa), An. sacharovi in Turkey, An. Subpictus in Srilanka, An. culicifacies (Singh et al., 2009) in India, An. arabiensis (Ochomo et al., 2013; Stump et al., 2004) in Western Kenya, Cx. pipiens (Martinez-Torres et al., 1999; Xu et al., 2006) in East Coast USA (Figure 6A; Table 2). Many other mutations associated with knockdown resistance (kdr) were reported. L1014S was Cx. pipiens (Bkhache et al., 2016; Martinez‐Torres et al., 1998; Martinez-Torres et al., 1999), An. arabiensis (Hemingway and Ranson, 2000), An. sacharovi (Lüleyap et al., 2002), An. culicifacies (Singh et al., 2010; Figure 6B; Table 2). In An. culicifacies, apart from the two more common alternative kdr mutation L1014F/S in sodium channel, a resistance-associated mutation V1010L was reported from the population of Odisha (Singh et al., 2010) (Table 2; Figure 5).

Surprisingly, Kdr-associated L1014F mutation has not yet been identified in Aedes aegypti. Rather ten new resistance-associated mutation was identified in Aedes aegypti in various countries. G923V and I1011M Substitution were reported in Brazil, Guyana and Martinque, L982W in Vietnam, V1016G in Indonesia and Thailand, I1011V and V1016I in latin America, F1534C in Thailand and Vietnam, S989P + V1016G and D1736Y + V1016G found to be coexisted in Thailand and Taiwan respectively (Du et al., 2016) (Table 2; Figure 5). S989P Frequently co-occurring with other mutations, and it contributes significantly to the overall resistance profile of mosquito populations (Tognarelli et al., 2025) (Figure 5). A recent study by Taranjeet Kaur et al. (2025) investigated knockdown resistance (kdr) mutations in Indian Aedes aegypti populations, which are a key mechanism of insecticide resistance, with a particular focus on the V1016G, F1534C, and S989P mutations, which confer high pyrethroid resistance when combined on a single haplotype (Figure 5). Collected mosquito specimens from eight locations, genotyping them for kdr alleles and intron haplotypes using PCR-based assays and DNA sequencing. And identified five kdr mutations (S989P, V1016G, T1520I, F1534C, F1534L) and two distinct intron haplotypes (intron-A and intron-B). These mutations were classified into three distinct haplogroups: haplogroup G (V1016G with/without S989P and intron-A), haplogroup L (F1534L and intron-A), and haplogroup C (F1534C with/without T1520I and intron-B) (Figure 5). Crucially, the study found no evidence of recombination among these three haplogroups within Indian populations. This lack of recombination implies that the highly potent triple kdr mutation haplotype (S989P, V1016G, F1534C) is currently absent in these populations, suggesting that existing vector control measures may remain effective and could delay the emergence of highly resistant mosquito phenotypes (Kaur et al., 2025).

Acetylcholinesterase

Acetylcholinesterase (AChE) is an enzyme that hydrolyzes the acetylcholine neurotransmitter on the post-synaptic neuron. This enzyme is identified as a potent target site for insecticides like organophosphates and carbamates. Upon binding of these insecticides in the acetylcholinesterase active site, the enzyme gets inhibited and continuously stimulates the postsynaptic neuronal membrane, resulting in the death of insects. Many insects have developed resistance against organophosphates and carbamates and these resistant insects have altered AChE, which makes them resistant to insecticides. Two different isoforms of AChE have been found: AChE1 (Cholinergic- the protein responsible for synaptic transmission) and AChE2 (non-cholinergic). In insects, these two enzymes (AChE1 and AChE2) are coded by ace-1 and ace-2 genes, respectively (N. Liu, 2015). The ace-1 gene is closely linked to resistance development in mosquitoes. A mutation in the ace-1 gene reduces the affinity of substrate binding sites in the acetylcholinesterase enzyme. Many mutations have been reported to be linked to the development of resistance to organophosphates and Carbamates. G119S is one of the most commonly found mutations associated with acetylcholinesterase (AChE) inactivation in several mosquito species: Anopheles albimanus, Anopheles nigerimus, Anopheles atroparvus, Anopheles subpictus, Anopheles gambiae, Culex pipiens (in Africa and Caribbean, Europe), Culex vishnui (in China) (N. Liu, 2015) (Table 2; Figure 7). Structural analysis has indicated that a larger size of the serine sidechain (-CH₂OH) in comparison to the glycine sidechain (-H) would impart steric hindrance in the inhibitor binding site. Hence, the mutation of Glycine to Serine has been shown to decrease the accessibility of the inhibitors as well as the substrate (Alout and Weill, 2008).

Metabolic detoxification

Insects are exposed to various toxins known as xenobiotics, which are produced by plants (naturally) or humans (artificially). Insects have evolved various detoxification mechanisms over time to survive these toxins. Mosquitoes undergo detoxification of insecticides through three primary metabolic detoxification gene families: cytochrome P450S (P450S), glutathione S-transferases (GSTs), and carboxylesterases (Hazarika et al., 2025; N. Liu, 2015) (Figure 8). Metabolic enzyme genes have a higher degree of adaptability compared to insecticide target site genes (P. Karunaratne et al., 2018) and hence increased levels of metabolic enzyme cytochrome P450s, GSTs, and esterase confer metabolic detoxification. This increased level of cytochrome P450s, GSTs, and esterase capability to break down or sequester toxic compounds is achieved through broad categories of mechanisms: gene amplifications, gene copy-number variations (CNVs), regulatory changes, and structural mutations. (a) Gene amplification and copy-number variations (CNVs) involve the physical multiplication of specific genomic loci or entire gene clusters, expanding the cellular gene pool to yield massive protein production; this is widespread at the cytochrome P450 cluster Cyp6aa/p, the glutathione S-transferase Gste2, and alpha-esterase clusters (Coeae1f-2f) across major malaria vectors like Anopheles gambiae and Anopheles arabiensis (Eukubay et al., 2026; Nagi et al., 2024). In contrast, (b) gene overexpression/transcriptional upregulation can occur independently of physical gene duplication, where elevated enzyme quantities are driven by regulatory variations. These regulatory variations encompass modifications in cis-acting promoter or enhancer elements (Wilding et al., 2012), or changes in trans-acting transcription factors (such as Maf-S signaling) that orchestrate broad-spectrum metabolic networks. Beyond quantitative changes in enzyme abundance, metabolic detoxification is also driven by structural mutations, which are non-synonymous point mutations within the protein-coding sequences of metabolic genes (Alout and Weill, 2008). These structural polymorphisms alter the spatial conformation of the enzyme’s catalytic pocket, significantly enhancing its specific substrate affinity or degradation velocity toward chemical insecticides. Additionally, structural duplications can occasionally manifest as heterogeneous duplications- combining an intact wild-type allele with a structurally mutated resistance allele- allowing field populations to maintain vital physiological function while simultaneously acquiring profound chemical tolerance (Xiao et al., 2026).

Monooxygenase

Monooxygenase, otherwise known as Cytochrome P450s dependent mono-oxygenase, is crucial as it is involved in catabolism and anabolism (synthesis) of xenobiotics (Scott, 1999; N. Liu, 2015). These enzymes belong to the oxidoreductase class and are quite versatile, oxidizing a wide range of substrates and triggering cascades of metabolic reactions. Cytochrome P450s, being the terminal oxidase enzyme in the mono-oxygenase system, bind to molecular oxygen (O2) and are subsequently reduced by electrons transferred from NADPH to incorporate one hydroxyl group (-OH) in the substrate (Scott, 1999). This enzymatic network is ubiquitously distributed across diverse taxa, including plants, mammals, bacteria, and insects. In insects, increased activity of the monooxygenase enzyme through the metabolic detoxification of insecticides confers resistance. The significance of this enzyme in resistance development was apparent when a study showed that sesamex, a P450 inhibitor could eliminate the resistance against carbaryl (Eldefrawi et al., 1960). A large number of P450s were purified and characterized, leading to specific nomenclature of each P450 superfamily with CYP as a prefix (corresponding to CYtochrome P450) followed by a number (for family), letter (for subfamily) and number (for isoform).

Metabolic and genomic P450 drivers in Anopheles

Within major malaria vectors, the molecular and genomic architecture governing pyrethroid resistance has evolved far beyond simple gene expression alterations, transitioning into highly selected macro-genomic structural modifications and novel locus deployments. In Anopheles funestus, the physical duplication and corresponding transcriptional overexpression of the Cyp6p9a, Cyp6p9b, and Cyp6p4 genes remain the foundational drivers of metabolic pyrethroid resistance across Southern and East Africa (Riveron et al., 2013; Wondji et al., 2009) (Table 2). However, recent multi-omics temporal profiling reveals a much more intricate network of parallel markers—including upregulation of V-ATPase, tubulin alpha-1, and various transposases—driving the escalation of high-intensity resistance across the African continent (Gadji et al., 2025).

Beyond the foundational Cyp6p9 cluster, Cyp6p5 and Cyp9k1 are now recognized as critical parallel drivers of pyrethroidi clearance (Mugenzi et al., 2020; Djoko Tagne et al., 2025). In East and Central Africa, high-intensity pyrethroid resistance and the resulting failure of insecticidal bed net interventions have been directly linked to a highly selected, non-synonymous single nucleotide polymorphism (SNP) within Cyp9k1—specifically the G454A mutation—which significantly increases the degradation velocity of both Type I and Type II pyrethroids (Tchouakui et al., 2025).

Concurrently, adaptive evolution in An. funestus field populations have favored prominent structural variants (SVs) that systematically modulate gene expression landscapes inside these major P450 clusters (Mugenzi et al., 2024). Genomic mapping has identified key structural variants, notably a 6.5kb insertion (Mugenzi et al., 2020) and a rapidly selected, retrotransposon-containing 4.3kb insertion localized within the intergenic sequence separating Cyp6p5 and Cyp6p9b (Mugenzi et al., 2024). This 4.3kb structural variant harbors enriched binding motifs for key xenobiotic-responsive transcription factors and functions as a potent enhancer element that robustly upregulates transcription of adjacent Cyp6p9a/b alleles without altering physical gene copy number (Mugenzi et al., 2024). This transposable element-driven driver has undergone intense directional selection across Central and East Africa, illustrating how macro-structural genomic alterations facilitate rapid phenotypic escalation against both pyrethroid-only and piperonyl butoxide (PBO)-synergized vector control tools (Gadji et al., 2025; Mugenzi et al., 2024).

Other Anopheles species exhibit similar reliance on oxidative detoxification pathways. In Anopheles gambiae, metabolic detoxification acts in concert with target-site insensitivity to nullify pyrethroid efficacy, primarily driven by the transcriptional upregulation of Cyp6p3, Cyp6z1, and Cyp6m2 in resistant wild populations (Boussougou-Sambe et al., 2024) (Figure 9). Similarly, comparative mRNA expression studies in Thailand have demonstrated that the Cyp6aa2 transcript is explicitly upregulated in deltamethrin-resistant Anopheles minimus, whereas Cyp6p5 remains equally expressed between susceptible and resistant strains, confirming that Cyp6aa2 overexpression is the principal driver of the resistant phenotype in this species (Rongnoparut et al., 2003) (Table 2).

P450-mediated resistance in genera Culex and Aedes

In Culex species, metabolic resistance is conferred by localized gene amplification and duplication events. For instance, the Cyp9m10 gene exhibits a massive 260-fold overexpression in pyrethroid-resistant strains of Culex quinquefasciatus. This extreme overexpression is driven by the macro-genomic duplication of an approximately 100kb region, accompanied by specific transposable element insertions and a critical cis-acting regulatory mutation (Itokawa et al., 2010, 2011). In addition to Cyp9m10, another P450 gene, Cyp6pf1, is notably upregulated in resistant Cx quinquefasciatus strains compared to susceptible cohorts (Gong et al., 2022; Itokawa et al., 2010) (Table 2). Further cross-species analysis has confirmed that the overexpression of Cyp6pf1, along with specific Cyp4 family genes, directly underpins deltamethrin resistance in another closely related vector, Culex pipiens pallens (P. Karunaratne et al., 2018; Shen et al., 2003; Gong et al., 2005) (Figure 9).

Finally, in the genus Aedes, pyrethroid resistance is mediated by a distinct suite of phase-I metabolic enzymes. Specifically, the coordinated transcriptional upregulation of the Cyp9j24, Cyp9j26, Cyp9j28, and Cyp9j32 genes in Aedes aegypti allows the mosquito to rapidly metabolize and inactivate pyrethroid compounds, establishing the Cyp9j gene subfamily as the dominant oxidative driver of chemical tolerance within this arbovirus vector (Saavedra‐Rodriguez et al., 2012; Stevenson et al., 2012; Strode et al., 2008) (Table 2).

Glutathione S transferase

Glutathione S Transferase (GSTs) are a large family of hetero or homo dimeric multifunctional enzymes found extensively in all aerobic organisms. GSTs are essential for detoxifying and metabolizing a wide range of endogenous and xenobiotic compounds (including insecticides). They also protect the cell against oxidative stress from ROS. GSTs function by catalyzing dehydrochlorination reactions or by conjugation reactions. Conjugation reaction makes the compounds more water soluble and excretable by conjugating electrophilic compound to the thiol group of reduced glutathione (GSH) (Enayati et al., 2005).

Many studies about insect GSTs are primarily focused on the detoxification mechanism of nonself compounds like insecticides and the development of resistance (Hemingway et al., 2002; Ranson et al., 2001; Sawicki et al., 2003). Increased activity of GST in insects is the main cause and multiple pathways of GSTs are involved in generating resistance to different insecticides like O-dealkylation or O-dearylation conjugation mechanism for organophosphates, dehydrochlorination and conjugation pathways for organochlorines (DDT) (Che-Mendoza et al., 2009) (Figure 10). GSTs also play a crucial role in reduction of the oxidative stress due to insecticides. Atleast six classes of GSTs are identified based on greek letter nomenclature out of which delta and epsilon (earlier referred as class I and III GST respectively) are insect-specific and confer resistance to many insecticides (Ranson et al., 2001; Ortelli et al., 2003) (Table 2).

Novel class III insect GSTs which include AgGST3-2, confer DDT resistance in Anopheles gambiae, with high dehydrochlorinase activity and overexpression in resistant strains (Ranson et al., 2001) (Table 2). agGSTd1–6 increased activity was also reported to be associated with DDT resistance in Anopheles gambiae but was found to be inhibited by pyrethroid insecticides (Chen et al., 2003; Y. Wang et al., 2008) (Figure 11). agGSTe2 which belongs to epsilon class GSTs found to be crucial in Anopheles gambiae because of its high DDT-detoxifying activity conferring DDT resistance (Y. Wang et al., 2008). In addition to GSTe2, GSTd3 a delta class GST is upregulated in resistant strains of many malaria vector species. Recombinant expression of GSTe2 and GSTd3 from Anopheles arabiensis suggest that although both metabolize DDT, GSTe2 has high DDT dehydrochlorinase activity in comparison to GSTd3 suggesting that GSTd3 in Anopheles arabiensis sequester DDT (Lu et al., 2023; N’Dri et al., 2023). Purification of GST-1 and GST-2 from larvae of Aedes aegypti using chromatographic techniques and characterization shows high enzyme activity in resistant strain in comparison to the susceptible strain of Aedes aegypti (Grant et al., 1991; Grant and Matsumura, 1989; Lumjuan et al., 2005) (Table 2). The molecular mechanism behind the GST-based resistance in Ae. aegypti, is probably caused by a trans-acting regulator.

Carboxyl esterase

Carboxylesterase is an enzyme belonging to the serine hydrolase superfamily involved in metabolic resistance in mosquitoes, aiding in xenobiotic detoxification alongside cytochrome P450 and GST enzymes. Carboxyl esterase functions by hydrolysis of ester, amides, thioester, and carbamates.

Esterase-based metabolic detoxification is widely studied in Culex genus, where increased enzyme activity underpins organophosphate resistance in more than 80% of populations worldwide (P. Karunaratne et al., 2018). Rather than rapidly metabolizing the organophosphate compound, these carboxylesterases predominantly operate via a mass-sequestration mechanism (Karunaratne et al., 1995). Gene amplification and copy-number duplication represent the major molecular drivers responsible for these elevated enzyme phenotypes.

The phenotype associated with organophosphates resistance involves the co-amplification of the estα2 and estβ2 alleles (historically designated as A2 and B2), which can be amplified more than 250-fold, driving an approximate 800-fold increase in organophosphate tolerance in resistant strains of Culex pipiens quinquefasciatus (Mouchès et al., 1986, 1987) (Table 2; Figure 12). Similarly, Culex tritaeniorhynchus has the Ctrestb1 gene that is amplified once, conferring organophosphate resistance (Mouches et al., 1990; Hemingway & Karunaratne, 1998) (Table 2). While increased esterase activity is usually linked to gene amplification, this is not ubiquitous. For instance, increased estα1 gene expression in Culex pipiens (from southern France) was not due to amplification but rather was speculated to be the involvement of a peculiar regulatory element (Gullemaud et al., 1997). In Culex tarsalis, two resistance mechanisms co-exist to provide resistance: one with elevated sequestering esterases, the other involving non-elevated metabolically active esterases (Ziegler et al., 1987).

While Culex species usually have an elevated esterase mechanism, some Anopheles species, in contrast, have a non-elevated esterase mechanism, which provides resistance against malathion, through increased metabolism of insecticide. (P. Karunaratne et al., 1995; P. Karunaratne and Hemingway, 1996; Hemingway and Ranson, 2000). In Anopheles gambiae, functional genomic profiling has identified the specific carboxylesterase COEAE80 as a primary metabolic determinant of field resistance to chlorfenapyr (Tchouakui et al., 2025). Chlorfenapyr functions as a pro-insecticide that requires activation by metabolic enzymes into its toxic form; however, wild-caught populations surviving diagnostic exposures exhibit a highly selected structural polymorphism—the E510D-COEAE80 variant. This specific non-synonymous substitution drastically impairs standard pro-insecticide processing, directly facilitating chlorfenapyr survival while simultaneously providing cross-resistance to pyrethroids (Tchouakui et al., 2025). This paradigm shift highlights that carboxylesterases are not merely legacy remnants of 20th-century chemical usage, but actively expanding genomic barriers threatening contemporary dual-active ingredient long-lasting insecticidal nets (LLINs) across Africa.

Cuticular resistance

Apart from the extensively studied resistance mechanisms like target site insensitivity and metabolic detoxification of enzymes, many other mechanisms do exist that are less studied, such as Cuticular resistance. Cuticular resistance in mosquitoes is mediated by altering the cuticle composition. In this resistance mechanism, resistance is accomplished by changing the cuticle’s composition or by making it thicker, structural elements such as epicuticular lipids and structural cuticular proteins are accumulated in the cuticle impeding the entry of insecticides to the target sites, and as a result, very less insecticide can penetrate the surface and reach the target site (Jacobs et al., 2023; Balabanidou et al., 2018).

Anopheles funestus Giles, the primary vector for malaria in South Africa, has developed resistance to Pyrethroids due to metabolic detoxification and cuticular thickening. The Mean cuticular thickness calculated (as shown in below figure) was greater in the resistant strain than in the susceptible strain of Anopheles funestus (Figure 13). As a result, this Cuticle thickening impedes the insecticide entry, which in turn increases the efficacy of metabolic enzymes (Jacobs et al., 2023; Balabanidou et al., 2018; Wood et al., 2010).

Resistance due to gut microbiota

The impact of insecticide resistance on the commensal microbiota of malaria vectors is significant, as emerging research indicates that gut bacteria play a crucial role in these mosquitoes’ resistance mechanisms. The relationship between insecticide resistance and gut microbiota diversity suggests that specific bacterial populations can influence the susceptibility of mosquitoes to insecticides, thereby affecting vector control strategies.

It has been shown that the gut microbiota of mosquitoes actively promotes insecticide resistance via two primary physiological pathways: directly isolating and biochemically degrading insecticidal xenobiotics, or systematically altering the expression of endogenous host genes and phase-I/II metabolic detoxification enzymes (Barnard et al., 2019; H. Liu et al., 2024). Crucially, recent functional metagenomic profiling has demonstrated that the mosquito microbiome actively contributes to the escalation of pyrethroid resistance in the primary African malaria vectors, Anopheles gambiae s.s. and Anopheles funestus s.s (Djondji Kamga et al., 2025). Moreover, targeted broad-spectrum antibiotic treatments that deplete these commensal microbial communities lead to a significant increase in insecticide susceptibility, thereby directly reducing mosquitoes’ tolerance to pyrethroids and highlighting the cooperative host-microbiota defense network (Djondji Kamga et al., 2025). Many other studies have shown insecticide-resistant Anopheles arabiensis exhibits lower gut bacterial diversity, with specific bacteria potentially enhancing resistance. Antibiotic treatments can alter gut microbiota, affecting detoxification enzyme activity and influencing insecticide tolerance, underscoring the complex interplay between gut bacteria and resistance phenotypes (Barnard et al., 2019). Furthermore, a study involving whole metagenome sequencing reveals that the gut microbiota of Anopheles albimanus shows significant differences between insecticide-resistant (FEN_Res) and susceptible (FEN_Sus) mosquitoes, with OP-degrading bacteria enriched in FEN_Res, suggesting a potential role of microbiota in conferring insecticide resistance (Dada et al., 2018) (Table 3).

Table 3. Comparative analysis of microbial communities in resistant vs. Susceptible mosquitoes.

Interestingly, 16S rRNA sequencing of the microbiota of the Anopheles coluzzii mosquito midgut in deltamethrin-resistant and susceptible strains reveals significant differences in microbiota diversity. Resistant mosquitoes had a wider variety of bacteria, including genera known to degrade insecticides, such as- Ochrobactrum, Lysinibacillus, and Stenotrophomonas. In contrast, susceptible mosquitoes had comparably reduced microbial diversity dominated by Serratia and Asaia (Pelloquin et al., 2021).

Novel mechanisms and genes involved in insecticide resistance

Insecticide resistance mechanisms in mosquitoes are highly complex and not yet fully understood (Kumar et al., 2024). Recently, several novel candidate genes have been implicated in resistance, despite no prior evidence of their involvement. These discoveries have unveiled previously unrecognized mechanisms of resistance in mosquito vectors (Ingham et al., 2018). For instance, the upregulation of ABC transporters has been linked to pyrethroid resistance in Anopheles gambiae (Pignatelli et al., 2018). The RNAi-mediated gene silencing of distinct functional genes such as hexamerins (AGAP001659), α-crystallins (AGAP007159), and F-type ATP synthase (AGAP006879) has been shown to enhance pyrethroid toxicity in An. gambiae (Ingham et al., 2018). α-crystallins are stress-induced, ATP-independent molecular chaperones that bind to misfolded proteins, preventing their toxic aggregation (Landis et al., 2012). Hexamerins, the most abundant proteins in the mosquito haemolymph, function as transport and storage proteins (Burmester and Scheller, 1999) whereas, F-type ATP synthase, found in the plasma membrane of the insect midgut and salivary glands, is involved in lipid transport (Fruttero et al., 2014, 2017). Further, studies have highlighted the role of the transcriptional factor Maf-S, which modulates key insecticide resistance genes, in contributing to pyrethroid resistance. Silencing Maf-S showed to increase the susceptibility to pyrethroid insecticides in An. Gambiae (Ingham et al., 2017). Overexpression of NYD-OP7 (GenBank: AY749413), a member of Gq-coupled opsin subfamily has been associated to pyrethroid resistance in Aedes albopictus and Cx. pipiens pallens (Hu et al., 2007; Wu et al., 2004; D. Zhou et al., 2018). The gene silencing of NYD-OP7 and its downstream effector molecule Phospholipase C also showed reversal to pyrethroid susceptibility in the resistant Cx. pipiens pallens(D. Zhou et al., 2018). Similarly, RNAi mediated gene silencing of the legs-associated Sensory Appendage Protein (SAP2) has been shown to reverse pyrethroid resistance through a sequestration mechanism in An. gambiae (Ingham et al., 2020). Uridine-diphosphate (UDP)-glycosyltransferases (UGTs), characterized as phase-II detoxification enzymes responsible for excreting xenobiotic compounds (Meech et al., 2012), have also been associated with pyrethroid resistance across multiple species of Anopheles, including An. gambiae, An. coluzzii, An. funestus, and An. arabiensis (Logan et al., 2024). A novel protein, Cub and Sushi Domain-containing Insecticide Resistance (CSDIR), has recently been identified as contributing to resistance against both organophosphates and pyrethroids in An. stephensi (Kumar et al., 2024). CSDIR is a membrane-associated protein that interacts with and metabolizes pyrethroid insecticides extracellularly, further emphasizing the expanding complexity of insecticide resistance mechanisms in mosquitoes.

Mitigation strategies for insecticide resistance

Insecticide resistance in mosquitoes poses a significant challenge to public health. Effective mitigation strategies are crucial to avert the spread of insecticide resistance and to prolong the efficacy of insecticides. An integrated approach that combines multiple strategies to reduce selection pressure and preserve the efficacy of available insecticides against mosquitoes. One of the most efficient methods is insecticide resistance management (IRM), which includes rotating insecticides, insecticide mixture and mosaic approaches, non-chemical interventions, and resistance monitoring.

Insecticide rotation involves periodically changing the different classes of insecticides (i.e, with different modes of action) so that mosquitoes do not experience consistent selection pressure from a class of insecticide. The basis for this method is that the resistance allele is less likely to persist in the population when classes of insecticides are changed periodically, which will potentially reverse resistance (Dusfour et al., 2019; Hobbs et al., 2023). Whereas, the Mosaic approach involves the application of different classes of insecticides in spatially separated areas, exposing neighbouring mosquito populations to distinct selective pressures and thereby limiting the overall spread of resistance alleles (Dusfour et al., 2019). Insecticide mixtures strategies to mitigate insecticide resistance involve two or more insecticides with different modes of action simultaneously, and it is less likely that a mosquito will concomitantly develop resistance to the insecticides, especially when cross-resistance is absent (Hobbs et al., 2023; South and Hastings, 2018). However, this mixture method is effective only if both compounds have similar residual efficacy and are not subject to cross-resistance (Dusfour et al., 2019). Non-chemical interventions include a combination of insecticidal approaches with non-chemical methods, such as reducing the source (removal of breeding sites), larval habitat management, biological control, and genetic technologies, which can help reduce reliance on chemicals, consequently negating the selection pressure and slowing down the evolution of resistance (Dusfour et al., 2019). Resistance monitoring through bioassays (e.g., CDC Bottle Bioassays, WHO Tube tests) and molecular markers helps tailor interventions appropriately and detect emerging resistance early, allowing for timely adjustment of control strategies (Hazarika et al., 2025).

The development and judicious use of new insecticides with novel modes of action (such as Sulfoxaflor, fipronil, Pyrazoles and pyridazinones, Anthranilic diamides and Neonicotinoids neuroactive insecticides) as well as innovative methods such as genetically modified mosquitoes or Wolbachia-based strategies, Sterile insect technique (SIT) and incompatible insect techniques are necessary for sustainable long-term resistance management (Hazarika et al., 2025). Many cases have been reported, such as in Tapachula, Mexico, where the cessation of pyrethroid use led to a significant decline in resistance levels in Aedes aegypti, demonstrating that IRM can effectively reverse resistance when implemented correctly (Penilla-Navarro et al., 2024).

Wolbachia-based strategies

Mosquito-borne arboviral diseases pose a global threat due to rapid resistance development in mosquitoes and the lack of authorized vaccines and treatments, highlighting the need for efficient vector control (Utarini et al., 2021). Wolbachia are intracellular bacteria that infect many insect species, including several major mosquito vectors, and impart a pathogen-blocking effect. Wolbachia-based strategies provide an environmentally friendly, sustainable biological alternative to heavy insecticide reliance, which reduces the selection pressure imparted on mosquitoes and impedes the development of resistance in mosquito populations. Their use in vector control programs offers unique avenues for mitigating insecticide resistance. However, an incomplete understanding of the mechanism by which Wolbachia blocks pathogens in mosquitoes reduces its reliability in this control method (Yen and Failloux, 2020). Environmental factors, such as temperature and host diet, reportedly determine Wolbachia infection. These factors can affect the bacterial densities within mosquitoes, thereby impacting their virus-blocking ability (Yen and Failloux, 2020). Some specific strains of Wolbachia in Ae. aegypti upregulate the host immune gene expression, leading to proliferation of many immune effector molecules, thus inhibiting dengue and chikungunya virus replication (Moreira et al., 2009). Wolbachia-infected Culex quinquefasciatus have also been shown to increase resistance to West Nile Virus infection (Glaser and Meola, 2010). Wolbachia strain wMelPop infection upregulates several immune genes, such as LRIM1 (Leucine-rich repeat immune molecule 1) and TEP1 (thioester-containing protein 1) in An. gambiae, leading to inhibition of the Plasmodium parasite (Blandin et al., 2004; Hughes et al., 2011; Kambris et al., 2010; Povelones et al., 2009). Wolbachia has been seen to alter the reproduction of insects by cytoplasmic incompatibility (CI). This gene drive mechanism leads to no development (or embryonic lethality) of eggs (from Wolbachia uninfected females) when fertilized with sperm from Wolbachia-infected males. In contrast, male and female Wolbachia-infected insects give viable embryos. This CI-based mechanism shows a new avenue for insect control (Adams et al., 2021; Beckmann et al., 2017). Recent Studies have also shown cytoplasmic incompatibility induction due to Wolbachia cifB genes (a prophage gene that encodes CI) in the malaria vector (Adams et al., 2021).

Genetically modified mosquitoes

Genetically modified (GM) mosquitoes are emerging as a frontier strategy to mitigate insecticide resistance and restore the effectiveness of vector control methods against diseases like malaria and dengue. These approaches use advanced genetic engineering to alter mosquito populations in ways that target resistance mechanisms directly or reduce selective pressure for resistance genes. Major milestones in vector control have been achieved by conducting caged field trials and preparing for environmental release trials of these genetically modified (GM) insects. However, these breakthroughs also bring forward important questions about how to properly design and carry out such trials, ensuring that they are ethically sound and socially and culturally acceptable to the communities involved (G.-H. Wang et al., 2021; Xia et al., 2021). Researchers have used CRISPR/Cas9-based gene drives to replace insecticide resistance alleles (such as kdr mutations in the voltage-gated sodium channel gene) with the natural, susceptible version. This drives successfully reversed 83% resistant and 17% susceptible Drosophila population to 87% susceptible, 13% resistant population in just 10 generations in laboratory experiments, demonstrating the potential to restore insecticide sensitivity in wild populations (Kaduskar et al., 2022). Similar CRISPR/Cas9 based gene drive can also be develop for mosquitoes. Transgenic mosquitoes with Gene drives can also spread traits that suppress mosquito populations or render them incapable of transmitting pathogens, changing the mosquito behavior (such as increased outdoor biting than indoor), indirectly reducing reliance on chemical insecticides and thereby lowering resistance selection pressure (Selvaraj et al., 2020). So, Mosquito population suppression and mosquito replacement are two broad strategies of Gene drive methods to control vectors (Burt, 2003; Ribeiro and Kidwell, 1994; Selvaraj et al., 2020).

Sterile insect techniques

Sterile Insect Technique (SIT) is a promising, environmentally friendly strategy for mitigating insecticide resistance and controlling mosquito populations, especially where resistance to chemical treatments is widespread. Large numbers of male mosquitoes are reared and then sterilized using irradiation (X-rays or gamma rays), leading to germ cell degeneration and dominant lethal mutation in sperm. These males are unable to produce viable offspring but remain competitive for mating (Robinson, 2021).By releasing sterilized male insects to mate with wild females, SIT disrupts reproduction and gradually reduces pest populations. Recent advancement to mitigate mosquito populations includes Boosted SIT, a version of conventional SIT that releases sterile males into the population that have the ability to release a biocide (e.g., pyriproxyfen) that further impairs the development of immature mosquito stages, leading to higher suppression rates (Bouyer et al., 2025). Field trials were done against Aedes aegypti in La Reunion and Aedes albopictus in Spain using the Boosted SIT technique, leading to higher suppression in comparison to Non-boosted SIT (Bouyer et al., 2025). Incompatible insect mosquito (IIT) is a potential alternative to the established Sterile Insect Techniques (Ritchie et al., 2018).Instead of relying on irradiation, this approach involves releasing male mosquitoes carrying Wolbachia, which-when they mate with uninfected wild females-cannot produce viable offspring (Beckmann et al., 2017; Laven, 1967).

Implications for vector control (microbiome)

Surveillance and Monitoring: The identification of specific microbial markers linked to insecticide resistance presents a valuable opportunity to enhance current resistance monitoring programs. By integrating microbiota profiling into surveillance systems, it may become possible to detect early signs of resistance development even before phenotypic resistance becomes apparent. This approach could complement traditional methods, providing a more sensitive and predictive tool for managing vector populations Strategic Control Measures: Understanding the microbiota’s role in modulating insecticide resistance opens the door to innovative vector control strategies. For example, interventions targeting or manipulating the microbial communities of vectors-such as through probiotics, microbiome engineering, or microbiota-disrupting agents-could potentially reverse resistance or increase susceptibility to insecticides. This microbiota-based approach represents a novel paradigm in integrated vector management, emphasizing the importance of host-microbiota interactions in resistance dynamics. This could lead to innovative control strategies that leverage the microbiota’s role in resistance (Pelloquin et al., 2021; Barnard et al., 2019). Understanding the gut microbiome’s contribution to resistance may open avenues for developing new insecticides or treatments that target these microbial communities, potentially reducing resistance development.

Gene drive technology

CRISPR-Cas9 presents a novel non reversible tool for mosquito vector control. Unlike the conventional mode of inheritance, where there is a 50% likelihood of genes being passed to offspring, gene drives bias a specific genetic trait to be inherited amongst nearly all progeny, allowing the rapid spread of desired modifications through wild population (Burt, 2003). It utilizes enzymatic cleavage of the target DNA at a specific site and copying the drive sequence into the cut site during DNA repair, a process known as homology-directed repair (Gantz and Bier, 2015). This system has been successfully demonstrated in Anopheles mosquitoes to spread genes that either suppress populations by targeting fertility genes or modify populations to render mosquito’s incapable of transmitting pathogens (Hammond et al., 2016; Kyrou et al., 2018).

As a proof of concept, targetting a doublesex (dsx) gene in Anopheles gambiae via gene drive technique led to population suppression in laboratory settings (Hammond et al., 2016). This concurs with a similar study wherein; disrupting dsx gene could lead to female sterility, effectively collapsing mosquito populations over successive generations (Kyrou et al., 2018).

Despite its promise, gene drive technology raises ecological, ethical, and regulatory concerns. Issues include potential off-target effects, the development of resistance alleles, and unintended ecological consequences if gene drives spread to non-target species. Therefore, WHO and other international bodies advocates for rigorous risk assessment, community engagement, and regulatory oversight before environmental releases.

Discussion

Insecticide resistance in mosquito vectors represents one of the most significant biological and operational threats to vector control programmes worldwide. Since the beginning of 21st century, resistance has rapidly spread across Anopheles, Aedes, and Culex species, undermining the effectiveness of indoor residual spraying (IRS), long-lasting insecticidal nets (LLINs), and space-spraying interventions. As a result, the tropical and subtropical countries such as India, where malaria, dengue, chikungunya, lymphatic filariasis, and other vector-borne diseases remain endemic. The global response to insecticide resistance over the past decade reflects an increasing alignment between mechanistic insights into resistance biology and evidence-based vector control policy.

Molecular and biochemical foundations of insecticide resistance

The culmination and synergism of biochemical, genetic, physiological, and behavioral adaptations have resulted in rise of resistance in mosquito populations. Among these, metabolic resistance is the most widespread and operationally consequential mechanism. Overexpression of detoxification enzymes enables mosquitoes to inactivate or sequester insecticides before they reach their neuronal targets. Cytochrome P450 monooxygenases (P450s) genes such as Cyp9m10 in Culex quinquefasciatus, Cyp6p3 and Cyp6z1 in Anopheles gambiae, and Cyp6p9a/b in Anopheles funestus have been reported to be strongly associated with pyrethroid resistance (Itokawa et al., 2010, 2011; Riveron et al., 2013a; Stevenson et al., 2012) (Figure 9).

Similarly, glutathione S-transferases (GSTs), particularly GSTe2, which catalyzes dehydrochlorination of DDT, also confer cross-resistance to pyrethroids (Riveron et al., 2014). While Carboxylesterases, such as estα2 and estβ2, are key drivers of sequestering or hydrolyzing organophosphate insecticides in Culex species (Bharati and Saha, 2018; S. H. P. P. Karunaratne et al., 1995; Mouchès et al., 1986).

Target-site resistance constitutes another major resistance axis. Knockdown resistance (kdr) mutations in the voltage-gated sodium channel, notably L1014F and L1014S, are widespread in An. gambiae, An. culicifacies, and Cx. pipiens which reduces sensitivity to both DDT and pyrethroids (Martinez‐Torres et al., 1998; Singh et al., 2009). Mutations in the ace-1 gene, such as G119S, decrease acetylcholinesterase sensitivity to organophosphates and carbamates and have been reported in An. gambiae, An. subpictus, and Cx. vishnui (Alout and Weill, 2008). Additional complexity due to GABA receptor mutations in the Rdl gene (A296S/G), confers resistance to cyclodienes and phenylpyrazoles in Aedes and Anopheles species (W. Du et al., 2005).

Emergence of novel resistance pathways

Recent research has broadened the resistance landscape beyond the traditional enzymatic and target-site models. ABC transporters, such as ABCG3, contribute to active efflux of pyrethroids from mosquito cells, reducing intracellular accumulation (Pignatelli et al., 2018) Uridine diphosphate glycosyltransferases (UGTs) have been implicated in phase-II detoxification in An. gambiae and An. funestus, highlighting additional metabolic layers of resistance (Logan et al., 2024). In Anopheles stephensi, the discovery of the Cub and Sushi Domain-containing Insecticide Resistance (CSDIR) protein, which enzymatically degrades deltamethrin and malathion extracellularly, represents a fundamentally new resistance paradigm (Kumar et al., 2024). Non-enzymatic sequestration mechanisms, such as SAP2-mediated pyrethroid binding in the legs of An. gambiae, further demonstrate how resistance can evolve outside classical detoxification pathways (Ingham et al., 2020). Sensory and signaling pathways, including opsin-mediated regulation of phospholipase C, have also been shown to modulate deltamethrin resistance in Culex pipiens pallens and Aedes albopictus (Li et al., 2018).

Another emerging frontier is the role of gut microbiota in resistance modulation. Comparative studies in An. albimanus and An. coluzzii reveal enrichment of insecticide-degrading bacterial taxa—such as Lysinibacillus, Stenotrophomonas, and Ochrobactrum—in resistant populations, suggesting microbial contributions to detoxification or host gene regulation (Barnard et al., 2019; Pelloquin et al., 2021).

Beyond these classical mechanisms, cuticular thickening or composition changes serve to reduce the insecticide penetration (Balabanidou et al., 2018; Jacobs et al., 2023). Furthermore, behavioral changes that alter feeding times, resting locations, or host-seeking behavior to avoid insecticide exposure have contributed to phenotypic resistance in mosquitoes. Of particular concern is the emergence of synergist resistance, where reduced susceptibility to piperonyl butoxide (PBO) compromises the effectiveness of PBO-based interventions (G. Zhou et al., 2022).

Since, PBO operates strictly as a metabolic inhibitor rather than a standalone adulticide, a decline in the operational efficacy of PBO-based interventions is inherently complex, polygenic, and multifactorial (G. Zhou et al., 2022). Rather than representing true toxicological resistance to the synergist molecule itself, this reduced net performance can be driven by a combination of compounding mechanisms (G. Zhou et al., 2022). These include the extreme over-expression of PBO-targeted cytochrome P450 monooxygenases (such as the CYP6P9 or CYP6P4 loci) that mechanically out-saturate available PBO concentrations, concurrent cuticular modifications that restrict the passive uptake of both molecules simultaneously, or a metabolic reliance on non-P450 pathways- such as carboxylesterases or glutathione S-transferases (GSTs)- which naturally evade PBO-mediated inhibition and preserve pyrethroid degradation capacity (G. Zhou et al., 2022).

Global policy responses informed by resistance mechanisms

Recognition of the biological complexity of insecticide resistance has driven a paradigm shift in vector control policy. The Global Plan for Insecticide Resistance Management in Malaria Vectors (GPIRM), launched in 2012, provided the first comprehensive framework linking resistance surveillance, insecticide rotation, and evidence-based decision-making (Mnzava et al., 2015) By the early 2020s, many malaria-endemic countries in Africa and Asia had incorporated GPIRM principles into national malaria strategic plans. However, the results have been sub-optimal due to significant resource and coordination hurdles in establishing Insecticide Resistance Monitoring and Management Plans (IRMMPs) and sentinel surveillance networks (Tabue et al., 2025).

Based on GPIRM, the Global Vector Control Response (GVCR) 2017–2030 institutionalized Integrated Vector Management (IVM) across malaria and arboviral disease programmes (World Health Assembly, 2017). GVCR emphasized reducing reliance on chemical insecticides, strengthening intersectoral coordination, and integrating biological, environmental, and community-based interventions. Countries such as India, Brazil, Indonesia, Kenya, Ghana, and Tanzania formally adopted GVCR-aligned strategies between 2017 and 2020 (WHO, 2020).

One of the most tangible outcomes of resistance-informed policy has been the deployment of next-generation LLINs. WHO’s 2017 interim recommendation for pyrethroid–PBO LLINs directly targeted metabolic resistance mediated by P450s. Large cluster-randomized trials in Tanzania and Burkina Faso demonstrated 20–40% reductions in malaria prevalence compared with standard pyrethroid LLINs in high-resistance settings (Sih et al, 2024). More recently, WHO’s 2023 recommendation of dual-active ingredient LLINs (pyrethroid–chlorfenapyr and pyrethroid–pyriproxyfen) provided compelling evidence that combining unrelated modes of action can restore vector control efficacy, with significant reductions in malaria incidence reported in West African trials (Barker et al., 2023).

Besides, the management of Aedes species has increasingly moved away from chemical escalation. Widespread resistance to temephos and pyrethroids prompted countries such as Brazil, India, Thailand, and Indonesia to prioritize source reduction, biological larvicides (Bti), insect growth regulators, and community engagement. These integrated approaches slow resistance evolution but require sustained operational commitment (Dusfour et al., 2019). The most striking resistance-independent success has been the Wolbachia-based population replacement strategy, implemented in Brazil and Indonesia since 2016. A large randomized trial in Indonesia demonstrated a 77% reduction in dengue incidence, underscoring the potential of non-insecticidal tools to circumvent resistance entirely (Utarini et al., 2021).

The Indian context: challenges and policy adaptation

India exemplifies the complexity of resistance management in a diverse ecological and epidemiological settings. Since the first reporting of resistance to DDT in An. culicifacies in Gujarat (1959), resistance has expanded extensively against multiple insecticide classes across diverse geo-climatic states (WHO, 2020). The country’s transmission dynamics are sustained by distinct primary malaria vectors, each expressing unique bionomic patterns and localized resistance profiles. Anopheles culicifacies serves as the principal rural vector, exhibiting deep-seated resistance to organochlorines (DDT) and rapidly spreading multi-insecticide resistance to organophosphates (malathion) and synthetic pyrethroids (Raghavendra et al., 2011; Singh et al., 2010). Concurrently, Anopheles stephensi acts as the primary urban malaria vector, presenting intense pyrethroid resistance and expanding its footprint due to rapid urbanization. In forested and sub-Himalayan regions, specialized vectors like Anopheles fluviatilis, Anopheles minimus, and Anopheles dirus present localized transmission dynamics that require targeted environmental management (Sharma, 1999).

To track this evolving landscape, national surveillance activities are systematically integrated through a collaborative network of sentinel stations overseen by the National Center for Vector Borne Diseases Control (NCVBDC) and the Indian Council of Medical Research-National Institute of Malaria Research (ICMR-NIMR). Routine monitoring utilizes standard WHO tube tests and CDC bottle bioassays alongside PCR-based molecular genotyping to detect target-site mutations, notably the L1014F, L1014S, and V1010L knockdown resistance (kdr) alleles in the voltage-gated sodium channels, as well as ace-1 modifications (Singh et al., 2009, 2010).

Persistent use of pyrethroids in IRS and LLINs has selected for kdr mutations in malaria vectors, while emerging pyrethroid resistance in Ae. aegypti and elevated esterase-mediated organophosphate resistance in Cx. quinquefasciatus threaten dengue and filariasis control. In response, India has adopted Integrated Vector Management under the National Vector Borne Disease Control Programme, emphasizing insecticide rotation, biolarvicides, and exploration of novel chemistries such as fluralaner as a potential novel mosquito-control compound (Shah et al., 2024). Operationally, resistance-management strategies implemented in India are pivoting away from legacy chemical protocols. Current frameworks enforce the strict rotational deployment of unrelated chemical classes for IRS, the distribution of next-generation dual-active LLINs to combat pyrethroid-resistant vector populations, and the use of chemical larvicides (e.g., temephos) in tandem with environmentally safe biological larvicides like Bacillus thuringiensis serotype israelensis (Bti) in aquatic habitats (NCVBDC, 2023). By linking real-time molecular surveillance with tailored regional interventions, India’s vector control policy aims to delay the selection of multi-resistant phenotypes and safeguard long-term elimination targets.

Conclusion and future perspectives

Insecticide resistance in mosquitoes is a dynamic and multifactorial phenomenon with profound implications for public health. It requires an understanding of localized resistance patterns and region-specific interventions, improved molecular surveillance, and the incorporation of new tools within integrated vector management frameworks. A deeper understanding of these resistance mechanisms, including their genetic and ecological underpinnings, is essential for designing sustainable, effective, and adaptable vector control strategies.

Correction note

This article has been corrected with minor changes. These changes do not impact the scientific content of the article.

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Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/fmala.2026.1890327/full#supplementary-material

Summary of the mechanisms of insecticide resistance in mosquitoes, illustrating metabolic detoxification, target-site insensitivity, and cuticular resistance pathways contributing to resistance evolution. Supplementary material at the source

Footnotes

Edited by: Emmanuel Elanga-Ndille, Centre for Research in Infectious Diseases (CRID), Cameroon

Reviewed by: Brice Natchema Soh Fonkou, Centre for Research in Infectious Diseases (CRID), Cameroon

Eric Tossou, International Institute of Tropical Agriculture (Benin), Benin