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Article

Biochemical Effects of Petroselinum crispum (Umbellifereae) Essential Oil on the Pyrethroid Resistant Strains of Aedes aegypti (Diptera: Culicidae)

by
Jitrawadee Intirach
1,2,
Anuluck Junkum
1,*,
Nongkran Lumjuan
3,
Udom Chaithong
1,
Pradya Somboon
1,
Atchariya Jitpakdi
1,
Doungrat Riyong
1,
Danita Champakaew
1,2,
Roongtawan Muangmoon
1,2,
Arpaporn Chansang
1,2 and
Benjawan Pitasawat
1
1
Center of Insect Vector Study, Department of Parasitology, Faculty of Medicine, Chiang Mai University, Chiang Mai 50200, Thailand
2
Graduate PhD’s Degree Program in Parasitology, Faculty of Medicine, Chiang Mai University, Chiang Mai 50200, Thailand
3
Research Institute for Health Sciences, Chiang Mai University, Chiang Mai 50200, Thailand
*
Author to whom correspondence should be addressed.
Insects 2019, 10(1), 1; https://doi.org/10.3390/insects10010001
Submission received: 22 November 2018 / Revised: 17 December 2018 / Accepted: 18 December 2018 / Published: 24 December 2018
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Abstract

:
In ongoing screening research for edible plants, Petroselinum crispum essential oil was considered as a potential bioinsecticide with proven antimosquito activity against both the pyrethroid susceptible and resistant strains of Aedes aegypti. Due to the comparative mosquitocidal efficacy on these mosquitoes, this plant essential oil is promoted as an attractive candidate for further study in monitoring resistance of mosquito vectors. Therefore, the aim of this study was to evaluate the impact of P. crispum essential oil on the biochemical characteristics of the target mosquito larvae of Ae. aegypti, by determining quantitative changes of key enzymes responsible for xenobiotic detoxification, including glutathione-S-transferases (GSTs), α- and β-esterases (α-/β-ESTs), acetylcholinesterase (AChE), acid and alkaline phosphatases (ACP and ALP) and mixed-function oxidases (MFO). Three populations of Ae. aegypti, comprising the pyrethroid susceptible Muang Chiang Mai-susceptible (MCM-S) strain and the pyrethroid resistant Pang Mai Dang-resistant (PMD-R) and Upakut-resistant (UPK-R) strains, were used as test organisms. Biochemical study of Ae. aegypti larvae prior to treatment with P. crispum essential oil revealed that apart from AChE, the baseline activity of most defensive enzymes, such as GSTs, α-/β-ESTs, ACP, ALP and MFO, in resistant UPK-R or PMD-R, was higher than that determined in susceptible MCM-S. However, after 24-h exposure to P. crispum essential oil, the pyrethroid susceptible and resistant Ae. aegypti showed similarity in biochemical features, with alterations of enzyme activity in the treated larvae, as compared to the controls. An increase in the activity levels of GSTs, α-/β-ESTs, ACP and ALP was recorded in all strains of P. crispum oil-treated Ae. aegypti larvae, whereas MFO and AChE activity in these mosquitoes was decreased. The recognizable larvicidal capability on pyrethroid resistant Ae. aegypti, and the inhibitory effect on AChE and MFO, emphasized the potential of P. crispum essential oil as an attractive alternative application for management of mosquito resistance in current and future control programs.

1. Introduction

The mosquito, Aedes (Stegomyia) aegypti (Linnaeus, 1762), is an important vector for transmitting life-threatening human diseases like dengue, chikungunya, Zika, and yellow fever in tropical and subtropical regions worldwide [1,2,3]. Dengue is the World’s most critical mosquito-borne viral disease, which inflicts millions of deaths each year. Approximately 3.9 billion people, in 128 countries, are currently at risk of dengue infection, with an estimated 50–100 million and 250,000–500,000 cases of dengue fever and dengue hemorrhagic fever, respectively [4,5]. The number of dengue cases reported annually to the World Health Organization (WHO) has increased from 0.4 to 1.3 million in the decade 1996 to 2005, reaching 2.2, 3.2, and 3.8 million in 2010, 2015, and 2016, respectively [2,6,7]. Since several dengue vaccine candidates are currently in the developmental stage, there is no specific treatment or vaccine commercially available. Therefore, personal protection, environmental management, and mosquito control remain the most important tools in preventing and controlling transmission of this disease [5,8,9,10].
Management of mosquito vectors by applying appropriate chemical compounds such as larvicides, adulticides, attractants, deterrents, and repellents is an essential element to minimize disease transmission [11,12,13]. At least four main classes of synthetic insecticides; namely organochlorines (exclusively DDT), organophosphates, carbamates, and pyrethroids have been in use over decades in vector control programs. In addition to low mammalian toxicity, synthetic pyrethroids have become the most popular and prevalent active ingredients in public health vector control programs, due to their high invertebrate potency at low levels, resulting in rapid immobilization (‘knockdown’) and killing of mosquitoes [14,15]. Although synthetic substances that dramatically reduce the risk of vector-borne diseases have been documented, the overuse and misuse of conventional chemicals, for example, pyrethroids and other insecticides, have led to mosquito resistance, which threatens the potentiality of vector control [16,17,18,19]. A recommended approach for monitoring and managing insecticide resistance in mosquito populations is to search for alternative interventions in order to restrain or reduce the evolution of further resistance as well as preserve the efficiency of existing insecticides [19,20]. Use of biopesticides is encouraged as a promising economic and eco-friendly strategy for fighting the resistance problem [21,22]. Plant essential oils with insecticidal activity have been developed as pesticides for many years, and some have been used commercially as pesticides or repellents against garden and household pests. However, their use as mosquito larvicides is limited, due to their relatively low toxicity against mosquito larvae [22,23,24]. In consequence, research and development of new natural alternatives have been designed and performed by focusing on the improvement of their biological activity [25,26,27]. A literature survey revealed that there are several studies elucidating the behavioral responses or biochemical constituents of target mosquitoes, which could enable a better understanding of plant essential oil effects and their possible mode of action [28,29,30].
As part of the ongoing screening research for local edible plants in Thailand, Petroselinum crispum fruit oil was considered as a potential bioinsecticide, with proven antimosquito activity against both pyrethroid susceptible and resistant strains of Ae. aegypti [31]. The larvicidal potential of P. crispum oil is effective, with an insignificant difference of LC50 values of 43.22, 44.50 and 44.03 ppm against Muang Chiang Mai-susceptible (MCM-S), Pang Mai Dang-resistant (PMD-R), and Upakut-resistant (UPK-R) strains of Ae. aegypti, respectively. The GC-MS analysis of P. crispum oil, which is an identical sample used in this study, revealed the presence of 19 bioactive compounds, accounting for 98.25% of the whole oil, with the principal constituents being thymol (42.41%), p-cymene (27.71%) and γ-terpinene (20.98%) [31]. Despite P. crispum oil producing less larvicidal potential than synthetic insecticides such as temephos, permethrin, and deltamethrin; the comparative efficacy on pyrethroid susceptible and resistant Ae. aegypti makes it an attractive candidate for further research on monitoring resistance in mosquito vectors. As the essential oil is a mixture of bioactive materials that functions through the different modes of actions [32,33], P. crispum oil potentially reduces the risk of resistance development in mosquitoes. The literature, however, offers no data about the insect detoxification mechanisms and biochemical responses to this plant essential oil.
Major mechanisms of pyrethroid resistance in insects involve mutation within the target site of the insecticide and/or increase in the rate of insecticide detoxification. Metabolic detoxification of insecticides is associated with alterations in the activity of defensive enzymes such as glutathione-S-transferases (GSTs), α- and β-esterases (α-/β-ESTs), acetylcholinesterase (AChE), acid and alkaline phosphatases (ACP and ALP), and mixed-function oxidases (MFO), which help organisms to transform and/or eliminate endogenous and exogenous compounds [34,35,36,37]. An understanding of biochemical changes to plant-derived product, such as P. crispum essential oil, plays a vital role in vector management, especially resolution of insecticide resistance. Therefore, the aim of this study was to evaluate the biochemical effects of the P. crispum essential oil caused in Ae. aegypti strains. Quantitative changes of important marker enzymes responsible for xenobiotic detoxification, including GSTs, α-/β-ESTs, AChE, ACP, ALP, and MFO, were determined in either the susceptible or resistant strain of Ae. aegypti larvae treated with P. crispum essential oil. The information obtained from this study has enhanced basic knowledge and useful guidelines on how to use and improve plant products for future management of vector control in areas with high levels of pyrethroid resistance.

2. Materials and Methods

2.1. Preparation of P. crispum Oil

Dried fruit of P. crispum was obtained commercially from medicinal plant suppliers in Chiang Mai province, Thailand. Scientific identification of this plant specimen was accomplished by Miss Wannaree Charoensup, a scientist at the Department of Pharmaceutical Science, Faculty of Pharmacy, Chiang Mai University (CMU). A reference voucher, PARA-PE-001-Fr/1, was deposited at the Department of Parasitology, Faculty of Medicine, CMU. The identified P. crispum fruit was shade-dried for one week in an open area, with active ventilation and ambient temperature of about 30 ± 5 °C, in order to remove the moisture content before extracting essential oil. After grinding, the fruit powder of P. crispum was subjected to extractions by steam distillation at 100 °C for at least 3 h. The essential oil obtained was collected, dried over anhydrous sodium sulfate (Na2SO4) to make it moisture free, and preserved in an airtight brown bottle at 4 °C in a refrigerator until its use in experimental bioassays.

2.2. Chemicals and Reagents

The albumin standard Thermo Fisher Scientific (Waltham, MA, USA) and Bio Rad protein reagent (Bio-Rad Laboratories, Bercules, CA, USA) were used to determine total protein. Cytochrome c from bovine heart (≥95%) (Sigma-Aldrich, St. Louis, MO, USA), α-naphthol (≥99%) (Sigma-Aldrich, Buchs, SG, Switzerland), β-naphthol (99%) (Sigma-Aldrich, Shanghai, China), and p-nitrophenol (99%) (Sigma-Aldrich, St. Louis, MO, USA) were used in the biochemical assays of MFO, α-/β-ESTs, and ACP/ALP, respectively. The substrates of enzyme analysis, including 3,3′,5,5′-tetramethylbenzidine dihydrochloride hydrate powder (≥98%) (Sigma-Aldrich, Buchs, SG, Switzerland), α-naphthyl acetate (≥98%) (Sigma-Aldrich, Buchs, SG, Switzerland), β-naphthyl acetate (98%) (Sigma-Aldrich, Buchs, SG, Switzerland), 1-chloro-2, 4-dinitrobenzene (minimum 98%) (Sigma-Aldrich, St. Louis, MO, USA), L-glutathione reduced (minimum 98%) (Sigma-Aldrich, Tokyo, Japan), 5,5′-Dithiobis (2-nitrobenzoic acid) (≥98%) (Sigma-Aldrich, St. Louis, MO, USA), acetylthiocholine iodide (≥98%) (Sigma-Aldrich, Gillingham, Kent, UK), and p-nitrophenyl phosphate (≥97%) (Sigma-Aldrich, Buchs, SG, Switzerland), were dissolved in a suitable solvent such as absolute ethanol, methanol and phosphate buffer. Fast Blue B Salt-Dye content (~95%) was purchased from Sigma-Aldrich, St. Louis, MO, USA. All other chemicals and reagents used were of analytical grade and procured from local agencies in Chiang Mai province, Thailand.

2.3. Mosquito Test Population and Colonization

The three mosquito populations used in this study were pyrethroid-susceptible and resistant strains of Ae. aegypti, comprising Muang Chiang Mai-susceptible (MCM-S), Pang Mai Dang-resistant (PMD-R), and Upakut-resistant (UPK-R), which were established from field larvae collected from clean stagnant water at various breeding sites in Chiang Mai province: Muang Chiang Mai district in 1995 [38], Ban Pang Mai Dang, Mae Tang district in 1997 [39], and Upakut Temple, Muang Chiang Mai district in 2006 [40], respectively. The MCM-S was resistant to DDT, but susceptible to pyrethroids such as permethrin and deltamethrin [31]. The PMD-R strain was resistant to both DDT and permethrin, but susceptible to deltamethrin [39,41,42,43], whereas the UPK-R strain was resistant to DDT, permethrin, and deltamethrin [40]. The PMD-R and UPK-R strains were reared under selection pressure by regular exposure to the WHO [44] discriminating dose (0.75% permethrin and 0.05% deltamethrin, respectively) in order to maintain the resistance level. All of the mosquitoes were maintained continually from their dates of collection in a climate-controlled insectary (25 ± 2 °C, 80 ± 10% Relative Humidity, and 14:10 h L/D photoperiod cycle) at the Department of Parasitology, Faculty of Medicine, CMU. The mosquito larvae were reared in plastic trays with tap water and fed on finely ground dog biscuits twice daily. The adults reared in humidified cages had continual access to 10% sucrose solution containing 10% multivitamin syrup, while females were blood-fed for egg production. Early 4th stage Ae. aegypti larvae of each strain were available for biochemical assays.

2.4. Determination of Lethal Threshold Time for the Lethal Effect of P. crispum Oil

Larvicidal bioassays, according to a slightly modified version of the WHO standard protocol [45], were performed to determine the threshold time for lethal effect of P. crispum oil on the three strains of Ae. aegypti under laboratory conditions. Four batches of 25 newly hatched 4th instar larvae were exposed to P. crispum essential oil solution at the median lethal concentrations (LC50) of 43.22, 44.50 and 44.03 ppm for MCM-S, PMD-R, and UPK-R strains, respectively, as estimated in an earlier study [31]. The control group comprised the larvae maintained in 0.4% DMSO-distilled water solution, whereas the untreated group was maintained in distilled water only. No food was provided for the larvae during the experiment. Bioassays were carried out under controlled conditions at 25 ± 2 °C and 80 ± 10% Relative Humidity, with four replicates. The lethal threshold time points for larvae were determined after recording the percentage of their mortality at time-intervals up until 24 h (0, 3, 6, 12, and 24 h). Exposure times that yielded larval mortality of 20–50% were considered as the threshold time for the lethal effect of P. crispum essential oil.

2.5. Preparation of Whole Body Homogenates Used for Enzyme Assay

After exposure to the test solutions at time-intervals up until 24 h (0, 3, 6 and 24 h), the larvae of treated, untreated and control groups obtained from each time-interval were washed with deionized water. These larvae were blotted with tissue paper for complete removal of adhered water and then kept in 1.5 mL tubes at −80 °C until their use for enzyme analysis. In order to prepare whole body homogenates, the stored larvae (2–10 individuals) of treated, untreated and control groups of each mosquito strain were homogenized separately in suitable buffer required for each enzyme assay. The whole body homogenates were centrifuged at 10,000× g at 4 °C for 20 min and the clear supernatants were used immediately for the determination of enzyme activity. All enzyme bioassays in this study were performed together with negative controls in the presence of homogenate buffer, but not enzyme, in the reactions.

2.6. Total Protein Assay

Total protein content in each larval homogenate was estimated according to the method of Bradford [46], with some minor modifications. Two hundred microliters of BIO Rad protein reagent diluted 1:4 in buffer (distilled water) were added to 10 µL of crude larval homogenate. The mixture was incubated in a 96-well microtiter plate at room temperature. Absorbance reading was taken at 595 nm after 5 min of reaction against homogenate buffer as a blank. Protein concentrations in mg/mL were calculated from a standard curve of bovine serum albumin (0–0.5 mg/mL).

2.7. Glutathione S-Transferases (GSTs) Assay

GSTs activity was determined according to the modified method of Habig et al. [47]. Ten live larvae were homogenized in 200 µL of 0.1 M potassium phosphate buffer, pH 6.5. Two hundred microliters of GSH/CDNB working solution (10 mM reduced glutathione prepared in 0.1 M potassium phosphate buffer, pH 6.5 and 3 mM chlorodinitrobenzene diluted in methanol) were added to 10 µL of larval homogenate. Enzyme activity was measured at 340 nm, with the optimum time for reading of 2 min at room temperature. The GSTs activity was expressed as µmol CDNB conjugated/min/mg protein.

2.8. α-/β-Esterases (α-/β-ESTs) Assays

The activity of α- and β-ESTs was determined according to the methodology of van Asperen [48], with slight modifications. Two live larvae were homogenized in 1 mL of 20 mM potassium phosphate buffer, pH 7.2. The larval homogenates were diluted with homogenization buffer at the ratio of 1:4 before measuring. Two hundred microliters of α- or β-naphthyl acetate solution (200 µL of 30 mM α- or β-NA in ethanol in 20 mL of 20 mM potassium phosphate buffer, pH 7.2) were added to 20 µL of larval homogenate. The mixture was incubated in a 96-well microtiter plate for 30 min at room temperature. The enzymatic reaction was stopped by the addition of 50 µL of Fast Blue B stain solution (22.5 mg of Fast Blue in 2.25 mL of distilled water, then 5.25 mL of 5% sodium lauryl sulphate diluted in distilled water). Replicate blanks contained 20 µL of homogenization buffer, 200 µL of substrate solution, and 50 µL of stop solution. Enzyme activity was read at 570 nm as an end point. The activities of α-/β-ESTs were calculated from the α- or β-naphthyl acetate standard curve. The activity of α-/β-ESTs was expressed as nmol of the α- or β-naphthol released/min/mg protein.

2.9. Acetylcholinesterase (AChE) Assay

Evaluation of AChE activity was performed by following the modified method of Ellman et al. [49]. Five live larvae were homogenized in 250 µL of 0.1 M potassium phosphate buffer, pH 7.0 with 1 µM DL-dithiothreitol and 1% Triton X-100. An aliquot of 25 µL of larval homogenates was mixed successively with 155 µL of 0.65 mM dithiobis 2-nitrobenzoic acid solution and 25 µL of 10 mM acetylthiocholine iodide in a microliter plate. The absorbance reading was monitored at 405 nm after incubation for 5 min at room temperature. The AChE activity was reported as nmol of the acetylthiocholine hydrolyzed/min/mg protein.

2.10. Acid and Alkaline Phosphatases (ACP and ALP) Assays

The levels of ACP and ALP were measured by following the method of Asakura [50], with slight modifications. Ten live larvae were homogenized in 500 µL of appropriate homogenization buffer, which comprised 50 mM sodium acetate buffer, pH 4.0 and 50 mM Tris-HCl buffer, pH of 8.0, with 1 mM DL-dithiothreitol for the ACP and ALP assay, respectively. ACP activity was analyzed by mixing 10 µL of larval homogenate with 200 µL of substrate mixture containing 25 mM sodium acetate buffer, pH 4.0 and 6.25 mM p-nitrophenyl phosphate. For estimating ALP activity, buffer in substrate mixture also was substituted by 50 mM Tris-HCl buffer, pH 8.0. After incubation at 37 °C for 15 min in 96-well microtiter plates, the enzymatic reactions were terminated by adding 50 µL of 0.5 N NaOH. The absorbance was read at 405 nm and compared with the standard curve of known p-nitrophenol concentrations (0.0625–4 µg/µL). ACP/ALP activity was reported as nmol of the p-nitrophenol released/min/mg protein.

2.11. Mixed-Function Oxidases (MFO) Assay

MFO activity was determined by using the heme-peroxidase assay of Brogdon et al. [51], with minor modifications. Ten live larvae were homogenized in 700 µL of 0.25 M sodium acetate buffer, pH 5.0. One hundred microliters of larval homogenate were mixed with 200 μL of substrate mixture and 6.3 mM TMBZ solution (0.01 g of 3,3′,5,5′-tetramethylbenzidine in 5 mL of absolute methanol mixed with 15 mL of 0.25 M sodium acetate buffer, pH 5.0, freshly prepared daily), in a 96-well microtiter plate. Finally, 25 μL of 3% H2O2 were added. The plate was incubated for 5 min at room temperature and then read at 630 nm by using a microplate reader (Spectra MR, DYNEX technologies, Chantilly, VA, USA). MFO values were compared with known concentrations of cytochrome c from horse heart type VI (0–1.4 ng/µL) and expressed as nmol of cytochrome c equivalent unit/mg protein.

2.12. Statistical Analysis

Data from biochemical assays were subjected to analysis of variance (One-way ANOVA) and expressed as a mean of 6–10 replicates using samples from different preparations. Significant differences between treatment groups among the three strains of mosquitoes were analyzed by Tukey’s HSD test (p < 0.05) using IBM SPSS statistics 24 (IBM Corp., Armonk, NY, USA).

3. Results

3.1. Enzyme Activity Levels in the Three Strains of Ae. aegypti Prior to Treatment (0 h Time Point)

Prior to treatment with P. crispum oil, the baseline activity of some detoxifying enzymes, such as GSTs (F = 867.722, df = 2, p < 0.0001), α-EST (F = 74.345, df = 2, p < 0.0001), β-EST (F = 78.497, df = 2, p < 0.0001) and ACP (F = 82.421, df = 2, p < 0.0001) in resistant PMD-R and UPK-R larvae, was found to be significantly higher than that in susceptible MCM-S (Figure 1 and Table 1). A significant difference in highest GSTs and β-EST activity was recorded in UPK-R and PMD-R (p < 0.0001), with enzyme levels of 0.260 and 0.204 µmol, respectively, and 188.44 and 156.67 nmol, respectively. The maximum activity of α-EST was detected in UPK-R, followed by PMD-R and MCM-S at 195.23, 183.82 and 127.73 nmol, respectively. PMD-R showed the greatest activity of ACP, followed by UPK-R and MCM-S, with enzyme levels of 96.21, 83.29 and 71.31 nmol, respectively. UPK-R showed the significantly highest level of ALP activity (20.82 nmol), with the lowest one being detected in PMD-R (12.65 nmol), which was not statistically different (p = 0.571) from that detected in MCM-S (13.21 nmol). MFO activity also was higher in resistant UPK-R and PMD-R (0.158 and 0.124 nmol, respectively) than in MCM-S (0.111 nmol), but an insignificant difference of MFO level was observed between PMD-R and MCM-S. There was no significant difference among AChE activities estimated in UPK-R, PMD-R and MCM-S, with enzyme levels of 11.40, 10.51 and 10.96 nmol, respectively.

3.2. Threshold Time for the Lethal Effect of P. crispum Oil on Ae. aegypti Larvae

Determination of threshold time for the lethal effect of P. crispum oil on 4th instar Ae. aegypti larvae revealed that the mortality rate depended on the exposure period (Figure 2). Higher mortality was recorded with increasing exposure time. Approximately 20% of larvae were killed at 3 h and about 50% mortality was recorded at 24 h of treatment. All larvae maintained in control and untreated conditions survived and were still active during the experimental period. The results clearly indicated that exposure to LC50 of P. crispum oil for 3, 6, 12, and 24 h generated varied larval mortalities in the range of 20–50%, with no significant difference among three strains of Ae. aegypti. As larval mortalities from 12- and 24-h exposure were not significantly different, only three time points, such as 3, 6 and 24 h, were selected as suitable lethal threshold times for investigating the effect of P. crispum oil on biochemical constituents of Ae. aegypti larvae.

3.3. Effects of P. crispum Oil on Biochemical Features of Ae. aegypti Larvae

The activity levels of all evaluated enzymes, including GSTs, α-/β-ESTs, AChE, ACP, ALP and MFO in the control and untreated groups of each strain of Ae. aegypti, were not significantly different (p > 0.05) at any of the time points (Figure 3 and Figure 4, Table 1). The enzyme profiles, except for α-/β-ESTs, also were altered similarly among the three strains of Ae. aegypti in these two groups during 24-h exposure. The activity level of GSTs in the control and untreated larvae of all strains was increased slightly from their baseline activity (0 h treatment). The increased AChE and ACP as well as decreased ALP and MFO activities in the control and untreated larvae of all strains were significantly different (p < 0.0001) from their baselines, except for the increased AChE activity in PMD-R.
After treatment with P. crispum oil for 3, 6 and 24 h, the activity levels of enzymes determined in the treated groups of each Ae. aegypti strain differed from those of the control and untreated groups (Figure 3 and Figure 4, Table 1). When comparing with the controls, the levels of GSTs in all strains of treated Ae. aegypti larvae were increased at every exposure time point (Figure 3A). Significant differences of GSTs activity between the treated and control groups were observed in MCM-S at 3 and 24 h, PMD-R at 24 h, and UPK-R at 3, 6 and 24 h of treatment (p < 0.0001). Although activity of α-/β-ESTs, ACP and ALP either increased or decreased in the three strains of treated larvae at all-time points during the exposure period, the levels of these enzymes, when detected at 24 h of treatment, eventually up-regulated when compared to those of the controls (Figure 3 and Figure 4, Table 1). Conversely, AChE activity in all strains of the treated larvae was reduced at every time point of exposure, when compared to that of the controls (Figure 3D). A statistically significant reduction of AChE activity was estimated in all strains of the treated larvae at every time point (MCM-S: 6 h, p = 0.000137; 24 h, p < 0.0001; PMD-R: 3 h, p = 0.002; 6 h, p = 0.025; 24 h, p = 0.000017; UPK-R: 3 h, p = 0.027; 6 h, p < 0.0001; 24 h, p = 0.002), apart from MCM-S at 3 h of treatment. Similar to AChE, the activity of MFO in all strains of treated larvae also decreased at every time point, except for those detected in PMD-R at 3- and 6-h exposure (Figure 4C).

4. Discussion

Plant essential oils have been recognized as potential sources of bioinsecticides used in agricultural pest and public health vector control programs. Among essential oils and ethanolic extracts of medicinal plants evaluated previously in the laboratory at Chiang Mai University, P. crispum essential oil exhibited larvicidal and adulticidal potential against both pyrethroid susceptible and resistant strains of Ae. aegypti [31]. With comparably mosquitocidal activity against pyrethroid susceptible and resistant Ae. aegypti, P. crispum essential oil became the candidate investigated herein for its effects on target mosquitoes, specifically in biochemical alterations. Three laboratory strains of Ae. aegypti, including MCM-S, PMD-R and UPK-R used in this study had been investigated by several investigators for status and mechanisms of insecticide resistance [39,40,42,43]. It was reported that while PMD-R showed resistance to permethrin, but susceptibility to deltamethrin, UPK-R proved to be resistant to both insecticides. However, our previous study revealed that both PMD-R and UPK-R were resistant to temephos, permethrin and deltamethrin in either the larval or adult stage, as compared to MCM-S [31]. Based on LC50 values obtained from a larvicidal bioassay, it is estimated that the susceptibility levels toward temephos, permethrin, and deltamethrin observed in PMD-R and UPK-R were lower than those of MCM-S by 1.9- and 1.9-fold; 4.1- and 286.0-fold; and 9.4- and 58.6-fold, respectively. According to Mazzarri and Georghiou [52], it can be clarified that PMD-R has low resistance to temephos and permethrin, but moderate resistance to deltamethrin, whereas UPK-R has low resistance to temephos, but high resistance to permethrin and deltamethrin.
Biochemical profiles of Ae. aegypti larvae prior to treatment with P. crispum essential oil demonstrated higher levels of some detoxifying enzymes, such as GSTs, α-/β- ESTs, ACP and MFO in resistant PMD-R and/or UPK-R, than those in susceptible MCM-S. These findings were in agreement with those of many biochemical studies, which evaluated various pest species in either laboratory or field populations, and demonstrated increased levels of detoxifying enzymes such as GSTs [53,54], α-/β-ESTs [34,55,56,57,58], ACP [59,60], and MFO [61,62,63] in insecticide-resistant insects. Evidence of insecticide-resistant populations manifesting a higher level of defensive enzymes than susceptible ones, suggests that these enzymes possibly play a role in developing insect resistance. Many researchers have indicated that qualitative and/or quantitative changes of defensive enzymes may be an important process associated with the resistance mechanism [15,64], and elevation of enzyme activity implicate an enhancer in the tolerance of mosquitoes to insecticides [55,56,65,66]. In consequence, resistance to some insecticides has been linked to an increment in insecticide metabolism through the up-regulation or structural changes of detoxifying enzymes [67,68]. In this regard, resistance of PMD-R and UPK-R to temephos, permethrin, and deltamethrin may be due partly to alteration in the rate of insecticide detoxification via increased enzyme activity of the insect organism. Thus, the expression of detoxification enzymes such as GSTs, α-/β-ESTs, ACP and MFO might be used as an important biomarker of insecticide resistance in the insect population.
Determining the threshold time for the lethal effect of P. crispum essential oil at LC50, demonstrated the exposure-time dependent efficacy against 4th instar of Ae. aegypti, with no significant difference of larval mortality among the three strains at any time point of treatment. Koodalingam et al. [37] investigated threshold time for the lethal effect of the soapnut, Sapindus emarginatus, on 4th instar larvae and pupae of Ae. aegypti by exposing them to lethal concentrations that produced 100% mortality within 24 h of exposure. Their study suggested that suitable lethal threshold times should be considered from the earliest time point of exposure, which generated the mortality outcome of 20–30% of test organisms upon their exposure to a specific test concentration. In this regard, 12 and 18 h of exposure were chosen as the threshold time points used for determining the lethal effect of soapnut extract on the larvae and pupae of Ae. aegypti, respectively. In order to gain more detailed insights of metabolic alterations affected by P. crispum oil in this study, sublethal concentrations (LC50) with different exposure periods, which generated a wide range of mortality, were applied on the three strains of Ae. aegypti. As a consequence, exposure times of 3, 6 and 24 h to LC50 of P. crispum oil producing 20–50% larval mortality were considered as appropriate threshold time points for determining the impact of this botanical oil on biochemical alterations in Ae. aegypti larvae.
Evaluation of biochemical enzymes at three points of the lethal threshold times (3, 6 and 24 h of exposure) showed that after maintaining at a similar condition to the treated group, activities of all the evaluated enzymes, including GSTs, α-/β-ESTs, AChE, ACP, ALP and MFO observed in all strains of the control and untreated Ae. aegypti larvae, were not different at any of the time points. Furthermore, alterations of enzyme activity at 24-h intervals in these two groups were similar among the three strains of Ae. aegypti. Slight alterations either increased or decreased activity of GSTs and α-/β-ESTs in the control and untreated larvae of all mosquito strains, except for the decreased activity of β-EST in UPK-R, were different from their baseline activity (0 h treatment). However, apart from the AChE activity in PMD-R that insignificantly increased, significant changes in the enzyme activity of increased AChE and ACP as well as decreased ALP and MFO, were detected in the control and untreated larvae of all mosquito strains. It can be postulated that the activity of these defensive enzymes in the control and untreated groups was modulated during the normal developmental period (24 h of experiment), thus showing their importance in diverse physiological processes in the development of mosquito larvae, as reported in various insect [69,70].
Different levels of some detoxifying enzymes in this study also were detected between resistant PMD-R and UPK-R strains, in particular, higher levels of GSTs, β-EST, and MFO in UPK-R, either before or after treatment with P. crispum essential oil. As indicated previously, elevation of these enzymes presents insecticide resistance to pyrethroids [71,72,73]. Therefore, differences in enzyme activity between PMD-R and UPK-R probably contribute to their differential susceptibility or resistance to insecticides. In support of these findings, previous results [31] also proved that resistance was higher in UPK-R than in PMD-R by 69.8- and 6.2-fold for permethrin and deltamethrin, respectively. The activity levels of phosphatase enzymes that are responsible for detoxifying organophosphate insecticides, also were different between resistant PMD-R and UPK-R, which showed the highest level of ACP and ALP activity, respectively. Therefore, comparable resistance to organophosphate temephos (1.9-fold of MCM-S), as previously recorded in PMD-R and UPK-R [31], may be a result of balanced activity between two phosphatases, ACP, and ALP, in these resistant strains.
Upon exposing Ae. aegypti larvae to sublethal concentrations of P. crispum essential oil for 3, 6 and 24 h, enzymatic alterations of either up-regulated or down-regulated activity were detected in all of these mosquito strains. It was recognized that the activity levels of evaluated enzymes in the treated larvae were different from those of the control and untreated groups, depending on the type of enzyme, strain of mosquito, and time point after treatment. However, the enzyme expression patterns in the treated larvae altered similarly among the three strains of Ae. aegypti during 24 h of treatment, except for those of β-EST recorded at the 3- and 6-h time points as well as ACP and ALP at the 3-h time point. Although the baseline activity of most defensive enzymes was lower in susceptible MCM-S, than in resistant PMD-R and/or UPK-R, the higher up-regulation of α-/β-ESTs and ALP activity in MCM-S, after treatment with P. crispum essential oil, resulted in comparable enzyme levels in susceptible and resistant mosquitoes at the 24-h time point. Unlike, the similar elevation rates of GSTs and ACP activity in susceptible and resistant strains of mosquitoes contributed to lower levels of these enzymes in treated MCM-S, as compared to those of PMD-R and/or UPK-R at the 24-h time point. However, the activities of GSTs, α-/β-ESTs, ACP, and ALP in treated larvae of all the Ae. aegypti strains were found to elevate after 24 h of treatment. These findings corresponded to those of many studies that demonstrated increased levels of GSTs [27,74,75], α-/β-ESTs [66,76,77,78], and ACP and ALP [69,79,80] in plant product-treated insects. The prominent alterations of these enzymes possibly correlate to the phytochemicals present in plant products, and their increased expression might suggest a probable role in detoxifying tested biocides into nontoxic substances [81,82].
GSTs, esterases (α-/β-ESTs), and phosphatases (ACP and ALP) are enzymes involved in the transformation and/or elimination of endogenous and exogenous compounds through several metabolic pathways in insects [34,35,36,37]. In addition to catalyzing the conjugation of glutathione with electrophilic compounds, GSTs also act as ligand-binding proteins that lead to the sequestration of xenobiotics [83,84,85,86]. Esterases are classified as hydrolases, a large and diverse group of enzymes that catalyze ester bonds found in a wide range of insecticides such as organophosphates, carbamates, and pyrethroids [87]. Phosphatases play an important role in diverse physiological processes and act as hydrolytic enzymes that catalyze removal of phosphate groups by hydrolyzing phosphate ester bonds, which are found in organophosphate insecticides [88,89]. Regarding the results obtained from the present and previous study, the increased activity of defensive enzymes, including GSTs, α-/β-ESTs, ACP, and ALP, in P. crispum oil-treated larvae of all the Ae. aegypti strains, indicated their possible role in metabolizing oil components. Elevation of these enzymes, observed during treatment with P. crispum essential oil, may be attributed to their overproduction. It is hypothesized that some oil constituents that enter the body of insects could be metabolized directly by these detoxifying enzymes, while the rest of the tested oil that binds to receptors could induce more enzyme production [66].
AChE and MFO activity in all of the treated Ae. aegypti strains in this study reduced when compared to the controls, as recorded after exposure to P. crispum essential oil for 24 h. Similar results of reduced activity of AChE [37,90,91] and MFO [22,26,92,93,94] were observed in several insect species exposed to different botanical products. These findings were contrary to those in many studies, which demonstrated biochemical responses with elevation of AChE or MFO activity after treatment with either synthetic substances or plant products. High activity of AChE and MFO was observed in all developmental stages of Culex pipiens treated with agricultural waste extracts, black and white liquors [95]. Evaluation of resistance to treatment with temephos and Pb-CVO (plant oil product) was carried out in a field collected wild strain (WS) and susceptible laboratory strain (LS) of Ae. aegypti [96]. The results demonstrated increased levels of cytochrome P450 monooxygenases (CYP450) in both strains of Pb-CVO-treated Ae. aegypti. However, CYP450 levels were increased in the LS, but decreased in the WS after treatment with temephos. Up-regulation of CYP450 level also was observed in Ae. aegypti larvae treated with the lethal concentration of Alangium salvifolium methanolic leaf extract [27]. The effects of girgensohnine alkaloid analog (DPPA) and Cymbopogon flexuosus essential oil (CFEO) on the behavior of detoxifying enzymes were evaluated in Rockefeller laboratory (Rock) and Santander wild (WSant) strains of Ae. aegypti [97]. It was found that the AChE activity in Rock and WSant was not affected by the evaluated dosages of DPPA and CFEO (p > 0.05), while MFO activity was significantly influenced by all of the CFEO dosages in WSant (p < 0.05). It was noticeable from these findings that differences in biochemical response in the target insects may be influenced by a variety of factors such as type of enzyme, type, and dosage of test substance, as well as developmental stages, species, and susceptibility of the insect.
AChE, a serine esterase (hydrolase) mainly found in nerve synapses, is a key enzyme that terminates nerve impulses by catalyzing hydrolysis of the neurotransmitter acetylcholine in the nervous system [98]. AChE also is the primary target of organophosphorus and carbamate compounds, which inhibit this enzyme, resulting in a general nervous system failure [99,100,101]. MFO is a family of enzymes induced when organisms are exposed to a variety of xenobiotics, and also is the most important enzyme involved in detoxifying pyrethroid and organophosphorus insecticides [102,103,104]. The increase and decrease of AChE and MFO after exposure to natural or synthetic compounds would probably result from the impact of these substances on insect metabolism by enzyme regulations, either inhibiting or stimulating enzyme activity. This appears to show biochemical interruption of diversified physiological mechanisms of insects [69]. Decreased AChE and MFO activity in all of the P. crispum oil-treated larvae strains in this study led to the postulation that these enzymes play no or a minimal role in metabolizing the tested oil, but they are a possible target of oil components. Due to the important role in various physiological processes, reduction of AChE and MFO, resulting from inhibitory effect of P. crispum essential oil, probably generates physiological disturbance and eventual death of Ae. aegypti larvae.
The comparatively pronounced larvicidal capability in pyrethroid susceptible MCM-S and resistant PMD-R and UPK-R of Ae. aegypti [31], and the inhibitory effect on AChE and MFO, emphasizes P. crispum essential oil as a qualified alternative for further development and application in current and future mosquito control programs. While AChE inhibitor affects nerve transmission, inhibition of MFO results in insecticide detoxification. Of interest, reduction of MFO activity in resistant UPK-R (21.8%) and PMD-R (13.2%), after treatment with P. crispum essential oil, sparked the idea of using this plant oil in managing mosquito resistance by combining it with existing conventional insecticides that are degraded by MFO such as pyrethroids and organophosphates, and this topic needs to be studied further. The combination of P. crispum essential oil and chemical compounds is a targetable prospect for synergist addition, which not only decreases dose application of insecticides, but also increases their mosquitocidal efficacy. As the target of MFO, pyrethroids and organophosphates can be detoxified by this enzyme, the lowered level of MFO leads to lowered insecticide detoxification, resulting in enhanced insecticidal efficacy. The synergism potential from MFO inhibition of P. crispum essential oil, which increased the toxic effects of pyrethroids and organophosphates, might be helpful in fighting against resistant mosquitoes, which have a high level of MFO activity. Furthermore, as P. crispum essential oil is a natural product comprising varied and complex active principles, with multiple modes of action through different targets such as AChE and MFO, a combination of P. crispum oil-insecticide may contribute to prolong the generation of resistance in vector populations. In consequence, this cumulative strategy may improve the performance of integrated mosquito management programs.

5. Conclusions

Biochemical profiles in the pyrethroid susceptible and resistant strains of Ae. aegypti larvae before and after exposure to P. crispum oil showed alterations of key enzymes responsible for xenobiotic detoxification, including GSTs, α-/β-ESTs, AChE, ACP, ALP, and MFO. The recognizable larvicidal capability on pyrethroid resistant Ae. aegypti and the inhibitory effect on AChE and MFO emphasize the potential of P. crispum oil as an attractive alternative for the management of mosquito resistance in current and future control programs.

Author Contributions

Conceptualization, J.I., A.J. (Anuluck Junkum) and B.P.; Data curation, J.I., A.J. (Anuluck Junkum), N.L. and D.C.; Formal analysis, J.I., A.J. (Anuluck Junkum), N.L., A.J. (Atchariya Jitpakdi), D.R., D.C. and B.P.; Funding acquisition, A.J. (Anuluck Junkum), P.S. and B.P.; Methodology, J.I., A.J. (Anuluck Junkum), N.L., U.C., A.J. (Atchariya Jitpakdi), D.R., D.C., R.M., A.C. and B.P.; Project administration, A.J. (Anuluck Junkum) and B.P.; Writing–original draft, J.I.; Writing–review & editing, A.J. (Anuluck Junkum), N.L. and B.P.

Funding

We are very grateful to the financial support by the Faculty of Medicine Research Fund (PAR-2559-04337, contract number 023/2558 to A.J. (Anuluck Junkum)), Diamond Research Grant of the Faculty of Medicine (PAR-2560-04663, contract number 120/2560 to P.S., B.P., A.J. (Anuluck Junkum)), the Excellence Center in Insect Vector Study of the research administration (grant number: 2562 to P.S., B.P., A.J. (Anuluck Junkum)), Chiang Mai University (CMU), and the Bio & Medical Technology Development Program of the National Research Foundation (NRF) funded by the Ministry of Science & ICT, South Korea (grant number: 2017M3A9E40707 to P.S., B.P., A.J. (Anuluck Junkum)).

Acknowledgments

We would like to thank Wannaree Charoensup, a scientist at the Department of Pharmaceutical Science, Faculty of Pharmacy, CMU, Chiang Mai province, Thailand, for her assistance in taxonomic identification of the medicinal plants investigated.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Knudsen, A.B.; Sloof, R. Vector-borne disease problems in rapid urbanization: NEW approaches to vector control. Bull. World Health Organ. 1992, 70, 1–6. [Google Scholar] [PubMed]
  2. World Health Organization (WHO). Global Strategy for Dengue Prevention and Control, 2012–2020; WHO Press: Geneva, Switzerland, 2012; ISBN 978-9-241-50403-4. [Google Scholar]
  3. Weaver, S.C.; Costa, F.; Garcia-Blanco, M.A.; Ko, A.I.; Ribeiro, G.S.; Saade, G.; Shi, P.Y.; Vasilakis, N. Zika virus: History, emergence, biology, and prospects for control. Antivir. Res. 2016, 130, 69–80. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  4. Gubler, D.J. Epidemic dengue/dengue hemorrhagic fever as a public health, social and economic problem in the 21st century. Trends Microbiol. 2002, 10, 100–103. [Google Scholar] [CrossRef]
  5. World Health Organization (WHO). Dengue: GUIDELINES for Diagnosis, Treatment, Prevention and Control; WHO Press: Paris, France, 2009; ISBN 978-9-241-54787-1. [Google Scholar]
  6. World Health Organization (WHO). Dengue Fact Sheet. 2016. Available online: http://www.searo.who.int/entity/vector_borne_tropical_diseases/data/data_factsheet/en/ (accessed on 5 May 2018).
  7. World Health Organization (WHO). Dengue and Severe Dengue. 2018. Available online: http://www.who.int/news-room/fact-sheets/detail/dengue-and-severe-dengue/ (accessed on 17 May 2018).
  8. World Health Organization (WHO). Dengue Vaccine Research. 2016. Available online: https://www.who.int/immunization/research/development/dengue_vaccines/en/ (accessed on 17 May 2018).
  9. Champakaew, D.; Junkum, A.; Chaithong, U.; Jitpakdi, A.; Riyong, D.; Sanghong, R.; Intirach, J.; Muangmoon, R.; Chansang, A.; Tuetun, B.; et al. Angelica sinensis (Umbelliferae) with proven repellent properties against Aedes aegypti, the primary dengue fever vector in Thailand. Parasitol. Res. 2015, 114, 2187–2198. [Google Scholar] [CrossRef] [PubMed]
  10. Scott, L.J. Tetravalent Dengue Vaccine: A review in the prevention of dengue disease. Drugs 2016, 76, 1301–1312. [Google Scholar] [CrossRef] [PubMed]
  11. Badolo, A.; Ilboudo-Sanogo, E.; Ouédraogo, A.P.; Costantini, C. Evaluation of the sensitivity of Aedes aegypti and Anopheles gambiae complex mosquitoes to two insect repellents: DEET and KBR 3023. Trop. Med. Int. Health 2004, 9, 330–334. [Google Scholar] [CrossRef] [PubMed]
  12. Waliwitiya, R.; Kennedy, C.J.; Lowenberger, C.A. Larvicidal and oviposition-altering activity of monoterpenoids, trans-anethole and rosemary oil to the yellow fever mosquito Aedes aegypti (Diptera: Culicidae). Pest Manag. Sci. 2008, 65, 241–248. [Google Scholar] [CrossRef]
  13. Monnerat, R.; Dumas, V.; Ramos, F.; Pimentel, L.; Nunes, A.; Sujii, E.; Praca, L.; Vilarinhos, P. Evaluation of different larvicides for the control of Aedes aegypti (Linnaeus) (Diptera: Culicidae) under simulated field conditions. Bioassay 2012, 7, 3. [Google Scholar] [CrossRef]
  14. Zaim, M.; Aitio, A.; Nakashima, N. Safety of pyrethroid-treated mosquito nets. Med. Vet. Entomol. 2000, 14, 1–5. [Google Scholar] [CrossRef] [Green Version]
  15. Chareonviriyaphap, T.; Bangs, M.J.; Suwonkerd, W.; Kongmee, M.; Corbel, V.; Ngoen-Klan, R. Review of insecticide resistance and behavioral avoidance of vectors of human diseases in Thailand. Parasit. Vectors 2013, 6, 280. [Google Scholar] [CrossRef] [Green Version]
  16. Grieco, J.P.; Achee, N.L.; Chareonviriyaphap, T. A new classification system for the actions of IRS chemicals traditionally used for malaria control. PLoS ONE 2007, 2, e716. [Google Scholar] [CrossRef]
  17. Thanispong, K.; Sathantriphop, S.; Chareonviriyaphap, T. Insecticide resistance of Aedes aegypti and Culex quinquefasciatus in Thailand. J. Pestic. Sci. 2008, 33, 351–356. [Google Scholar] [CrossRef]
  18. Ranson, H.; Burhani, J.; Lumjuan, N.; Black, W.C. Insecticide resistance in dengue vectors. TropIKAnet J. 2010, 1, 1–12. [Google Scholar]
  19. Karunamoorthi, K.; Sabesan, S. Insecticide resistance in insect vectors of disease with special reference to mosquitoes: A potential threat to global public health. Health Scope 2013, 2, 4–18. [Google Scholar] [CrossRef]
  20. World Health Organization (WHO). Monitoring and Managing Insecticide Resistance in Aedes aegypti Populations, Interim Guidance for Entomologist. 2016. Available online: http://www.who.int/csr/resources/publications/zika/insecticide-resistance/en/ (accessed on 20 March 2016).
  21. Chansang, A.; Champakaew, D.; Junkum, A.; Jitpakdi, A.; Amornlerdpison, D.; Aldred, A.K.; Riyong, D.; Wannasan, A.; Intirach, J.; Muangmoon, R.; et al. Synergy in the adulticidal efficacy of essential oils for the improvement of permethrin toxicity against Aedes aegypti L. (Diptera: Culicidae). Parasit. Vectors 2018, 11, 417. [Google Scholar] [CrossRef] [PubMed]
  22. Tong, F.; Bloomquist, J.R. Plant essential oils affect the toxicities of carbaryl and permethrin against Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 2013, 50, 826–832. [Google Scholar] [CrossRef] [PubMed]
  23. Pitasawat, B.; Champakaew, D.; Choochote, W.; Jitpakdi, A.; Chaithong, U.; Kanjanapothi, D.; Rattanachanpichai, E.; Tippawangkosol, P.; Riyong, D.; Tuetun, B.; et al. Aromatic plant-derived essential oil: AN alternative larvicide for mosquito control. Fitoterapia 2007, 78, 205–210. [Google Scholar] [CrossRef]
  24. Kweka, E.J.; Nyindo, M.; Mosha, F.; Sila, A.G. Insecticidal activity of the essential oil from fruits and seeds of Schinus terebinthifolia Raddi against African malaria vectors. Parasit. Vectors 2011, 4, 129. [Google Scholar] [CrossRef]
  25. Champakaew, D.; Choochote, W.; Pongpaibul, Y.; Chaithong, U.; Jitpakdi, A.; Tuetun, B.; Pitasawat, B. Larvicidal efficacy and biological stability of a botanical natural product, zedoary oil-impregnated sand granules, against Aedes aegypti (Diptera, Culicidae). Parasitol. Res. 2007, 100, 729–737. [Google Scholar] [CrossRef]
  26. Waliwitiya, R.; Nicholson, R.A.; Kennedy, C.J.; Lowenberger, C.A. The synergistic effects of insecticidal essential oils and piperonyl butoxide on biotransformational enzyme activities in Aedes aegypti (Diptera: Culicidae). J. Med. Entomol. 2012, 49, 614–623. [Google Scholar] [CrossRef]
  27. Thanigaivel, A.; Vasantha-Srinivasan, P.; Edwin, E.S.; Ponsankar, A.; Selin-Rani, S.; Chellappandian, M.; Kalaivani, K.; Senthil-Nathan, S.; Benelli, G. Development of an eco-friendly mosquitocidal agent from Alangium salvifolium against the dengue vector Aedes aegypti and its biosafety on the aquatic predator. Environ. Sci. Pollut. Res. Int. 2018, 25, 10340–10352. [Google Scholar] [CrossRef]
  28. Tehri, K.; Singh, N. The role of botanicals as green pesticides in integrated mosquito management—A review. Int. J. Mosq. Res. 2015, 2, 18–23. [Google Scholar]
  29. Bekele, D. Review on insecticidal and repellent activity of plant products for malaria mosquito control. Biomed. Res. Rev. 2018. [Google Scholar] [CrossRef]
  30. Jankowska, M.; Rogalska, J.; Wyszkowska, J.; Stankiewicz, M. Molecular targets for components of essential oils in the insect nervous system-A review. Molecules 2018, 23, 34. [Google Scholar] [CrossRef]
  31. Intirach, J.; Junkum, A.; Lumjuan, N.; Chaithong, U.; Jitpakdi, A.; Riyong, D.; Wannasan, A.; Champakaew, D.; Muangmoon, R.; Chansang, A.; et al. Antimosquito property of Petroselinum crispum (Umbellifereae) against the pyrethroid resistant and susceptible strains of Aedes aegypti (Diptera: Culicidae). Environ. Sci. Pollut. Res. Int. 2016, 23, 23994–24008. [Google Scholar] [CrossRef]
  32. Okumu, F.O.; Knols, B.G.J.; Fillinger, U. Larvicidal effects of a neem (Azadirachta indica) oil formulation on the malaria vector Anopheles gambiae. Malar. J. 2007, 6, 63. [Google Scholar] [CrossRef] [PubMed]
  33. Liao, M.; Xiao, J.J.; Zhou, L.J.; Liu, Y.; Wu, X.W.; Hua, R.M.; Wang, G.R.; Cao, H.Q. Insecticidal activity of Melaleuca alternifolia essential oil and RNA-Seq analysis of Sitophilus zeamais transcriptome in response to oil fumigation. PLoS ONE 2016, 11, e0167748. [Google Scholar] [CrossRef]
  34. Brengues, C.; Hawkes, N.J.; Chandre, F.; McCarroll, L.; Duchon, S.; Guillet, P.; Manguin, S.; Morgan, J.C.; Hemingway, J. Pyrethroid and DDT cross-resistance in Aedes aegypti correlated with novel mutations in the voltage-gated sodium channel gene. Med. Vet. Entomol. 2003, 17, 87–94. [Google Scholar] [CrossRef] [PubMed]
  35. Mukherjee, P.K.; Kumar, V.; Mal, M.; Houghton, P.J. Acetylcholinesterase inhibitors from plants. Phytomedicine 2007, 14, 289–300. [Google Scholar] [CrossRef]
  36. Polson, K.; Brogdon, W.G.; Rawlins, S.C.; Chadee, D.D. Characterization of insecticide resistance in Trinidadian strains of Aedes aegypti mosquitoes. Acta Trop. 2011, 117, 31–38. [Google Scholar] [CrossRef] [PubMed]
  37. Koodalingam, A.; Mullainadhan, P.; Arumugam, M. Effects of extract of soapnut Sapindus emarginatus on esterases and phosphatases of the vector mosquito, Aedes aegypti (Diptera: Culicidae). Acta Trop. 2011, 118, 27–36. [Google Scholar] [CrossRef] [PubMed]
  38. Sutthanont, N.; Choochote, W.; Tuetun, B.; Junkum, A.; Jitpakdi, A.; Chaithong, U.; Riyong, D.; Pitasawat, B. Chemical composition and larvicidal activity of edible plant-derived essential oils against the pyrethroid-susceptible and -resistant strains of Aedes aegypti (Diptera: Culicidae). J. Vector Ecol. 2010, 35, 106–115. [Google Scholar] [CrossRef] [PubMed]
  39. Prapanthadara, L.; Promtet, N.; Koottathep, S.; Somboon, P.; Suwonkerd, W.; McCarroll, L.; Hemingway, J. Mechanisms of DDT and permethrin resistance in Aedes aegypti from Chiang Mai, Thailand. Dengue Bull. 2002, 26, 185–189. [Google Scholar]
  40. Plernsub, S.; Saingamsook, J.; Yanola, J.; Lumjuan, N.; Tippawangkosol, P.; Sukontason, K.; Somboon, P. Additive effect of knockdown resistance mutations, S989P, V1016G and F1534C, in a heterozygous genotype conferring pyrethroid resistance in Aedes aegypti in Thailand. Parasit. Vectors 2016, 9, 417. [Google Scholar] [CrossRef] [PubMed]
  41. Yanola, J.; Somboon, P.; Walton, C.; Nachaiwieng, W.; Prapanthadara, L. A novel F1552/C1552 point mutation in the Aedes aegypti voltage-gated sodium channel gene associated with permethrin resistance. Pest Biochem. Physiol. 2010, 96, 127–131. [Google Scholar] [CrossRef]
  42. Somwang, P.; Yanola, J.; Suwan, W.; Walton, C.; Lumjuan, N.; Prapanthadara, L.A.; Somboon, P. Enzymes-based resistant mechanism in pyrethroid resistant and susceptible Aedes aegypti strains from northern Thailand. Parasitol. Res. 2011, 109, 531–537. [Google Scholar] [CrossRef]
  43. Lumjuan, N.; Wicheer, J.; Leelapat, P.; Choochote, W.; Somboon, P. Identification and characterisation of Aedes aegypti aldehyde dehydrogenases involved in pyrethroid metabolism. PLoS ONE 2014, 9, e102746. [Google Scholar] [CrossRef]
  44. World Health Organization. Test Procedures for Insecticide Resistance Monitoring in Malaria Vectors, Bio-Efficacy and Persistence of Insecticide on Treated Surfaces: Report of the WHO Informal Consultation; WHO Press: Geneva, Switzerland, 1998. [Google Scholar]
  45. World Health Organization. Instruction for Determining the Susceptibility or Resistance of Mosquito Larvae to Insecticide; WHO/VBC/81.807; WHO Press: Geneva, Switzerland, 1981. [Google Scholar]
  46. Bradford, M.M. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 1976, 72, 248–254. [Google Scholar] [CrossRef]
  47. Habig, W.H.; Pabst, M.J.; Jakoby, W.B. Glutathione S-transferases: The first enzymatic step in mercapturic acid formation. J. Biol. Chem. 1974, 249, 7130–7139. [Google Scholar]
  48. Van Asperen, K. A study of housefly esterases by means of a sensitive colorimetric method. J. Insect Physiol. 1962, 8, 401–416. [Google Scholar] [CrossRef]
  49. Ellman, G.L.; Courtney, K.; Andres, V.; Featherstone, R.M. A new and rapid colorimetric determination of acetylcholinesterase activity. Biochem. Pharmacol. 1961, 7, 88–95. [Google Scholar] [CrossRef] [Green Version]
  50. Asakura, K. Phosphatase activity in the larva of the euryhalinemosquito, Aedes togoi Theobold, with special reference to sea water adaptation. J. Exp. Mar. Biol. Ecol. 1978, 31, 325–337. [Google Scholar] [CrossRef]
  51. Brogdon, W.G.; McAllister, J.C.; Vulule, J. Heme peroxidase activity measured in single mosquitoes identifies individuals expressing an elevated oxidase for insecticide resistance. J. Am. Mosq. Control Assoc. 1997, 13, 233–237. [Google Scholar] [PubMed]
  52. Mazzarri, M.B.; Georghiou, G.P. Characterization of resistance to organophosphate, carbamate, and pyrethroid insecticides in field populations of Aedes aegypti from Venezuela. J. Am. Mosq. Control Assoc. 1995, 11, 315–322. [Google Scholar]
  53. Grant, D.F. Evolution of glutathione S-transferase subunits in Culicidae and related Nematocera: ELECTROPHORETIC and immunological evidence for conserved enzyme structure and expression. Insect Biochem. 1991, 21, 435–445. [Google Scholar] [CrossRef]
  54. Grant, D.F.; Dietze, E.C.; Hammock, B.D. Glutathione S-transferase isozymes in Aedes aegypti: PURIFICATION, characterization, and isozyme specific regulation. Insect Biochem. 1991, 4, 421–433. [Google Scholar] [CrossRef]
  55. Macoris, M.L.G.; Andrighetti, L.; Takaku, L.; Glasser, C.M.; Garbeloto, V.V.; Bracco, J.E. Resistance of Aedes aegypti from the state of Sao Paulo, Brazil, to organophosphate insecticides. Mem. Inst. Oswaldo Cruz 2003, 98, 703–708. [Google Scholar] [CrossRef]
  56. Yaicharoen, R.; Kiatfuengfoo, R.; Chareonviriyaphap, T.; Rongnoparut, P. Characterization of deltamethrin resistance in field populations of Aedes aegypti in Thailand. J. Vector Ecol. 2005, 30, 144–150. [Google Scholar] [PubMed]
  57. Jagadeshwaran, U.; Vijayan, V. Biochemical characterization of deltamethrin resistance in a laboratory-selected strain of Aedes aegypti. Parasitol. Res. 2009, 104, 1431–1438. [Google Scholar]
  58. Pimsamarn, S.; Sornpengb, W.; Akksilp, S.; Paeporn, P.; Limpawitthayakul, M. Detection of insecticide resistance in Aedes aegypti to organophosphate and synthetic pyrethroid compounds in the north-east of Thailand. Dengue Bull. 2009, 33, 194–202. [Google Scholar]
  59. Awan, D.A.; Saleem, M.A.; Nadeem, M.S.; Shakoori, A.R. Toxicological and biochemical studies on spinosad and synergism with piperonyl butoxide in susceptible and resistant strains of Tribolium castaneum. Pak. J. Zool. 2012, 44, 649–662. [Google Scholar]
  60. Hariprasad, T.P.N.; Shetty, N.J. Biochemical basis of alphamethrin resistance in different life stages of Anopheles stephensi strains of Bangalore, India. Pest Manag. Sci. 2016, 72, 1689–1701. [Google Scholar] [CrossRef] [PubMed]
  61. Amin, A.M. Preliminary investigation of the mechanisms of DDT and pyrethroid resistance in Culex quinquefasciatus Say. (Diptera: Culicidae) from Saudi Arabia. Bull. Entomol. Res. 1989, 79, 361–366. [Google Scholar] [CrossRef]
  62. Putra, R.E.; Ahmad, I.; Prasetyo, D.B.; Susanti, S.; Rahayu, R.; Hariani, N. Detection of insecticide resistance in the larvae of some Aedes aegypti (Diptera: Culicidae) strains from Java, Indonesia to temephos, malathion and permethrin. Int. J. Mosq. Res. 2016, 3, 23–28. [Google Scholar]
  63. Choovattanapakorn, N.; Yanola, J.; Lumjuan, N.; Saingamsook, J.; Somboon, P. Characterization of metabolic detoxifying enzymes in an insecticide resistant strain of Aedes aegypti harboring homozygous S989P and V1016G kdr mutations. Med. Entomol. Zool. 2017, 68, 19–26. [Google Scholar] [CrossRef]
  64. Hemingway, J.; Karunaratne, S.H.P.P. Mosquito carboxylesterases: A review of the molecular biology and biochemistry of a major insecticide resistance mechanism. Med. Vet. Entomol. 1998, 12, 1–12. [Google Scholar] [CrossRef]
  65. Chareonviriyaphap, T.; Rongnoparut, P.; Chantarumporn, P.; Bangs, M.J. Biochemical detection of pyrethroid resistance mechanisms in Anopheles minimus in Thailand. J. Vector Ecol. 2003, 28, 108–116. [Google Scholar]
  66. Ahmad, I.; Astari, S.; Tan, M. Resistance of Aedes aegypti (Diptera: Culicidae) in 2006 to pyrethroid insecticides in Indonesia and its association with oxidase and esterase levels. Pak. J. Biol. Sci. 2007, 10, 3688–3692. [Google Scholar]
  67. Braga, I.A.; Valle, D. Aedes aegypti: INSETICIDAS, mecanismos de ação e resistência. Epidemiol. Serv. Saúde 2007, 16, 279–293. [Google Scholar] [CrossRef]
  68. Diniz, D.F.A.; de Melo-Santos, M.A.V.; de Mendonca Santos, E.A.; Beserra, E.B.; Helvecio, E.; de Carvalho-Leandro, D.; dos Santos, B.S.; de Menezes Lima, V.L.; Ayres, C.V.J. Fitness cost in field and laboratory Aedes aegypti populations associated with resistance to the insecticide temephos. Parasit. Vectors 2015, 8, 662. [Google Scholar] [CrossRef]
  69. Koodalingam, A.; Mullainadhan, P.; Rajalakshmi, A.; Deepalakshmi, R.; Ammua, M. Effect of a Bt-based product (Vectobar) on esterases and phosphatases from larvae of the mosquito Aedes aegypti. Pest Biochem. Physiol. 2012, 104, 267–272. [Google Scholar] [CrossRef]
  70. Singh, K.; Kaur, M.; Rup, P.J.; Singh, J. Effects of Indian coral tree, Erythrina indica lectin on eggs and larval development of melon fruit fly, Bactrocera cucurbitae. J. Environ. Biol. 2009, 30, 509–514. [Google Scholar] [PubMed]
  71. Berge, J.B.; Feyereisen, R.; Amichot, M. Cytochrome P450 monooxygenases and insecticide resistance in insects. Philos. Trans. R. Soc. Lond. B. Biol. Sci. 1998, 353, 1701–1705. [Google Scholar] [CrossRef] [PubMed] [Green Version]
  72. Vulule, J.M.; Beach, R.F.; Atieli, F.K.; McAllister, J.C.; Brogdon, W.G.; Roberts, J.M.; Mwangi, R.W.; Hawley, W.A. Elevated oxidase and esterase levels associated with permethrin tolerance in Anopheles gambiae from Kenyan villages using permethrin-impregnated nets. Med. Vet. Entomol. 1999, 3, 239–244. [Google Scholar] [CrossRef]
  73. Vontas, J.; Ranson, H.; Alphey, L. Transcriptomics and disease vector control. BMC Biol. 2010, 8, 52. [Google Scholar] [CrossRef] [PubMed]
  74. Sarita, N.; Moharil, M.P.; Ghodki, B.S.; Lande, G.K.; Bisane, K.D.; Thakare, A.S.; Barkhade, U.P. Biochemical analysis and synergistic suppression of indoxacarb resistance in Plutella xylostella L. J. Asia-Pac. Entomol. 2010, 13, 91–95. [Google Scholar]
  75. Zibaee, A.; Bandani, A.R. A study on the toxicity of a medicinal plant, Artemisia annua L. (Asteracea) extracts to the sunn pest, Eurygaster integriceps Puton (Hemiptera: Scutelleridae). J. Plant Protect. Res. 2010, 50, 79–85. [Google Scholar] [CrossRef]
  76. Wang, K.Y.; Yong, Z.; Yan, W.H.; Ming, X.X.; Xian, L.T. Influence of three diets on susceptibility of selected insecticides and activities of detoxification esterases of Helicoverpa assulta (Lepidoptera: Noctuidae). Pest Biochem. Physiol. 2010, 96, 51–55. [Google Scholar] [CrossRef]
  77. Gamil, W.E.; Mariy, F.M.; Youssef, L.A.; Abdel Halim, S.M. Effect of Indoxacarb on some biological and biochemical aspects of Spodoptera littoralis (Boisd.) larvae. Ann. Agric. Sci. 2011, 56, 121–126. [Google Scholar] [CrossRef]
  78. Huron, E.N.; Abbas, A.A.; El-Hamid, N.A.A.; Nada, M.S.; Amin, T.R. Toxicity and acute macromolecular abnormalities induced by some plant extracts against the Cowpea aphid; Aphis carricivora Koch. J. Plant Prot. Path. 2016, 7, 445–449. [Google Scholar]
  79. Shekari, M.; Sendi, J.J.; Etebari, K.; Zibaee, A.; Shadparvar, A. Effects of Artemisia annua L. (Asteracea) on nutritional physiology and enzyme activities of elm leaf beetle, Xanthogaleruca luteola Mull. (Coleoptera: Chrysomellidae). Pest Biochem. Physiol. 2008, 91, 66–74. [Google Scholar] [CrossRef]
  80. Ghoneim, K.; Amer, M.; Al-Daly, A.; Mohammad, A.; Khadrawy, F.; Mahmoud, M.A. Disturbed acid and alkaline phosphatase activities in desert locust Schistocerca gregaria (Forskal) (Orthoptera: Acrididae) by extracts from the khella plant Ammi visnaga L. (Apiaceae). Int. J. Adv. Res. 2014, 2, 584–596. [Google Scholar]
  81. Després, L.; David, J.P.; Gallet, C. The evolutionary ecology of insect resistance to plant chemicals. Trends Ecol. Evol. 2007, 22, 298–307. [Google Scholar] [CrossRef] [PubMed]
  82. Panini, M.; Manicardi, G.; Moores, G.; Mazzoni, E. An overview of the main pathways of metabolic resistance in insects. Invertebr. Surv. J. 2016, 13, 326–335. [Google Scholar]
  83. Hayes, J.D.; Wolf, C.R. Role of glutathione transferase in drug resistance. In Glutathione Conjugation: Mechanisms and Biological Significance; Sies, H., Ketterer, B., Eds.; Academic Press: London, UK, 1988; pp. 315–355. [Google Scholar]
  84. Mannervik, B.; Danielson, U.H. Glutathione transferases-structure and catalytic activity. CRC Crit. Rev. Biochem. 1988, 23, 283–337. [Google Scholar] [CrossRef] [PubMed]
  85. Pickett, C.B.; Lu, A.Y. Glutathione S-transferases: GENE structure, regulation, and biological function. Annu. Rev. Biochem. 1989, 58, 743–764. [Google Scholar] [CrossRef] [PubMed]
  86. Yang, Y.; Cheng, J.Z.; Singhal, S.S.; Saini, M.; Pandya, U.; Awasthi, S. Role of glutathione S-transferases in protection against lipid peroxidation. J. Biol. Chem. 2001, 276, 19220–19230. [Google Scholar] [CrossRef]
  87. Dauterman, W.C. Insect metabolism: EXTRAMICROSOMAL. In Comprehensive Insect Physiology, Biochemistry, and Pharmacology; Kerkut, G.A., Gilbert, L.I., Eds.; Pergamon Press: Oxford, UK, 1985; pp. 713–730. [Google Scholar]
  88. Callaghan, A. Insecticide resistance: MECHANISM and detection methods. Sci. Progress 1991, 75, 423–438. [Google Scholar]
  89. Majerus, P.; Kisseleva, M.; Norris, F.A. The role of phosphatases in inositol signaling reactions. J. Biol. Chem. 1999, 274, 10669–10672. [Google Scholar] [CrossRef]
  90. Chaubey, M.K. Fumigant toxicity of essential oils against rice weevil Sitophilus oryzae L. (Coleoptera: Curculionidae). J. Biol. Sci. 2011, 11, 411–416. [Google Scholar] [CrossRef]
  91. Seo, S.M.; Jung, C.S.; Kang, J.; Lee, H.R.; Kim, S.W.; Hyun, J.; Park, I.K. Larvicidal and acetylcholinesterase inhibitory activities of Apiaceae plant essential oils and their constituents against Aedes albopictus and formulation development. J. Agric. Food Chem. 2015, 63, 9977–9986. [Google Scholar] [CrossRef] [PubMed]
  92. Mclaughlin, L.A.; Niazi, U.; Bibby, J.; David, J.P.; Vontas, J.; Hemingway, J.; Ranson, H.; Sutcliffe, M.J.; Paine, M.J. Characterization of inhibitors and substrates of Anopheles gambiae CYP6Z2. Insect Mol. Biol. 2008, 17, 125–135. [Google Scholar] [CrossRef] [PubMed]
  93. Joffe, T.; Gunning, R.V.; Allen, G.R.; Kristensen, M.; Alptekin, S.; Field, L.M.; Moores, G.D. Investigating the potential of selected natural compounds to increase the potency of pyrethrum against houseflies Musca domestica (Diptera:Muscidae). Pest Manag. Sci. 2012, 68, 178–184. [Google Scholar] [CrossRef] [PubMed]
  94. Ramirez, G.I.J.; Logan, J.G.; Loza-Reyes, E.; Stashenko, E.; Moores, G.D. Repellents inhibit P450 enzymes in Stegomyia (Aedes) aegypti. PLoS ONE 2012, 7, e48698. [Google Scholar]
  95. Helmy, N.; Bakr, R.F.A.; Nawwar, G.A.; Ibrahim, S.E.; Helmy, O.M. Biochemical effects of some agricultural waste extracts against Culex pipiens (Diptera: Culicidae). Egyptian Acad. J. Biol. Sci. 2010, 2, 75–81. [Google Scholar] [CrossRef]
  96. Vasantha-Srinivasan, P.; Senthil-Nathana, S.; Ponsankar, A.; Thanigaivel, A.; Edwina, E.S.; Selin-Rani, S.; Chellappandiana, M.; Pradeepaa, V.; Lija-Escalinea, J.; Kalaivani, K.; et al. Comparative analysis of mosquito (Diptera: Culicidae: Aedes aegypti Liston) responses to the insecticide temephos and plant derived essential oil derived from Piper betle L. Ecotoxicol. Environ. Saf. 2017, 139, 439–446. [Google Scholar] [CrossRef] [PubMed]
  97. Carreño Otero, A.L.; Palacio-Cortés, A.M.; Navarro-Silva, M.A.; Kouznetsov, V.V.; Duque L, J.E. Behavior of detoxifying enzymes of Aedes aegypti exposed to girgensohnine alkaloid analog and Cymbopogon flexuosus essential oil. Comp. Biochem. Physiol. C Toxicol. Pharmacol. 2018, 204, 14–25. [Google Scholar] [CrossRef]
  98. Wang, J.J.; Cheng, W.X.; Ding, W.; Zhao, Z.M. The effect of insecticide dichlorvos on esterase activity extracted from the psocids, Liposcelis bostrychophila and L. entomophila. J. Insect Sci. 2004, 4, 1–5. [Google Scholar] [CrossRef]
  99. Harel, M.; Kryger, G.; Rosenberry, T.L.; Mallender, W.D.; Lewis, T.; Fletcher, R.J.; Guss, J.M.; Silman, I.; Sussman, J.L. Three-dimensional structures of Drosophila melanogaster acetylcholinesterase and its complexes with two potent inhibitors. Protein Sci. 2000, 9, 1063–1072. [Google Scholar] [CrossRef]
  100. Walker, C.H.; Hopkin, S.P.; Sibly, R.M.; Peakall, D.B. Principles of Ecotoxicology, 3rd ed.; Taylor & Francis: London, UK, 2012. [Google Scholar]
  101. Rebechi, D.; Richardi, V.S.; Vicentini, M.; Guilosky, I.C.; de Assis, H.C.S.; Navarro-Silva, M.A. Low malathion concentrations influence metabolism in Chironomus sancticaroli (Diptera, Chironomidae) in acute and chronic toxicity test. Rev. Bras. Entomol. 2014, 58, 296–301. [Google Scholar] [CrossRef]
  102. Hemingway, J.; Hawkes, N.J.; McCarroll, L.; Ranson, H. The molecular basis of insecticide resistance in mosquitoes. Insect Biochem. Mol. Biol. 2004, 34, 653–665. [Google Scholar] [CrossRef] [PubMed]
  103. Melo-Santos, M.V.; Varjal-Melo, J.J.M.; Araújo, P.; Gomes, T.C.S.; Paiva, M.H.S.; Regis, L.N.; Furtado, F.; Magalhaes, T.; Macoris, M.L.G.; Andrighetti, M.T.M.; et al. Resistance to the organophosphate temephos: MECHANISMS, evolution and reversion in an Aedes aegypti laboratory strain from Brazil. Acta Trop. 2010, 113, 180–189. [Google Scholar] [CrossRef] [PubMed]
  104. Ocampo, C.B.; Salazar-Terreros, M.J.; Mina, N.J.; McAllister, J.; Brogdon, W. Insecticide resistance status of Aedes aegypti in 10 localities in Colombia. Acta Trop. 2011, 118, 37–44. [Google Scholar] [CrossRef] [PubMed]
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Figure 1. Enzyme activity in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti prior to treatment (untreated group, 0 h time point). Each bar represents Mean ± SE of six or ten biological samples from different preparations. Bars with different letters on top indicate statistically significant differences among mosquito strains (Tukey’s HSD test, p < 0.05).
Figure 1. Enzyme activity in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti prior to treatment (untreated group, 0 h time point). Each bar represents Mean ± SE of six or ten biological samples from different preparations. Bars with different letters on top indicate statistically significant differences among mosquito strains (Tukey’s HSD test, p < 0.05).
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Figure 2. Threshold time for the lethal effect of exposure to LC50 of P. crispum oil on the three strains of Ae. aegypti larvae (43.22, 44.50 and 44.03 ppm for MCM-S, PMD-R and UPK-R strains, respectively). The error bars represent the standard errors of four replicates.
Figure 2. Threshold time for the lethal effect of exposure to LC50 of P. crispum oil on the three strains of Ae. aegypti larvae (43.22, 44.50 and 44.03 ppm for MCM-S, PMD-R and UPK-R strains, respectively). The error bars represent the standard errors of four replicates.
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Figure 3. Activity levels of: GSTs (A); α-EST (B); β-EST (C); and AChE (D) in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti after treatment with P. crispum oil (LC50) for 3, 6 and 24 h. Vertical bars represent standard error (±SE) of six or ten biological samples from different preparations. The asterisk shows statistically significant differences of means among the different groups (Tukey’s HSD test, p < 0.05).
Figure 3. Activity levels of: GSTs (A); α-EST (B); β-EST (C); and AChE (D) in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti after treatment with P. crispum oil (LC50) for 3, 6 and 24 h. Vertical bars represent standard error (±SE) of six or ten biological samples from different preparations. The asterisk shows statistically significant differences of means among the different groups (Tukey’s HSD test, p < 0.05).
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Figure 4. Activity levels of: ACP (A); ALP (B); and MFO (C) in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti after treatment with P. crispum oil (LC50) for 3, 6 and 24 h. Vertical bars represent standard error (±SE) of six biological samples from different preparations. The asterisk shows statistically significant differences of means among the different groups (Tukey’s HSD test, p < 0.05).
Figure 4. Activity levels of: ACP (A); ALP (B); and MFO (C) in 4th instar larvae of the pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) Ae. aegypti after treatment with P. crispum oil (LC50) for 3, 6 and 24 h. Vertical bars represent standard error (±SE) of six biological samples from different preparations. The asterisk shows statistically significant differences of means among the different groups (Tukey’s HSD test, p < 0.05).
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Table
Table 1. Enzyme activity in pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) strains of Ae. aegypti after treatment with LC50 of P. crispum oil.
Table 1. Enzyme activity in pyrethroid-susceptible (MCM-S) and resistant (PMD-R and UPK-R) strains of Ae. aegypti after treatment with LC50 of P. crispum oil.
EnzymeEnzyme Activity (Mean ± SE) *
MCM-SPMD-RUPK-R
GSTs 1
0 hUntreated0.109 ± 0.002 a,A0.204 ± 0.002 a,B0.260 ± 0.003 a,C
24 hUntreated0.116 ± 0.004 a,A0.227 ± 0.004 a,B 0.276 ± 0.006 a,b,C
Control0.112 ± 0.004 a,A0.242 ± 0.006 a,B0.280 ± 0.007 b,C
Treated0.210 ± 0.007 b,A0.442 ± 0.021 b,B0.521 ± 0.007 c,C
α-EST 2
0 hUntreated127.73 ± 3.73 a,A183.82 ± 4.94 a,B195.23 ± 3.79 a,B
24 hUntreated133.10 ± 3.52 a,A178.64 ± 5.35 a,B192.88 ± 3.79 a,B
Control125.92 ± 5.56 a,A174.59 ± 3.40 a,B193.33 ± 4.40 a,C
Treated202.19 ± 5.08 b,A219.44 ± 6.71 b,A215.14 ± 3.67 b,A
β-EST 3
0 hUntreated121.07 ± 3.84 a,A156.67 ± 4.10 a,B188.44 ± 3.44 a,C
24 hUntreated128.42 ± 4.33 a,A158.87 ± 6.04 a,B170.79 ± 4.15 b,B
Control125.28 ± 4.16 a,A157.25 ± 2.83 a,B169.66 ± 2.92 b,C
Treated 203.34 ± 8.31 b,A,B185.03 ± 7.41 b,A212.09 ± 5.99 c,B
AChE 4
0 hUntreated10.96 ± 0.24 a,A10.51 ± 0.33 a,A11.40 ± 0.36 a,A
24 hUntreated12.81 ± 0.42 b,A12.12 ± 0.60 a,A15.06 ± 0.46 b,B
Control12.90 ± 0.46 b,A11.90 ± 0.44 a,A15.27 ± 0.35 b,B
Treated8.65 ± 0.32 c,A8.67 ± 0.37 b,A12.93 ± 0.35 c,B
ACP 5
0 hUntreated71.31 ± 1.34 a,A96.21 ± 1.21 a,C83.29 ± 1.55 a,B
24 hUntreated100.02 ± 4.81 b,A125.55 ± 2.07 b,B101.57 ± 3.14 b,A
Control104.56 ± 4.20 b,A126.39 ± 1.94 b,B103.13 ± 1.60 b,A
Treated121.12 ± 2.52 c,A168.97 ± 2.85 c,B123.35 ± 2.41c,A
ALP 6
0 hUntreated13.21 ± 0.28 a,A12.65 ± 0.54 a,A20.82 ± 0.40 a,B
24 hUntreated7.24 ± 0.33 b,A6.48 ± 0.35 b,A9.44 ± 0.36 b,B
Control7.33 ± 0.23 b,A6.90 ± 0.31 b,A10.38 ± 0.25 b,B
Treated15.54 ± 0.56 c,A13.03 ± 0.55 a,B16.93 ± 0.65 c,A
MFO 7
0 hUntreated0.111 ± 0.003 a,A0.124 ± 0.006 a,A0.158 ± 0.005 a,B
24 hUntreated0.077 ± 0.005 b,A0.075 ± 0.006 b,A 0.115 ± 0.004 b,c,B
Control0.080 ± 0.003 b,A0.076 ± 0.004 b,A0.119 ± 0.008 b,B
Treated0.073 ± 0.003 b,A0.066 ± 0.005 b,A0.093 ± 0.004 c,B
* Values followed by different lowercase letters in a column and uppercase letters in a row are significantly different (Tukey’s HSD test, p < 0.05); 1 GSTs activity was expressed as µmol/min/mg protein; 2 α-EST activity was expressed as nmol/α-naphthol released/min/mg protein; 3 β-EST activity was expressed as nmol/β-naphthol released/min/mg protein; 4 AChE activity was expressed as nmol/min/mg protein; 5 ACP activity was expressed as nmol p-nitrophenol released/min/mg protein; 6 ALP activity was expressed as nmol p-nitrophenol released/min/mg protein; 7 MFO activity was expressed as nmol cytochrome c equivalent unit/mg protein.

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Intirach, J.; Junkum, A.; Lumjuan, N.; Chaithong, U.; Somboon, P.; Jitpakdi, A.; Riyong, D.; Champakaew, D.; Muangmoon, R.; Chansang, A.; et al. Biochemical Effects of Petroselinum crispum (Umbellifereae) Essential Oil on the Pyrethroid Resistant Strains of Aedes aegypti (Diptera: Culicidae). Insects 2019, 10, 1. https://doi.org/10.3390/insects10010001

AMA Style

Intirach J, Junkum A, Lumjuan N, Chaithong U, Somboon P, Jitpakdi A, Riyong D, Champakaew D, Muangmoon R, Chansang A, et al. Biochemical Effects of Petroselinum crispum (Umbellifereae) Essential Oil on the Pyrethroid Resistant Strains of Aedes aegypti (Diptera: Culicidae). Insects. 2019; 10(1):1. https://doi.org/10.3390/insects10010001

Chicago/Turabian Style

Intirach, Jitrawadee, Anuluck Junkum, Nongkran Lumjuan, Udom Chaithong, Pradya Somboon, Atchariya Jitpakdi, Doungrat Riyong, Danita Champakaew, Roongtawan Muangmoon, Arpaporn Chansang, and et al. 2019. "Biochemical Effects of Petroselinum crispum (Umbellifereae) Essential Oil on the Pyrethroid Resistant Strains of Aedes aegypti (Diptera: Culicidae)" Insects 10, no. 1: 1. https://doi.org/10.3390/insects10010001

APA Style

Intirach, J., Junkum, A., Lumjuan, N., Chaithong, U., Somboon, P., Jitpakdi, A., Riyong, D., Champakaew, D., Muangmoon, R., Chansang, A., & Pitasawat, B. (2019). Biochemical Effects of Petroselinum crispum (Umbellifereae) Essential Oil on the Pyrethroid Resistant Strains of Aedes aegypti (Diptera: Culicidae). Insects, 10(1), 1. https://doi.org/10.3390/insects10010001

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