Revista Mexicana Ciencias Agrícolas volume 12 number 1 January 01 - February 14, 2021
Quantification of enzymes related to insecticide resistance in Bemisia tabaci from the state of Sinaloa
Leslie Carnero Avilés1
Ernesto Cerna Chávez1§
José Francisco Rodríguez Rodríguez1
Mariana Beltrán Beache2
Yisa M. Ochoa Fuentes1
Sixto Velarde Félix3
1Postgraduate of Sciences in Agricultural Parasitology-Antonio Narro Autonomous Agrarian University. Roadway Antonio Narro 1923, Col. Buenavista, Saltillo, Coahuila, Mexico. (firstname.lastname@example.org; email@example.com; firstname.lastname@example.org). 2FORDECYT-CULTA, SA de CV. (email@example.com). 3Culiacán Valley Experimental Field-INIFAP. Culiacán-El Dorado Highway km 17.5, Ejido Canán, Pueblo Costa Rica, Culiacán Rosales, Sinaloa. ZC. 80130. (firstname.lastname@example.org).
§Corresponding author: email@example.com.
The whitefly Bemisia tabaci (Gennadius) is one of the most invasive pests and causes direct damage to crop by feeding on the sap and indirect damage by being a vector of more than 100 phytopathogenic viruses. At present its control is based on the use of chemical insecticides, because the populations have been constantly subjected to a high selection pressure. An alternative that contributes to understanding the origin of resistance in a population are biochemical tests, which show the parameter of the detoxifying enzyme present. The objective of the present work was the quantification of these enzymes in B. tabaci in the three main solanaceae producing areas of the north (Guasave, Sinaloa de Leyva, Mochis), center (Culiacán, Navolato, Elota) and south (Concordia, Rosario, Esquinapa) of the state of Sinaloa. Whitefly adults were collected at these sites and the enzymatic levels of α and β esterases, glutathione S-transferases, aceticolinesterases and oxidases were determined, additionally a susceptible laboratory line was used as a reference. The enzymes with the highest presence were α-esterases, β-esterases and oxidases, followed by glutathione S-transferases and acetylcholinesterase. Therefore, it is concluded that resistance to insecticides in B. tabaci in the state of Sinaloa is due to the high content of α - β-esterases and oxidases, while acetylcholinesterase is not a relevant mechanism in the populations of this producing region.
Keywords: Bemisia tabaci, insecticides, resistance.
Reception date: November 2020
Acceptance date: January 2021
The state of Sinaloa is the main national producer of solanaceaes such as tomato, pepper and eggplant, whose combined production value exceeds 18 billion pesos (SIAP, 2019). The trade in these vegetables is highly dependent on the United States market since Sinaloa serves as the main producer and exporter during the winter to the United States of America (FAS-USDA, 2018).
The whitefly Bemisia tabaci is one of the most destructive and invasive pests in the world (GISD, 2020), it has a host range that amounts to more than 1 000 cultivated and wild plants (Oliveira et al., 2001; Simmons et al. , 2008; Abd-Rabou and Simmons, 2010) and causes direct damage to crops by feeding on plant sap and excreting sugary substances on leaves and fruits that damage their quality and promote the development of fungi such as Fumago spp., likewise, causes indirect damage by being a vector of more than 100 phytopathogenic viruses (Horowitz and Ishaaya, 2014).
Excessive populations of B. tabaci can reduce crop yields by up to 50% (Raveesh, 2018), therefore, continuous applications of chemical insecticides are carried out for their control, which caused the development of resistance (Palumbo et al., 2001; Ahmad et al., 2010).
World registries on B. tabaci mention resistance to 64 active ingredients of toxicological groups such as avermectins, neonicotinoids, buprofezin, organophosphates, pyrethroids, carbamates, phenylpyrazoles, chlorinated cyclodiene, butenolids, pyridine azomethine, acaricides and insecticides METI and pyriproxyfen (APRD, 2020).
This is due to a diverse set of resistance mechanisms, which were corroborated in countries that lead the world production of Solanaceae, China (Wang et al., 2018), India (Naveen et al., 2017), Turkey (Erdogan et al. al., 2008), Egypt (Farghaley et al., 2016) and the United States of America (Longhurst et al., 2014). In Mexico, Aguilar-Medel et al. (2007) evaluated the susceptibility of populations from the states of Baja California and Sinaloa to the insecticides acetamiprid, cypermethrin, imidacloprid, pymetrozine and thiamethoxam, being the population from Sinaloa the most resistant, in another study Gutiérrez-Olivares (2007) reports tolerance in populations of San Luis Potosi to imidacloprid and Servin-Villegas et al. (2006) to thiamethoxam and endosulfan in Baja California.
Resistance to insecticides involves mutations in the sites of action, on gene expression, less cuticular penetration, resistance to knockdown, greater storage and excretion, in addition to the production of detoxifying enzymes (Vais et al., 1997; Ahmad et al., 2006; Bass and Field, 2011).
The latter are the main mechanism of resistance, particularly the production of esterases, oxidases and glutathione S-transferans (GST) (Flores et al., 2006). The objective of the present work was the quantification of detoxifying enzymes in B. tabaci in the three main solanaceae producing areas of the state of Sinaloa.
Materials and methods
To carry out this study, three populations of B. tabaci were collected in the producing areas of the north (Guasave, Sinaloa de Leyva, Mochis), center (Culiacán, Navolato, Elota) and south (Concordia, Rosario, Esquinapa) of Solanaceae crops in the open field in the state of Sinaloa, Mexico, in the 2018-2019 agricultural cycle.
In addition, a susceptible line maintained in the laboratory was included, which has been free of selection pressure for a period of more than two years. The captured whitefly adults were collected in plastic containers for their conservation in 70% alcohol and at a temperature of 4 °C in refrigeration. The analyzes were carried out in the Toxicology Laboratory of the Department of Agricultural Parasitology of the Autonomous Agrarian University Antonio Narro.
The protein source was obtained from eight samples, with three repetitions, of 100, 150, 200, 250, 300, 350 and 400 individuals of B. tabaci, using the methodology described by Bradford (1976) modified by Brogdon (1984); Brogdon and Barber (1987). In 2 ml Eppendorf tubes, 1 000 μl of buffer solution (KPO4 0.05 M, pH 7.2) and the different densities of the insect mentioned above were added, to later crush them and make up to 1 ml.
In a 96-well microplate, 20 μl of homogenate, 80 μl of buffer solution and 200 μl of diluted dye (4: 1 dye:water) of Comassie Brilliant Blue (Bio-Rad Kit II) were placed in triplicate for each repetition. The absorbance readings were taken with a 630 nm filter. The values of μg ml-1 of protein comprised in the range of 80 to 120 μg (homogenate carried out with 300 individuals) were calculated.
Determination of enzyme levels
The enzymatic levels of α-esterases and β-esterases were determined with the methodology of Brogdon and Dickinson (1983). In a transparent 96-well microplate, 100 μl of the homogenate was added with 100 μl of the substrate α-naphthyl acetate for α-esterases and β-naphthyl acetate for the case of β-esterases, it was incubated for 10 min and 100 μl of Fast-Blue dye were added to incubate again for 2 min, the absorbances were taken with a 540 nm filter.
For the determination of glutathione S-transferases, the methodology described by Brogdon and Barber (1990) was used for this, in a microplate 100 μl of homogenate, 100 μl of reduced Glutathione as substrate and 100 μl of 1-chloro-2,4’-dinitrobenzene (CDNB) as a dye. The absorbances were recorded with a 340 nm filter, their readings were at time zero (T0) and time five (T5), the difference between the two readings (T5-T0), was considered for the result for the analysis.
Acetylcholinesterase levels were determined with the Brogdon (1988) methodology, applying 100 μl of the homogenate to a microplate, 100 μl acetylcholine-iodysed 3 mm as a substrate and 100 μl of 5,5’-dithio-bis-2-nitrobenzoic acid (DTNB) as a dye. The absorbances were recorded with a 414 nm filter and the readings were taken at time zero (T0) and ten (T10), the difference considered as a result for the analysis.
Finally, the determination of oxidase levels was obtained with the method of Brogdon et al. (1997), adding 100 μl of the homogenate, 200 μl of 3,3’,5,5’- Tetramethyl-Benzidine Dihydrochloride (TBMZ) as substrate and 25 μl of 3% H2O2 as dye, were incubated for 5 min and they took the readings with a 620 nm filter.
Analysis of results
With the absorbance readings obtained, a frequency distribution was made and a resistance threshold was established based on the highest value (absorbance) of the susceptible line. The percentage of resistance was obtained with the number of means that exceeded the resistance threshold and were classified according to Montella et al. (2007) with minor modifications such as: ‘unaltered’ (0-5%), ‘incipiently altered’ (6-30%), ‘moderately altered’ (31-50%), ‘altered’ (51-75%) and ‘very altered’ (> 76%). To know the variation in the enzymatic activity between populations, an Anova and Tukey’s test (p= 0.05) were carried out with the statistical program R version 3.3.1.
Results and discussion
The density of 300 adults reached the required levels of protein (80 to 120 μg), most of their repetitions were located near the threshold, while in the densities of 200 and 400 individuals the absorbance values were distributed in different protein contents. (Figure 1). Bradford (1976) mentions that values outside the range are not reliable for the quantification of enzymatic in tissues; for their part, Dary et al. (1990) report that there is a close relationship between sample size and amount of protein, so values outside this range may present differences in the results obtained.
Figure 1. Protein absorbances in the different densities of B. tabaci homogenate in buffer solution (KPO4 0.05 M, pH 7.2).
In relation to α-esterases and β-esterases, the three field populations behaved homogeneously (Table 1) with means of 3.453, 3.489 and 3.513, for the northern, central and southern populations, respectively; however, they were different from the susceptible line with the lowest mean of 1.488.
Table 1. Means and absorbances of the enzymes α-esterases, β-esterases, from the different solanaceae producing areas in the state of Sinaloa.
Mean ± SD1
Mean ± SD1
Means with different letters present a significant difference (p= 0.05). 1= standard deviation; 2= susceptible line.
The α-esterases and β-esterases can occur separately or together in insects (Bisset, 2002), depending on the toxic interaction, detoxification can be carried out by two mechanisms: catalytic hydrolysis that are not inhibited by organophosphates and are not catalytic as it is inhibited by organophosphates (Costa, 2006).
Previous studies reported that these enzymes confer resistance to insecticides such as pyrethroids (Yang et al., 2001; Flores et al., 2006; Landeros et al., 2010), organochlorines (Bisset et al., 2001), neonicotinoids (Dávila-Medina , 2012), organophosphates and carbamates (Cerna et al., 2013), the high activity of esterases in populations is directly related to resistance to these insecticides, coinciding with previous studies in whiteflies (Byrne and Devonshire, 1993; Kang et al., 2006; Roditakis et al., 2006; Liang et al., 2007; Alon et al., 2008). The oxidases were the enzymes with the highest values in the central zone with a mean of 1.216, followed by the north zone with 1.073 and south with 1.068 (Table 2), the means of the three zones were significantly different with respect to the susceptible line (p-value 0.002), which presented the lowest absorbance value with 0.971.
Table 2. Means and absorbances of the oxidase enzymes and glutathione S-transferases, of the different solanaceae producing areas in the state of Sinaloa.
Mean ± SD1
Mean ± SD1
Means with different letter present significant difference (Tukey> 0.05). 1= standard deviation; 2= susceptible line.
Oxidases act by oxidizing the insecticidal molecule allowing them to enter other enzymatic systems and be expelled (Pimentel et al., 2008), in B. tabaci their activity has been correlated with resistance to neonicotinoids and avermectins (Rauch and Nauen, 2003; Wang and Wu, 2007; Roditakis et al., 2010). In the case of glutathione S-transferases, the susceptible line presented an average of 0.040, when taken as resistance threshold, the highest average was presented in the center population with 0.152, the southern population that showed an average of 0.034, the highest glutathione S-transferases content (Table 2).
In B. tabaci, it disables the action of neonicotinoids (Yang et al., 2016), organophosphates and pyrethroids (Ortelli et al., 2003; Hu et al., 2014) pyriproxyfen (Ma et al., 2010) and diafenthiuron (Zhang et al. al., 2015). These enzymes have been reported to act by conjugating the glutathione thiol group (GSH;\(Lg-glutamyl-L-cysteinyl-glycine) to compounds that have an electrophilic center, thereby eliminating substrates from a cell by increasing its solubility in water (Low et al., 2010).
Finally, the presence of acetylcholinesterases in the populations of Sinaloa and in the laboratory susceptible line did not present significant differences (Table 3). Other studies mention that the resistance of B. tabaci to insecticides (such as methamidophos, chlorpyrifos, phoxim, fenvalerate, avermectin, emamectin benzoate, spinosad, fipronil and imidacloprid) is related to the insensitivity or absence of acetylcholinesterase Kang et al. (2006) since it is considered as non-metabolic resistance which is associated with a mutation in the Acetylcholinesterase site of action (Ramya et al., 2016).
Table 3. Means and absorbances of the acetylcholinesterase enzyme in the different solanaceae producing areas in the state of Sinaloa.
Mean ± SD
Means with different letter present significant difference (Tukey> 0.05). 1= standard deviation; 2= susceptible line.
The main detoxifying mechanisms in the state of Sinaloa were α-esterases, β-esterases and oxidases, presenting a resistance ratio of 100% and being categorized as highly ‘altered’ for the three areas under study. In the case of glutathione S-transferases, only in the central zone it was presented as an important detoxification mechanism by reporting a resistance factor of 66% and a classification of ‘altered’, while in the North and South zones, it is not considered as an important mechanism of resistance to insecticides by showing a null proportion of resistance and being classified as ‘unchanged’.
As regards acetylcholinesterase, it is considered as a mechanism of little relevance for the development of resistance to insecticides in B. tabaci in the state of Sinaloa, as it is not registered in the physical-chemical tests (Table 4).
Table 4. Proportion of resistance (%) in the North, Central and South zones of Sinaloa, in comparison.
Classification according to Montella et al. (2007). a= ‘unaltered’; b= ‘incipiently altered’; c= ‘moderately altered’; d= ‘altered’; e= ‘very altered’.
Understanding these resistance mechanisms is the most important aspect for managing resistance in insect pests (Guo et al., 2014; Zhang et al., 2016; Horowitz, et al., 2020). Kang et al. (2006) point out that the presence or absence of detoxifying enzymes and the difference in resistance to insecticides could be associated with different antecedents of chemical application in the field; the State of Sinaloa, was documented by Cortinas (2000) as one of the areas with the highest consumption of pesticides in the country, representing 30% of national consumption, while 70% is represented by the set of other producing areas (Jalisco-Nayarit-Colima, Sonora-Baja California, Tamaulipas, Michoacán, Chiapas, Veracruz, Tabasco, State of Mexico, Puebla-Oaxaca).
The most frequently used pesticides in Northwest Mexico are dithiocarbamates, bipyridyls, organophosphates, organochlorines, carbamates, pyrethroids and inorganic compounds (Leyva-Morales et al., 2014), which coincides with the high levels of resistance and detoxification mechanisms reported in the present work.
Through the analysis of the enzymatic levels of three populations of B. tabaci from the state of Sinaloa, it was possible to attribute the resistance to insecticides of the North Central and South populations of the state to the enzyme systems of α-esterases, β-esterases and Oxidases in addition to Glutathione S-transferases for the population of the center. These resistance mechanisms coincide with the chemical inputs used for the control of B. tabaci in the state, such as: pyrethroids, organochlorines, neonicotinoids, organophosphates and carbamates. Due to the above, the detection of the origin of resistance and its understanding allows us to take the correct measures and actions to manage physiological resistance and restore sensitivity in populations. Which would reduce the high costs in the control, the losses in the harvests, and the contamination of the environment.
Abd-Rabou, A. S. and Simmons, A. M. 2010. Survey of reproductive host plants of Bemisia tabaci (Hemiptera: Aleyrodidae) in Egypt, including new host records survey of reproductive host plants. Entomological News. 121(5):456-465. Doi: 10.3157/021.121.0507.
Aguilar-Medel, S.; Rodríguez-Maciel, J. C.; Santillán-Ortega, C.; Lagunes-Tejeda, A.; Díaz-Gómez, O. y Martínez-Carrillo, J. L. 2007. Susceptibilidad a insecticidas en dos poblaciones de Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae) biotipo B Colectadas en Baja California y Sinaloa, México. Interciencia. 32(4):266-269. http://www.redalyc.org/ articulo.oa?id=33932410.
Ahmad, M.; Denholm, I. and Bromilow, R. H. 2006. Delayed cuticular penetration and enhanced metabolism of deltamethrin in pyrethroid-resistant strains of Helicoverpa armigera from China and Pakistan. Pest Management Sci. 62(9):805-810. Doi: 10.1002/ps.1225.
Ahmad, M.; Arif, M. I. and Naveed, M. J. 2010. Dynamics of resistance to organophosphate and carbamate insecticides in the cotton whitefly Bemisia tabaci (Hemiptera: Aleyrodidae) from Pakistan. J. Pest Sci. 83(4):409-420. Doi: 10.1007/s10340-010-0311-8.
Alon, M.; Alon, F.; Nauen, R. and Morin, S. 2008. Organophosphates resistance in the B-biotype of Bemisia tabaci (Hemiptera: Aleyrodidae) is associated with a point mutation in an ace1-type acetylcholinesterase and overexpression of carboxylesterase. Insect Biochem. Mol. Biol. 38(10):940-949. Doi: 10.1016/j.ibmb.2008.07.007.
APRD. 2020. Arthropod pesticide resistance database. www.pesticideresistance.com.
Bass, C. and Field, L. M. 2011. Gene amplification and insecticide resistance. Pest Management Sci. 67(8):886-890. Doi: 10.1002/ps.2189.
Bisset, J. A.; Rodríguez, M. M.; Molina, D.; Díaz, C. y Soca, L. A. 2001. Esterasas elevadas como mecanismo de resistencia a insecticidas organofosforados en cepas de Aedes aegypti. Rev. Cubana de Medicina Tropical. 53(1):37-43. http://scielo.sld.cu/scielo.php?script=sci-arttext&pid=S037507602001000100007.
Bisset, J. A. 2002. Uso correcto de insecticidas: control de la resistencia. Rev. Cubana de Medicina Tropical. 54(3):202-219. http://scielo.sld.cu/pdf/mtr/v54n3/mtr05302.pdf.
Bradford, M. M. 1976. A rapid and sensitive method for quantification of microgram quantities of protein utilizing the principles of protein-dye binding. Anal. Biochem. 72(7):248-254. http://www.sciencedirect.com/science/article/pii/0003269776905273.
Brogdon, W. G. and Dickinson, M. C. 1983. A microassay system for measuring esterase activity and protein concentration in small samples and in hig-pressure liquid chromatography eluate fractions. Analytical Biochemistry. 131(2):499-503. https://www.ncbi.nlm.nih.gov/ pubmed/6614483.
Brogdon, W. G. 1984. Mosquito protein microassay-1, protein determinations from small portions of single-mosquito homogenates. Comp. Biochem. Physiol. 79(3):457-459. https://www.ncbi.nlm.nih.gov/pubmed/6509934.
Brogdon, W. G. and Barber, A. M. 1987. Microplate assay of acetylcholinesterase inhibition kinetics in single mosquitoes homogenates. Pest. Biochem. Phisiol. 29(3):252-259. http://www.sciencedirect.com/science/article/pii/0048357587901556.
Brogdon, W. G. 1988. Microassay of acetylcholinesterase activity in small portions of single mosquito homogenates. Comparative Bioch. Physiol. 90(1):145-150. http://www.sciencedirect.com/science/article/pii/0742841388901107.
Brogdon, W. G. and Barber, A. M. 1990. Microplate assay of glutathione s-transferase activity for resistance detection in single mosquito triturates. Comparative Biochemistry and Physiology. 96(2):339-342. http://www.sciencedirect.com/science/article/pii/03050491 90903857.
Byrne, F. J. and Devonshire, A. L. 1993. Insensitivive acetylcholi-nesterase and esterase polymorphism in susceptible andresistant populations of tobacco whitefly Bemisia tabaci (Genn.). Pestic. Biochem. Physiol. 45(1):34-42. Doi: 10.1006/pest.1993.1005.
Brogdon, W. G.; Allister, J. C. and Vulule, J. 1997. Hemeperoxidase activity measured in single mosquitoes identifies individuals expressing an elevated oxidase for insecticide resistance. J. Am. Mosquito Control Association. 13(3):233-237. http://www.biodiversitylibrary.org/ content/part/JAMCA/JAMCA-V13-N3-P23.
Cerna, C. E.; Hernández, B. O.; Landeros, F. J. y Ochoa, F. Y. 2013. Cuantificación de enzimas asociadas a la resistencia de insecticidas en Bactericera cockerelli (Sulc) de la zona papera de Coahuila y Nuevo León, México. Investigación y Ciencia. 21(59):5-12. http://www.redalyc.org/articulo.oa?id=67430113001.
Cortinas, C. 2000. Proyecto para habilitar a México a formular el plan nacional de implementación (PNI) para dar cumplimiento al Convenio de Estocolmo. Cortinas. http://www.pni-mexico.org. 246 p.
Costa, L. G. 2006. Current issues in organophosphate toxicology. Clin Chim Acta. 366(1-2):1-13. Doi: 10.1016/j.cca.2005.10.008.
Dary, O.; Georghiou, G. P.; Parsons, E. and Pasteur, N. 1990. Microplate adaptation of Gomori’s assay for quantitative determination of general esterase activity in single insects. J. Econ. Entomol. 83(6):2187-2192. Doi: 10.1093/jee/83.6.2187.
Dávila-Medina, M. D.; Cerna-Chávez, E.; Aguirre-Uribe, L. A.; García-Martínez, O.; Ochoa-Fuentes, Y. M.; Gallegos-Morales, G. y Landeros-Flores, J. 2012. Susceptibilidad y mecanismos de resistencia a insecticidas en Bactericera cockerelli (Sulc.) en Coahuila, México. Rev. Mex. Cien. Agríc. 3(6):1145-1155. Doi: 10.29312/remexca.v3i6.1365.
Erdogan, C.; Graham, D.; Oktay, M.; Gorman, K. and Denholm, I. 2008. Insecticide resistance and biotype status of populations of the tobacco whitefly Bemisia tabaci (Hemiptera: Aleyrodidae) from Turkey. Crop Protection. 27(3-5):600-605. Doi: 10.1016/j.cropro. 2007.09.002.
Farghaly, S. F. and Dawood, A. I. 2016. Role of cytochrome P450 gene in insecticide susceptibility of the whitefly, Bemisia tabaci (Homoptera, Aleyrodidae) in Egyptian Governorates. Egyptian Scientific Journal of Pesticides. 2(1):53-66.
FAS-USDA. 2018. Foreign Agricultural Service-United States. Departmentof Agriculture. https://www.fas.usda.gov/data/.
Flores, E. A.; Grajales, J. S.; Fernández, I. S.; Ponce, G. G.; Loaiza. M. H. B.; Lozano, S.; Brogdon, W. G.; Black, I. V. W. C. and Beaty, B. 2006. Mechanisms of insecticide resistance in field populations of Aedes aegypti (L.) from Quintana Roo, Souther Mexico. J. Am. Mosquito Control Association. 22(4):672-677. Doi: 10.2987/8756-971X (2006)22[672: MOIRIF] 2.0.CO; 2.
GISD. 2020. Global Invasive Species Database. http://www.issg.org/database.
Guo, L.; Xie, W.; Wang, S.; Wu, Q.; Li, R.; Yang, N.; Yang, X.; Pan, H. and Zhang, Y. 2014. Detoxification enzymes of Bemisia tabaci B and Q: biochemical characteristics and gene expression profiles. Pest Management Science. 70(10):1588-94. Doi: 10.1002/ps.3751.
Gutiérrez-Olivares, M.; Rodríguez-Maciel, J. C.; Llanderal-Cázaresk, C.; Terán-Vargas, A. P.; Lagunes-Tejeda, A. y Díaz-Gómez, O. 2007. Estabilidad de la resistencia a neonicotinoides en Bemisia tabaci (Gennadius), biotipo B de San Luis Potosí, México. Agrociencia. 41(8):913-920. http://www.redalyc.org/articulo.oa?id=30220203010.
Horowitz, A. R. and Ishaaya, I. 2014. Dynamics of biotypes B and Q of the whitefly Bemisia tabaci and its impact on insecticide resistance. Pest. Manag. Sci. 70(10):1568-1572. Doi: 10.1002/ps.3752.
Horowitz, A.; Ghanim, M.; Roditakis, E.; Nauen, R. and Ishaaya, I. 2020. Insecticide resistance and its management in Bemisia tabaci species. J. Pest Sci. 93(12):893-910. Doi: 10.1007/s10340-020-01210-0.
Hu, F.; Dou, W.; Wang, J. J. and Jia, F. X. 2014. Multiple glutathione S ‐ transferase genes: identification and expression in oriental fruit fly, Bactrocera dorsalis. Pest Manag. Sci. 70(2):295-303. Doi: 10.1002/ps.3558.
Kang, C. Y.; Wu, G. and Miyata, T. 2006. Synergism of enzyme inhibitors and mechanisms of insecticide resistance in Bemisia tabaci (Gennadius) (Hom, Aleyrodidae). J. Appl. Entomol. 130(6-7):377-385. DOI: 10.1111/j.1439-0418.2006.01075.x
Landeros, J.; Ail, C.; Cerna, E.; Ochoa, Y.; Guevara, L. y Aguirre, L. 2010. Susceptibilidad y mecanismos de resistencia de Tetranychus urticae en rosal de invernaderos. Rev. Colomb. Entomol. 36(1):5-9. http://www.scielo.org.co/pdf/rcen/v36n1/v36n1a02.pdf.
Leyva-Morales, J. B.: García-Parra, L. B.; Bastidas-Bastidas, P. D.; Astorga-Rodríguez, J. E.; Bejarano-Trujillo, J.; Cruz-Hernández, A.; Martínez-Rodríguez, I. E. y Betancourt-Lozano, M. 2014. Uso de plaguicidas en un valle agrícola tecnificado en el noroeste de México. Rev. Inter. Contaminación Ambiental. 30(3):247-261. http://www.scielo.org.mx/ scielo.php?script=sci-arttext&pid=S018849992014000300002&lng=es&nrm=iso.
Liang, P.; Cui, J. Z.; Yang, X. Q. and Gao, X. W. 2007. Effects of host plants on insecticide susceptibility and carboxylesterase activity in Bemisia tabaci biotype B and greenhouse whitefly, Trialeurodes vaporariorum. Pest Management Sci. 63(4):365-371. Doi: 10.1002/ps.1346.
Longhurst, C.; Babcock, J. M.; Denholm, I.; Gorman, K.; Thomas, J. D. and Sparks, T. C. 2013. Cross‐resistance relationships of the sulfoximine insecticide sulfoxaflor with neonicotinoids and other insecticides in the whiteflies Bemisia tabaci and Trialeurodes vaporariorum. Pest Management Sci. 69(7):809-813. Doi: https://doi.org/10.1002/ps.3439.
Low, W. Y.; Feil, S. C.; Ng, H. L.; Gorman, M. A.; Morton, C. J.; Pyke, J. and Gooley, P. R. 2010. Recognition and detoxification of the insecticide DDT by Drosophila melanogaster glutathione S-transferase D1. J. Mol. Biol. 399(3):358-366. Doi: https://doi.org/10.1016/j.jmb.2010.04.020.
Ma, W.; Li, X.; Dennehy, T. J.; Lei, C.; Wang, M.; Degain, B. A. and Nichols, R. L. 2010 b. Pyriproxyfen resistance of Bemisia tabaci (Homoptera: Aleyrodidae) biotype B: metabolic mechanism. J. Econ. Entomol. 103(1):158-165. Doi: 10.1603/ec09122.
Montella, I. R.; Martins, A. J.; Fernández, V.; Pereira, L. B.; Braga, I. A. y Valle, D. 2007. Insecticide resistance mechanism of Brazilian Aedes aegyti populations from 2001-2004. Am. J. Trop. Med. Hygiene. 77(3):467-477. Doi: https://doi.org/10.4269/ ajtmh.2007.77.467.
Naveen, N. C.; Chaubey, R.; Kumar, D.; Rebijith, K. B.; Rajagopal, R.; Subrahmanyam, B. and Subramanian, S. 2017. Insecticide resistance status in the whitefly, Bemisia tabaci genetic groups Asia-I, Asia-II-1 and Asia-II-7 on the Indian subcontinent. Scientific Reports, 7(1):1-15. Doi: 10.1038/srep40634.
Oliveira, M. R. V.; Henneberry, T. J. and Anderson, P. 2001. History, current status, and collaborative research projects for Bemisia tabaci. Crop Prot. 20(9):709-723. Doi: 10.1016/s0261-2194(01)00108-9.
Ortelli, F.; Rossiter, L. C.; Vontas, J.; Ranson, H. and Hemingway, J. 2003. Heterologous expression of four glutathione transferase genes genetically linked to a major insecticide-resistance locus from the malaria vector Anopheles gambiae. Biochem. J. 373(3):957-63. Doi: 10.1042/BJ20030169.
Palumbo, J.; Horowitz, A. and Prabhaker, N. 2001. Insecticidal control and resistance management for Bemisia tabaci. Crop Protection. 20(9):739-765. Doi: 10.1016/S0261-2194(01)00117-X.
Pimentel, M. A. G.; Antonino, F. L. R.; Duarte, B. M. y Humberto, S. 2008. Resistance of stored product insects to phosphine. Pesq. Agropec. Bras. 43(12):1671-1676. Doi: 10.1590/S0100-204X2008001200005.
Ramya, S. L.; Venkatesan, T.; Srinivasa, M. K.; Jalali, S. K. and Verghese, A. 2016. Detection of carboxylesterase and esterase activity in culturable gut bacterial flora isolated from diamondback moth, Plutella xylostella (Linnaeus), from India and its possible role in indoxacarb degradation. Brazilian J. Microbiol. 47(2):327-336. Doi: 10.1016/j.bjm. 2016.01.012.
Rauch, N. and Nauen, R. 2003. Identification of biochemical markers linked to neonicotinoid cross resistance in Bemisia tabaci (Hemiptera: Aleyrodidae). Arch. Insect Biochem. Physiol. 54(5):165-176. Doi: 10.1002/arch.10114.
Raveesh, K. G. and Gangwar, C. 2018. Lifecycle, distribution, nature of damage and economic importance of whitefly, Bemisia tabaci (Gennadius). Acta Scientific Agric. 2(4):36-39. https,://actascientific.com/ASAG/ASAG-02-0064.php.
Roditakis, E.; Morou, E.; Tsagkarakou, A.; Riga, M.; Nauen, R.; Paine, M. and Vontas, J. 2010. Assessment of the Bemisia tabaci CYP6CM1vQ transcript and protein levels in laboratory and field-derived imidacloprid-resistant insects and cross-metabolism potential of the recombinant enzyme. Insect Sci. 18(1):23-29. Doi:10.1111/j.1744-7917.2010.01384.x.
Roditakis, E.; Tsagkarakou, A. and Vontas, J. 2006. Identification of mutations in the para sodium channel of Bemisia tabaci from Crete, associated with resistance to pyrethroids. Pesticide Biochem. Physiol. 85(3):161-166. Doi: 10.1016/j.ijpara.2008.12.006.
Servín-Villegas, R.; García-Hernández, J. L.; Murillo-Amador, B.; Tejas, A. and Martínez-Carrillo, J. L. 2006. Estability of insecticide resistance of silverleaf whitefly (Homoptera: Aleyrodidae) in the absence of selection pressure. Folia Entomol. Mex. 45(1):27-34. http://www.redalyc.org/articulo.oa?id=42445104.
SIAP. 2019. Servicio de Información Agroalimentaria y Pesquera. https://nube.siap.gob.mx/ cierreagricola/.
Simmons, A. M.; Harrison, H. F. and Ling, K. S. 2008. Forty-nine new host plant species for Bemisia tabaci (Hemiptera: Aleyrodidae). Entomol. Sci. 11(4):385-390. Doi: 10.1111/j.1479-8298.2008.00288.x.
Vais, H.; Williamson, M. S.; Hick, C. A.; Eldursi, N.; Devonshire, A. L. and Usherwood, P. N. 1997. Functional analysis of a rat sodium channel carrying a mutation for insect knock-down resistance (kdr) to pyrethroids. FEBS Lett. 413(2):427-332. Doi: 10.1016/s0014-5793(97)00931-9.
Wang, L. and Wu, Y. 2007. Cross-resistance and biochemical mechanisms of abamectin resistance in the B-type Bemisia tabaci. J. Appl. Entomol. 131(2):98-103. Doi:10.1111/j.1439-0418.2006.01140.x.
Wang, R.; Wang J. D.; Che, W.; Sun, Y.; Li, W. and Luo, Ch. 2018. Characterization of field-evolved resistance to cyantraniliprole in Bemisia tabaci MED from China. J. Integrative Agric. 18(11):2571-2578. Doi: https://doi.org/10.1016/S2095-3119(19)62557-8.
Yang, X.; Margolies, D. C.; Zhu, K. Y. and Buschman, L. L. 2001. Host plant-induced changes in detoxification enzymes and susceptibility to pesticides in the two spotted spider mites (Acari: Tetranychidae). J. Econ. Entomol. 94(2):381-387. Doi: 10.1603/0022-0493-94.2.381.
Yang, X.; He, C.; Xie, W.; Liu,Y.; Xia, J.; Yang, Z.; Guo, L.; Wen, Y.; Wang, S.; Wu, Q.; Yang, F.; Zhou, X. and Zhang, Y. 2016. Glutathione S-transferases are involved in thiamethoxam resistance in the field whitefly Bemisia tabaci Q (Hemiptera: Aleyrodidae). Pesticide Biochem. Physiol. 134:73-78. Doi: 10.1016/j.pestbp.2016.04.003.
Zhang, B.; Kong, F. and Zeng, X. 2015. Detoxification enzyme activity and gene expression in diafenthiuron resistant whitefly, Bemisia tabaci. J. Agricultural Sci. 7(9):66-76. Doi:10.5539/jas.v7n9p66.
Zhang, S.; Zhang, X.; Shen, J.; Mao, K.; You, H. and Li, J. 2016. Susceptibility of field populations of the diamondback moth, Plutella xylostella, to a selection of insecticides in Central China. Pesticide Biochem. Physiol. 132:38-46. Doi: 10.1016/j.pestbp.2016.01.007.