Comparison of Biochemical and Molecular Tests for Detecting Insecticide Resistance Due to Insensitive Acetylcholinesterase in Culex quinquefasciatus 1

Article (PDF Available)inJournal of the American Mosquito Control Association 28(4):323-6 · December 2012with24 Reads
DOI: 10.2987/12-6280R.1 · Source: PubMed
Abstract
Insecticide resistance to organophosphates and carbamates can be the result of changes in acetylcholinesterase activity conferred by the ACE-1 mutation. Detection of this altered target site mutation is important in guiding informed decisions for resistance management. In this study we compared a competitive enzyme assay with a polymerase chain reaction assay utilizing a restriction enzyme. Both assays detected the ACE-1 mutation in Culex quinquefasciatus and agreement was 100%. The costs and benefits of each assay are presented.

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Comparison of Biochemical and Molecular Tests for Detecting
Insecticide Resistance Due to Insensitive Acetylcholinesterase in
Culex quinquefasciatus
Author(s): Mariah L. Scott and Janet C. McAllister
Source: Journal of the American Mosquito Control Association, 28(4):323-326.
2012.
Published By: The American Mosquito Control Association
DOI: http://dx.doi.org/10.2987/12-6280R.1
URL: http://www.bioone.org/doi/full/10.2987/12-6280R.1
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SCIENTIFIC NOTE
COMPARISON OF BIOCHEMICAL AND MOLECULAR TESTS FOR
DETECTING INSECTICIDE RESISTANCE DUE TO INSENSITIVE
ACETYLCHOLINESTERASE IN CULEX QUINQUEFASCIATUS
1
MARIAH L. SCOTT
AND
JANET C. M
C
ALLISTER
2
Arbovirus Diseases Branch, Division of Vector-Borne Diseases, Centers for Disease Control and Prevention,
3150 Rampart Road, Fort Collins, CO 80521
ABSTRACT. Insecticide resistance to organophosphates and carbamates can be the result of changes in
acetylcholinesterase activity conferred by the ACE-1 mutation. Detection of this altered target site mutation
is important in guiding informed decisions for resistance management. In this study we compared a
competitive enzyme assay with a polymerase chain reaction assay utilizing a restriction enzyme. Both assays
detected the ACE-1 mutation in Culex quinquefasciatus and agreement was 100%. The costs and benefits of
each assay are presented.
KEY WORDS Culex quinquefasciatus, acetylcholinesterase, organophosphate resistance, ACE-1 mutation
Because of biological fitness costs usually
associated with insensitivity of target sites to
insecticides, insects carrying modified target site
gene(s) generally remain at a low frequency in the
absence of selection pressure. Remarkably, it is
only minor changes in the gene(s), often a single
base change, which make target proteins (enzyme
or receptor) insensitive to insecticides (Chaudhry
and MacNicoll 1998). The gene, ACE-1, is
present worldwide and causes organophosphate
(OP) and carbamate resistance. A high level of
acetylcholinesterase (AChE)-1 protein insensitiv-
ity or resistance displayed in Culex pipiens L. is
due to a single amino acid substitution, G119S, a
mutation in the 3rd exon of the ACE-1 gene,
leading to the replacement of a glycine (GGC,
susceptible allele) by a serine (AGC) (Weill et al.
2003, 2004). This mutation is associated with
reduced susceptibility to OP insecticides, modifi-
cations of the catalytic properties of AChE-1, and
a high fitness cost (Weill et al. 2003). Under-
standing the underlying resistance mechanisms is
important in making informed decisions on
alternative control strategies. In our current
economic situation, with funding and programs
being cut, it is important to have access to a
variety of different testing methods for detecting
mechanisms of resistance depending on one’s
budget, personnel, and time. In this study we
compare a biochemical assay used in our
laboratory with a molecular assay, both of which
detect resistance due to insensitive acetylcholin-
esterase. We evaluated not only the efficacy of
each assay, but also calculated the time, cost, and
skill level each assay required.
Culex quinquefasciatus Say egg rafts were
collected in Harris County, TX from 2004 through
2008. Mosquitoes were reared in incubators at
27.5uC, 80–85% RH, and 14 : 10 light : dark cycle.
They were aspirated into 1.5-ml microtubes 1–2 wk
after they had emerged and stored in a 280uC
freezer (Thermo Scientific, Waltham, MA) for
future use. Fifteen mosquitoes from each collec-
tion site throughout Harris County were tested,
for a total sample size of 300. In order to compare
the competitive enzyme assay to the polymerase
chain reaction (PCR) assay, the same mosquitoes
were used for both tests. For the PCR assay and
sequencing, all 6 legs were removed from the
females, and the genomic DNA was extracted with
the use of DNAzolH (Molecular Research Center
Inc., Cincinnati, OH). The head and body were
homogenized in potassium phosphate (KPO
4
)
buffer for the enzyme assay.
The insensitive acetylcholinesterase assay fol-
lowed the procedures described by Brogdon
(1988), and outlined in McAllister et al. (2012).
The authors previously found that 10 min was
not a sufficient amount of ti me to see the
necessary color change (McAllister and Scott,
unpublished data). The protocol was modified,
and the microplate was read immediately (T
0
)on
a spectrophotometer with the use of a 414-nm
filter, and then the microplate was stored in the
refrigerator and read again after 24 h (T
24
)
(McAllister et al. 2012). The T
0
reading was
subtracted from the T
24
reading for statistical
analysis. Each mosquito sample was run in
triplicate on the microplate.
1
The views of the authors do not necessarily reflect
the position of the Centers for Disease Control and
Prevention.
2
To whom correspondence should be addressed.
Journal of the American Mosquito Control Association, 28(4):323–326, 2012
Copyright
E
2012 by The American Mosquito Control Association, Inc.
323
Mosquito genomic DNA was extracted from
the legs with the use of DNAzol. The legs from
individual mosquitoes were homogenized in 25
m
l
of DNAzol with the use of a Kontes pellet pestle
cordless motor with disposable pestles and the pestles
were rinsed with 25
m
l of DNAzol. The homogenate
was centrifuged for 2 min at 10,621 3 g to remove
insoluble tissue. A volume of 48
m
l of the resulting
viscous supernatant was transferred to a new 1.5-
ml microtube. The DNA was precipitated from
the homogenate by adding 25
m
lof100% ethanol
(CAS 64-17-5). The samples were mixed
by inversion to ensure a homogenous solution
and incubated at room temperature for 3 min. The
samples were centrifuged for 4 min at 14,000 rpm.
The DNA precipitate was washed twice with 500
m
l
of 75% ethanol, allowed to air dry for 5–15 sec,
and then resuspended in 25
m
lofdH
2
O (CAS
7789-20-0).
The genomic DNA extracted from the legs was
PCR amplified with the degenerated primers
Moustdir1 59-CCG GGN GCS ACY ATG TGG
AA-39 and Moustrev1 59-ACG ATM ACG TTC
TCY TCC GA-39 for 30 cycles of amplification
(94uC for 30 sec, 52uC for 30 sec, and 72uCfor
1 min), a procedure developed by Weill et al. (2004).
The PCR fragments were digested with AluI
restriction enzyme according to the manufactur-
er’s instructions and fractionated on a 2%
agarose gel. The 194 base-pair (bp) fragment
amplified by PCR on genomic DNA is cut by the
Alu1 restriction enzyme only in resistant mosqui-
toes. Frequencies of ACE-1 alleles were calculat-
ed as described in McAllister et al. (2012).
In order to check the identity of the amplified
fragments, sequences were performed directly on
PCR products of all 300 mosquitoes using the Big
DyeH terminator kit (Applied Biosystems, Carls-
bad, CA). The sequencing procedure outlined by
Labbe et al. (2007) was followed. Two specific
primers: CpEx3dir 59-CGA CTC GGA CCC
ACT CGT-39 and CpEx3rev 59-GAC TTG CGA
CAC GGT ACT GCA-39 generated a 457-bp
fragment that amplified part of exon 3 of the
ACE-1 gene, including position 119.
The concordance between insensitive AChE
and the presence of the G119S mutation, as
detected by the PCR test and enzyme assay, was
100% (n 5 300). This was confirmed by sequenc-
ing of the PCR product. The AChE activity
measured biochemically in adults is therefore due
to the ACE-1 mutation. Genomic DNA amplified
a 194-bp fragment that is undigested by Alu1for
susceptible homozygous mosquitoes, and cut into
2 fragments (74 bp and 120 bp) for resistant
homozygous individuals. Heterozygous individu-
als displayed a combined pattern, with 3 frag-
ments (74 bp, 120 bp, and 194 bp).
For the competitive enzyme assay, absor-
bance values were designated that signified
susceptible homozygous, resistant homozygous,
and heterozygous individuals by comparing the
results to the restriction enzyme and the
sequenced produ ct. An absorbance reading of
0–0.44 nm indicated a susceptible homozygous
individual, .2.45 nm a resistant homozygous
individual, and a reading fro m 0.45–2.44 nm
specified a heterozygous indivi dual.
As seen in Table 1, of the 300 mosquitoes
tested, 13 (4%) were resistant homozygous, 80
(27%) were susceptible homozygous, and 207
(69%) were heterozygous. Figure 1 shows absor-
bance (nm) levels detected in the tested mosqui-
toes. There was a wide range, 1.9 nm, of
absorbance values for the heterozygous mosqui-
toes with the homozygous mosquito values
spanning a range of 0.4 nm.
Competitive biochemical assays help detect
specific resistance mechanisms in individual
insects and can be used to estimate the frequency
of resistance genes in populations. Propoxur is
used in this assay to inhibit the activity of the
sensitive (i.e., susceptible) AChE, allowing the
detection of the altered enzyme when it is present.
In resistant mosquitoes, the insecticide fails to
inhibit AChE. The number of alleles of insensitive
AChE is greater as the yellow color darkens;
however, it can be difficult to determine exact
cut-off points with the naked eye.
The costs, time, and skill set needed for each
test vary significantly. For the enzyme assay, the
cost per sample is $0.07 (assuming a 96-well
plate); this includes the materials (acetone,
ATCH, DTNB, propoxur, and KPO
4
buffer)
and the consumables (microtiter plates, 1.5-ml
tubes, and pestles). The initial equipment (spec-
trophotometer and pH meter) costs can range
from $5,000 to $30,000. It is important to note
that a spectrophotometer is not necessarily
needed, as it is possible, using the naked eye,
to distinguish roughly between resistant ho mo-
zygous, heterozygous, and susceptible homozy-
gous individuals by their discrete absorbance
classes.
For the molecular assay, the cost per sample is
$1.66 (assuming a 96-well plate), which includes
the materials (Taq, DNAzol, ladder, primers,
dNTPs, agarose, EtBR, ethanol, etc.) and the
consumables (PCR plates, cap strips, 1.5-ml
tubes, and pestles). The initial equipment (centri-
fuge, thermal cycler, gel system, transilluminator,
and hot water bath) costs are $40,000+.
The enzyme assay (30 mosquitoes per plate)
requires 45 min, which includes making the stock
chemicals; homogenizing the mosquitoes; loading
the plates with homogenate, ATCH, and DNTB;
and running the plates on the spectrophotometer.
The plates must also be incubated for 24 h in the
refrigerator in order to see the necessary color
change. In total, the enzyme assay requires 45 min
to complete, and a 24-h incubation period. For
the molecular assay (96 mosquitoes total), one
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has to extract the DNA (2 h), run the PCR
program (2 h), incubate the PCR product with
the restriction enzyme (2–4 h), and run a gel (30–
45 min), which totals 6K to 7L h.
Training to conduct both assays is straightfor-
ward, but with the molecular assay it is advised that
the newly trained individual be evaluated to assure
proper technique. All samples are run in triplicate
in the enzyme assay, so outlying data points due to
pipetting errors or a large piece of homogenate can
be disregarded. If an error arises when running the
gel or sequencing the PCR product, it is necessary
to start the assay over minus the extraction step.
Also of importance is that the enzyme assay is not
species specific, but can be run on any species of
mosquito with no changes in chemicals or proce-
dures. However, it is advised that the absorbance
values indicating resistant/susceptible homozygous
and heterozygous individuals on each species and
on an individual-laborat ory basis be determined.
Table 1. Frequency of single-nucleotide polymorphism mutations associated with ACE-1 mutation in Culex
quinquefasciatus from Harris County, TX. Susceptible homozygous individuals are Gly/Gly; resistant homozygous
individuals are Ser/Ser, and heterozygotes are Gly/Ser.
Year Sampled area Gly/Gly Gly/Ser Ser/Ser N
Frequency resistant
allele (serine)
95% confidence
index
2004 109 3 12 0 15 0.400 0.202
2005 205 4 11 0 15 0.367 0.186
225 4 11 0 15 0.367 0.186
2006 51 1 13 1 15 0.500 0.253
91 4 11 0 15 0.367 0.186
93 2 13 0 15 0.433 0.219
225 7 7 1 15 0.300 0.152
904 3 12 0 15 0.400 0.202
936 9 4 2 15 0.267 0.135
2007 51 5 9 1 15 0.367 0.186
93 1 13 1 15 0.500 0.253
205 7 8 0 15 0.267 0.135
904 4 8 3 15 0.467 0.236
936 4 10 1 15 0.400 0.202
2008 51 0 15 0 15 0.500 0.253
55 0 14 1 15 0.533 0.270
93 7 8 0 15 0.267 0.135
109 10 5 0 15 0.167 0.084
225 2 12 1 15 0.467 0.236
604 3 11 1 15 0.433 0.219
Total 80 207 13 300 0.388 0.044
Fig. 1. Frequency of absorbance values (nm) detected with competitive enzyme assay of Culex quinquefasciatus
from Harris County, TX.
D
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The restriction enzyme assay was devised to
detect the ACE-1 mutation in Anopheles gambiae
Giles and Cx. pipiens, 2 mosquitoes belonging to
different genera. The primers have also been
shown to work on An. albimanus Wiedemann and
Cx. quinquefasciatus. This indicates that it prob-
ably has a broad applicability within the Culic-
idae family (Weil et al. 2004). The assay has also
been adapted to work on An. funestus Giles, An.
arabiensis Patton, and An. quadriannulatus (Theo-
bald) with the use of a different set of primers
(Djegbe et al. 2011, Yewhalaw et al. 2011).
Primers must be developed for each species of
interest if not already available.
In conclusion, both assays detected ACE-1
genotypes with 100% agreement. The enzyme
assay requires less time by personnel, costs less,
employs a simpler technique than the PCR assay,
and is not species specific. Either assay would be an
invaluable option for any laboratory, and would
greatly aid resistance management decisions.
The authors wish to thank the personnel of the
Harris County Public Health and Environmental
Services, Mosquito Control Division, who con-
tributed to collecting and shipping eggs to the
Centers for Disease Control and Prevention. In
particular we wish to thank Pamela Stark and
Nathan Vessey. They would also like to thank
Michael Hayashi for his work on the study.
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  • [Show abstract] [Hide abstract] ABSTRACT: Background Resistance to the carbamate insecticide bendiocarb is emerging in Anopheles gambiae populations from the city of Yaoundé in Cameroon. However, the molecular basis of this resistance remains uncharacterized. The present study objective is to investigate mechanisms promoting resistance to bendiocarb in An. gambiae populations from Yaoundé. Methods The level of susceptibility of An. gambiae s.l. to bendiocarb 0.1 % was assessed from 2010 to 2013 using bioassays. Mosquitoes resistant to bendiocarb, unexposed and susceptible mosquitoes were screened for the presence of the Ace-1R mutation using TaqMan assays. Microarray analyses were performed to assess the pattern of genes differentially expressed between resistant, unexposed and susceptible. ResultsBendiocarb resistance was more prevalent in mosquitoes originating from cultivated sites compared to those from polluted and unpolluted sites. Both An. gambiae and Anopheles coluzzii were found to display resistance to bendiocarb. No G119S mutation was detected suggesting that resistance was mainly metabolic. Microarray analysis revealed the over-expression of several cytochrome P450 s genes including cyp6z3, cyp6z1, cyp12f2, cyp6m3 and cyp6p4. Gene ontology (GO) enrichment analysis supported the detoxification role of cytochrome P450 s with several GO terms associated with P450 activity significantly enriched in resistant samples. Other detoxification genes included UDP-glucosyl transferases, glutathione-S transferases and ABC transporters. Conclusion The study highlights the probable implication of metabolic mechanisms in bendiocarb resistance in An. gambiae populations from Yaoundé and stresses the need for further studies leading to functional validation of detoxification genes involved in this resistance.
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