Dengue cases have risen sharply over the past decade, with global reports increasing nearly 29 times between 2000 and 2024 (WHO 2025a). Currently, about 5.6 billion people are at risk of dengue infection (Lim et al. 2025), and Aedes aegypti (Linnaeus) is the main vector. Vector control is still the main approach for preventing dengue (Ogunlade et al. 2023), and chemical insecticides are an essential component of these programs (Ranson & Lissenden 2016). However, using insecticides for long periods and on a large scale has put strong pressure on mosquito populations, causing them to develop resistance (Rahayu et al. 2018). There are four main ways mosquitoes become resistant: increased detoxification by metabolic enzymes (metabolic resistance), mutations in target sites that reduce sensitivity (target site resistance), changes in the cuticle that impede insecticide penetration (penetration resistance), and behaviors that help mosquitoes avoid treated surfaces (behavioral resistance) (Panini et al. 2016).

Indonesia provides favorable conditions for the spread of Ae. aegypti and is one of the countries with the highest rates of dengue (WHO 2025b). Pyrethroids are the most commonly used insecticides in Indonesia’s dengue control programs (Sayono et al. 2016), but resistance to these chemicals has been found in several Ae. aegypti populations (Zulfa et al. 2022). Because pyrethroids work by contact, it is important to study how the mosquito cuticle may contribute to penetration resistance. The insect cuticle is the main barrier to the entry of xenobiotics, including insecticides (Balabanidou et al. 2018; Ren et al. 2023), and changes like cuticle thickening can affect how well insecticides get through (WHO 2025b).

Previous studies have reported cuticle thickening associated with insecticide resistance in Ae. aegypti (Samal & Kumar 2021; Jacobs et al. 2023). However, to the best of our knowledge, information regarding this mechanism in Ae. aegypti populations from Indonesia remains limited. To address this gap, this study investigated cuticle thickening as a potential penetration resistance mechanism in Ae. aegypti population from Padang City, a region with multiple documented cases of insecticide resistance (Amelia-Yap et al. 2019; Hasmiwati & Supargiyono 2018; Hasmiwati et al. 2018; Rahayu & Fatimah 2020; Rahayu et al. 2022). The research was on Gunung Pangilun Village, where Ae. aegypti populations have been reported to exhibit resistance to the pyrethroid alpha-cypermethrin (Ardhiah et al. 2017) and the organophosphate temephos (Hasmiwati et al. 2018; Rahayu et al. 2021), as well as harbor mutations in the Ace-1 gene (Hasmiwati et al. 2018). This study was conducted to provide information on the potential contribution of cuticle thickening to insecticide resistance in this population, which could support dengue vector control strategies in the region.

Sampling was conducted from January to October 2020 in Gunung Pangilun Village, Padang, West Sumatra, Indonesia (0°55’00.8” S, 100°21’39.2” E) using purposive sampling. Rearing and specimen preparation prior to scanning electron microscopy (SEM) were performed in the Animal Physiology Laboratory, Department of Biology, Faculty of Mathematics and Natural Sciences, Universitas Andalas. Observations using SEM were conducted at the Biology Research Center, Indonesian Institute of Sciences (LIPI; now integrated into BRIN), Gedung Widya Satwaloka, Cibinong, West Java, Indonesia.

Mosquito eggs were collected using egg traps (ovitraps) placed in ten residential houses, with three ovitraps installed in each house. Ovitraps were made from plastic cups (~9 cm top diameter and ~10 cm height) coated with thick black plastic. A strip of Whatman paper (approximately 6 cm × 10 cm) was placed inside each cup and attached to the inner wall. The cups were filled with tap water to approximately one-quarter of their volume, allowing the paper to remain partially submerged while the upper portion remained exposed to air.

Egg traps were placed in a shady or dimly lit area indoors to minimize the risk of disturbance. The traps were left in place for one week, after which they were collected and replaced with other ovitraps containing fresh Whatman paper and tap water. This procedure was repeated weekly until sufficient egg samples were obtained.

^Eggs attached to the Whatman paper were gently brushed using a fine brush into plastic containers (30 cm × 50 cm × 12 cm) filled with tap water to approximately two-thirds of the container volume. The containers were covered with mesh to maintain hygiene and prevent contamination. Larvae of all instars were maintained in the same rearing container. Water was replaced as needed based on visual inspection to maintain cleanliness.

Pupae were transferred to plastic cups (~9 cm top diameter and ~10 cm height) using a pipette and covered with mesh to prevent the escape of emerging adults. Emerging adults were maintained until five days post-emergence prior to use in bioassays. Larvae were fed finely ground commercial chicken feed, while adult mosquitoes were provided with a 10% sucrose solution via cotton pads placed on top of the mesh covering the cups. Rearing was conducted at and 70–80% relative humidity (RH).

Female Ae. aegypti mosquitoes aged 5 days were used in this study. Species identification was performed based on the presence of a lyre-shaped scale pattern on the dorsal thorax, and sex differentiation was determined by antennal morphology (Christophers 1960).

Screening bioassays using a WHO-based bioassay protocol (WHO 2016) were conducted for phenotypic classification of resistant and susceptible mosquitoes using impregnated paper with 0.025% alphacypermethrin for exposure. Adult female mosquitoes were tested in batches of up to 20 individuals per exposure, depending on availability. Mosquitoes were introduced into holding tubes lined with untreated paper (12 cm × 15 cm) by aspirator and maintained vertically for 1 h under standard test conditions. The exposure tube containing the insecticide-impregnated paper was then connected to the holding tube, and mosquitoes in the holding tube were transferred to the exposure tube by opening the slide unit and gently blowing until all mosquitoes moved into the exposure tube. After 1 hour of exposure, the mosquitoes were returned to the holding tube by opening the slide unit and were provided with a 10% sucrose solution via cotton pads for a 24-hour recovery period.

After the 24 h post-exposure, surviving mosquitoes were classified as resistant, while those that died were classified as susceptible. No independent biological replicates were performed, and individuals were collected from multiple exposure batches until the required number of samples was obtained.

Ten individuals per group were used for scanning electron microscopy (SEM) analysis. The resistant group (n = 10) consisted of mosquitoes that survived 24 h after insecticide exposure using WHO bioassay, while 10 individuals from the susceptible group consisted of six field-collected mosquitoes that died during the WHO bioassay and four individuals from a laboratory susceptible strain (VCRU-WHO, Universiti Sains Malaysia). The limited number of laboratory strain individuals was due to low emergence success during colony maintenance. All susceptible individuals, both from the field and laboratory strains, were analyzed collectively as the susceptible group in this s tudy .

Mosquitoes were processed immediately after classification. Resistant individuals were euthanized by placing them in a freezer for several minutes prior to dissection. Next, the left middle leg was carefully detached from the mosquito body and gently rubbed with a fingertip to remove scales on the leg surface, particularly along the tibia. The leg was then placed on a glass slide and rinsed with 70% ethanol, and the midtibia of Ae. aegypti was transversely sectioned using a platinum-coated razor blade (Figure 1). The excised leg segment was transferred into a 1.5 ml microcentrifuge tube containing 70% ethanol and stored until scanning electron microscopy (SEM) analysis. The mid-tibia of the left middle leg was selected to ensure that the same anatomical region was examined consistently across all specimens, thereby providing a consistent basis for comparison among specimens. Previous studies have assessed cuticular thickness or cuticular resistance at different anatomical locations, including the mid-tibia, as in the study of Lilly et al. (2016) in Cimex lectularius Linnaeus (Table 2).

Sample preparation for SEM was performed using a modified protocol based on Goldstein et al. (1992). Specimens were immersed in cacodylate buffer for approximately 2 h, followed by ultrasonic agitation for 5 min. Then, samples were fixed in 2.5% glutaraldehyde for 24 h and were immersed in 2% tannic acid for 6 h, followed by rinsing in cacodylate buffer for 5 min, repeated four times. Dehydration was performed using a graded ethanol series: 50% ethanol for 5 min (four times), followed by 70%, 85%, and 95% ethanol for 20 min each at room temperature. Samples were subsequently immersed in absolute ethanol for 10 min, repeated twice. Specimens were then frozen in tertbutanol for 10 min (twice), stored in a freezer, and dried using a vacuum dryer until completely dry.

Specimens were mounted and sputter-coated with gold at 5–8 mA for 5 min using an ion coater (Eiko IB-2), followed by vacuum treatment for 15 min. Observations were performed using a JSM-5000LV SEM. SEM imaging was conducted at an accelerating voltage of 3.0 kV, working distance of 11.7–13.2 mm, and under highvacuum conditions. Images used for measurements were acquired at magnifications ranging from 1000× to 3000×.

Cuticle thickness was measured from SEM micrographs at the mid-tibia using ImageJ software (version 1.52p). Measurements were not performed under blinded conditions. However, identical measurement criteria were applied to all specimens to reduce potential measurement bias. Cuticle thickness was determined by delineating the outer and inner boundaries of the mosquito leg cuticle. Regions that do not have clearly defined boundaries, such as areas affected by structural damage, debris, or cuticle derivatives, were excluded from the measurement. This approach is similar to that used by Wood et al. (2010) for SEM-based measurement of leg cuticle thickness in Anopheles funestus Giles. Because different magnifications were used, each micrograph was independently calibrated using its corresponding scale bar prior to measurement.

For each mosquito, 25 measurement points were recorded. The mean cuticle thickness per mosquito was calculated as the average of these measurements and used for statistical analysis. The individual mosquito was treated as the unit of analysis. Data normality and homogeneity of variance were assessed using the Shapiro–Wilk and Levene’s tests, respectively. Differences in mean cuticle thickness between resistant and susceptible groups were evaluated using an independent-samples t-test. Statistical analyses were performed using R software (version 4.5.2) (R Core Team 2025).

The detailed records of the total number of individuals tested and classified in each bioassay were not retained, since the WHO bioassay was conducted as part of a phenotypic selection procedure to obtain individuals representing both susceptible and resistant phenotypes for subsequent cuticle thickness analysis. The distribution of cuticle thickness measurements across 25 points is shown in Figure 2. The mean cuticle thickness of the alpha-cypermethrin-resistant Ae. aegypti population was significantly higher (1.28-fold) than that of the susceptible population (Table 1; Figure 3). For comparison, four susceptible individuals of the laboratory strain had a mean cuticle thickness of 1.949 µm, whereas six susceptible individuals from the field had a mean cuticle thickness of 1.885 µm. Thus, the descriptive difference between the laboratory-strain and field-susceptible groups was small.

Statistical analysis indicated that the data were normally distributed based on the Shapiro–Wilk test for both the susceptible and resistant groups. Homogeneity of variance was confirmed by Levene’s test An independent-samples t-test revealed a significant difference in tibial cuticle thickness between groups , with a mean difference of . The effect size was large (Hedges’

This study indicates an association between pyrethroid resistance and increased cuticle thickness in the Ae. aegypti population from Gunung Pangilun. Similar observations have been reported in pyrethroid-resistant insects across multiple species, life stages, and body regions, where resistant individuals exhibited thicker cuticles than susceptible ones, with differences of up to 2.3-fold (Table 2). In addition, Table 2 shows that cuticle thickening associated with insecticide resistance has been reported not only in adult insects but also in immature developmental stages, including larvae and nymphs. This pattern suggests that differences in cuticle thickness associated with resistance are already present even before adulthood.

As contact insecticides, pyrethroids must penetrate the insect cuticle before reaching their neural targets, where the cuticle serves as the primary physical interface regulating insecticide entry (Ren et al. 2023). Therefore, increased cuticle thickness likely acts as a kinetic barrier that slows pyrethroid penetration, effectively extending the time available for detoxification systems to metabolize and neutralize the toxic compound (Pedrini et al. 2009; Balabanidou et al. 2018). Rather than acting alone, penetration resistance may function as a co-resistance mechanism that enhances the effectiveness of other resistance mechanisms, including elevated activity of detoxification enzymes (esterases, cytochrome P450 monooxygenases, and glutathione S-transferases) and target site mutations in voltagegated sodium channels (VGSCs) (Liu 2015), as well as behavioral avoidance responses that reduce effective exposure time (Yang et al. 2019; Sukkanon et al. 2020).

Comparative bioassay studies further support the functional role of the cuticle as a barrier. The increased susceptibility observed in injection assays, where insecticides are injected directly into the hemocoel, bypassing the cuticle, compared with topical applications that must pass through the cuticle, suggests the cuticle’s involvement in limiting insecticide penetration (Wood et al. 2010; Meng et al. 2023; Wang et al. 2025). These findings strengthen the interpretation that cuticular modifications contribute to reduced insecticide uptake.

The insect cuticle is structurally composed of two main, compositionally distinct layers: the epicuticle (outer layer) and the procuticle (inner layer). The epicuticle contains lipoproteins, fatty acids, and a waxy layer enriched with cuticular hydrocarbons (CHCs), while the procuticle is predominantly composed of chitin and structural proteins (Gillot 2005; Balabanidou et al. 2016). Cuticular thickening can occur in one or both layers (Jacobs et al. 2023; Li et al. 2025), suggesting multiple structural pathways that may delay insecticide uptake.

Alterations in the epicuticle (the outer layer of the cuticle), particularly increases in cuticular hydrocarbons, have been reported in resistant insects. Pyrethroid-resistant populations have been shown to possess higher levels of CHCs, a major component of the epicuticle (Balabanidou et al. 2016; Simma et al. 2019; Yahouédo et al. 2017). Increased CHC content can form a chemical barrier that reduces the diffusion of lipophilic compounds across the insect surface (Koganemaru 2015). A thicker epicuticle has been observed in Anopheles gambiae Giles (Balabanidou et al. 2016), where increased CHC levels have been correlated with reduced pyrethroid uptake (Yahouédo et al. 2017). Those findings suggest that both structural thickening and enrichment of cuticular hydrocarbons may contribute to penetration resistance.

Penetration resistance also involves transcriptional changes in cuticle-related genes. Structural proteins belonging to the CPR, CPAP, and CPF families, as well as genes involved in chitin synthesis such as chitin synthase (CHS), have been shown to contribute to both cuticle thickening and structural modifications (Fang et al. 2015; Huang et al. 2018; Balabanidou et al. 2019; Guo et al. 2019; Wang et al. 2025; Li et al. 2025; Xu et al. 2022; Sun et al. 2017). Upregulation of the CYP4G16 gene, which is involved in hydrocarbon biosynthesis and directly linked to increased cuticular hydrocarbon production, has also been reported (Balabanidou et al. 2016; Yahouédo et al. 2017; Balabanidou et al. 2019). Additionally, other studies have also found overexpression of ABC transporters, which facilitate the redistribution of cuticle components, thus supporting cuticle reinforcement (Matowo et al. 2014; Xu et al. 2023; Kefi et al. 2023). Together, these transcriptional changes in multiple cuticle-related genes indicate that cuticle thickening represents a regulated physiological adaptation contributing to resistance.

Furthermore, functional studies also provide direct evidence for the role of cuticle structure in resistance. Silencing of cuticle protein genes such as BgCPLCP1 has been shown to reduce cuticle thickness and restore susceptibility to β-cypermethrin in Blattella germanica (Linnaeus) (Cai et al. 2024). Several additional mechanisms affecting cuticle physical properties have also been linked to resistance-related traits. Alterations in pigmentation pathways, such as increased YELLOW-E gene expression, have been associated with increased cuticle tanning, which may indirectly contribute to penetration resistance through changes in cuticle rigidity and permeability (Simma et al. 2019). Furthermore, a denser chitin layer in the endocuticle has been observed in resistant Bactrocera dorsalis (Hendel) strains compared with susceptible strains, suggesting increased cuticle compaction (Lin et al. 2012). These findings are further supported by the overexpression of cuticle-related genes in leg regions, which are major sites of insecticide contact (Huang et al. 2018; Xu et al. 2020; Kefi et al. 2023; Li et al. 2025).

Taken together, previous studies have shown that several cuticle-related modifications may contribute to reduced insecticide penetration. The thicker cuticles in the resistant population found in this study support the possibility that structural changes in the cuticle play a role in penetration resistance.

However, this study is limited by its relatively small sample size, which reduces statistical generalizability. Additionally, the inclusion of field-collected mosquitoes in the susceptible group may contribute to biological variability compared with fully standardized laboratory strains. Further studies incorporating larger sample sizes, standardized susceptible reference strains, and complementary analyses such as molecular and biochemical assays are needed to validate and extend our findings.

Collectively, this study provides preliminary evidence of cuticular involvement in pyrethroidresistant Ae. aegypti from Gunung Pangilun Village, Padang. Our findings provide the basis for further comprehensive investigations to elucidate the resistance mechanisms and assess their implications for dengue vector control strategies.

This study provides evidence suggesting cuticle thickening as a potential resistance mechanism in pyrethroid-resistant Ae. aegypti from Gunung Pangilun Village, Padang, West Sumatra. To the best of our knowledge, this is the first report describing cuticle thickening associated with pyrethroid resistance in Ae. aegypti in Indonesia. These findings contribute to the understanding of insecticide resistance mechanisms and may support the development of more effective resistance management strategies for controlling this primary dengue vector.