Introduction:-
Insects have long been formidable adversaries
in agriculture, posing significant challenges to crop production and human
health. Historically, the use of chemical pesticides has been a primary method
for controlling insect pests and mitigating their impact. However, over time,
insects have demonstrated remarkable adaptability, evolving mechanisms to
withstand the effects of these control measures. This phenomenon, known as
"insect resistance," presents a pressing concern for farmers, public
health officials, and environmentalists alike. Insecticide resistance is a
result of accelerated microevolution. According to Insecticide Resistance
Action Committee (IRAC) insecticide resistance is a genetically heritable
reduction in the sensitivity of a target pest population to an insecticide that
arises from exposure of the population to the insecticide in the field, with
the potential to lead to control failure. Insecticide resistance is the added
ability to withstand an insecticide acquired by breeding of those individuals
which survive exposures to that particular insecticide sufficient to wipe out
the whole colony (Hoskins and Gordon ,1956).
Under
selection pressure the fittest survive, multiply and spread. It results from
the survival and spread of resistant insect genotypes that have the capability
to endure insecticide selection pressures in the environment. Insect
development of resistance to insecticides is an inevitable consequence of
insecticide use for pest control. When the frequency of resistant phenotypes
increases to a certain level in field populations, control efficacy with the
concerned insecticide becomes economically unacceptable. But poor efficacy
under field conditions is not always due to insecticide resistance. Amongst
other factors, the quality of technical grade material used, the formulation,
the application dose and the method of application can also play an important
role in impairing field control. However, if resistance is the major factor,
field control failure is inevitable, irrespective of quality, quantity or
methods of application. Thus resistance eventually is the single most important
phenomenon that threatens sustainable pest management. It is therefore
important to detect resistance when it is at incipient levels and monitor its
increase and geographical spread so that appropriate measures can be initiated
to curtail its increase.
Insecticide resistance monitoring is the
systematic process of assessing and tracking changes in the susceptibility of
insect populations to insecticides over time. It involves collecting data on
the efficacy of insecticides against target pest species and analysing this
information to detect trends and patterns indicative of resistance development.
Resistance monitoring is an important element of IRM plans for insecticides (as
well as for genetically modified crops – Bt
crops). Monitoring results can provide an early warning of resistance
evolution, advance the understanding of factors that drive resistance
evolution, document the effectiveness of IRM strategies, and provide relevant information
to guide implementation of effective pest management practices. Early detection
of resistance in a target pest population can facilitate early interventions
and extend product life (durability), thereby benefiting growers and
agricultural production systems. Additionally, resistance monitoring can
identify field-evolved resistance locally prior to broader spread and guide
better management practices in the affected and non-affected area. The major
objectives of resistance detection and monitoring must be to eventually ensure
effective and sustainable pest management.
Importance of insect resistance
monitoring:-
Resistance monitoring is crucial for several
reasons:
1. Preservation of Pest Control Effectiveness:
Monitoring resistance allows us to detect changes in the susceptibility of pest
populations to control measures such as insecticides. By identifying resistance
early, we can adapt pest management strategies to maintain their effectiveness
and prevent the escalation of resistance.
2. Timely Intervention: Early detection of
resistance enables prompt intervention, which can help prevent the spread of
resistant pest populations and minimize their impact on agricultural
productivity, public health, and environmental sustainability.
3. Informed Decision-Making: Monitoring provides
valuable data that can inform decision-making in pest management programs. By
understanding resistance trends and patterns, stakeholders can make informed
decisions about the selection and deployment of control measures, optimizing
their efficacy and cost-effectiveness.
4. Resistance Management Strategies: Monitoring
resistance allows for the development and implementation of targeted resistance
management strategies. By identifying areas or pest populations with high
levels of resistance, resources can be directed towards alternative control
methods or integrated pest management practices to mitigate the impact of
resistance.
5. Sustainable Agriculture and Environmental
Protection: Resistance monitoring supports sustainable agriculture by promoting
the judicious use of pesticides and minimizing environmental impacts. By
reducing reliance on chemical pesticides and incorporating alternative control
methods, resistance monitoring contributes to environmental protection and
biodiversity conservation.
Overall, resistance monitoring is essential
for safeguarding agricultural productivity, public health, and environmental
sustainability in the face of evolving pest challenges. By staying vigilant and
proactive in our efforts to monitor and manage resistance, we can ensure the
continued effectiveness of pest control measures and mitigate the impact of
resistant pest populations.
Scope of Insecticide Resistance Monitoring
Programs:
There are two types of insect resistance
monitoring programme viz.Proactive and Reactive.
A proactive monitoring program is intended to
detect early signs of resistance development in response to the use of a given
product, which would allow for evaluation and if necessary, change of
management practices that could increase product durability and or limit the
spread of the resistance. On the other hand, a reactive monitoring program
focuses on determining whether resistance evolution is responsible for
compromised field performance, followed by the implementation of a suitable
mitigation strategy.
Proactive Monitoring Programs
Proactive resistance monitoring measures
changes in insect susceptibility (at a population level) to a given active
insecticidal ingredient (a.i.) or tracks performance of a product over time at
a given location before performance failure occurs. Therefore, proactive
monitoring programs involve systematic testing of field collected populations
(preferably using IRAC approved methods)
and/or systematic field surveys of product performance. A proactive
monitoring program consists of two parts: establishing baseline susceptibility
or product performance baseline, and subsequent systematic monitoring of insect
susceptibility (resistance) or product performance with comparison to the
baseline data. Establishing baseline susceptibility involves measuring the
initial variability in sensitivity of a given insect population to an a.i. or
determining the field performance of an a.i. prior to large scale exposure of
target insect field populations (i.e., commercialization in major crop production
areas). After baseline establishment, the susceptibility of field populations
is systematically monitored using methods in line with those used for
establishing the baseline and or insecticide performance (i.e., efficacy) is
evaluated over time in regions of high use to identify deviations from the
baseline.
Reactive Monitoring Programs
Reactive resistance monitoring relies on
detection and report of reduced efficacy of an insecticide in the field. Control
failure or unexpected damage reports by growers, crop consultants and extension
advisors can be collected and investigated. The reporting and documentation of
these cases enable the identification of potential resistant populations of the
target pest and can trigger remedial actions (e.g., altering product use
patterns, best management practices, and recommending additional pest
management tools). Ideally, insects are sampled from the area with a control
failure and tested using an appropriate bioassay method to confirm resistance
by comparing to previously established sensitivity baselines. The confirmation
of resistance may lead to additional management and/or mitigation measures at
the regional level.
Case study:
Ø The baseline-susceptibility tests conducted on P. brassicae in 11
diferent feld populations from Meghalaya revealed that Smit population strains
seemed to show less tolerance to both the Bt Cry toxins (Cry1C and Cry2Ab).
Ø Compared to the Cry1C toxin, Cry2Ab was found more potent against P.
brassicae.
Ø The median lethal concentrations, LC50 72 h, varied from 0.535 to 1.725 µg/ml
for Cry2Ab and 0.546–1.803 µg/ml for Cry1C toxin.
Ø The screening using leaf-dip bioassay resulted in a tolerance ratio of
3.3-fold and 3.2-fold for Cry1C and Cry2Ab, respectively.
Ø The most tolerant strains of P. brassicae from Umiam and Pepbah regions
were observed to show discriminating concentrations of 19.30 µg/ml for Cry1C
and 24.03 µg/ml for Cry2Ab (LC99, 72 h).
Protocols for Resistance Monitoring
To evaluate the resistance in insect a
protocol was given which include three steps viz. sampling and rearing
of technique, bioassay and statistical analysis of bioassay data.
Sampling and Rearing of technique :
Sampling and rearing techniques are specific
for each species. The insect are sampled from the field and reared and are used for bioassay.
Bioassay:
The bioassay methods should closely simulate
field conditions to ensure predictability of control efficacy in the field from
data obtained through lab-measured resistance. However, based on several
practical considerations resistance detection and monitoring methods are
developed in such a manner so as to ensure that the bioassays are reliable,
replicable, consistent and robust enough not to be influenced by variations in
operator skills, materials, extraneous factors and handling procedures. For
example, direct bioassays on simulated field conditions using larvae of varying
resistant levels, were designed and used to monitor resistance. But, due to
space and operational constraints, such assays can be performed only on a
limited sample size. Over the past two decades, bioassay methods based on
leaf-dip, larval dip, topical application and vial-residue were developed as
viable alternatives to simulated field conditions.
Commonly used bioassay methods:
Topical application: The method is very useful for contact poisons. Conventional techniques
involving a Potter's tower and even the not-so-old method of Burkhard’s microapplicators, have given way to the hand held
Hamilton repeating dispenser. The technique has emerged as one of the most
convenient methods of dispensing known amount of toxins accurately on insects.
Technical grade insecticides are dissolved in acetone and a pre-calibrated 1μl solution is applied on the dorsal surface of the prothoracic
region of third instar H. armigera larvae using a 50 μl Hamilton repetitive manual dispenser.
Ideally the topical application method is
suitable to treat organisms, which have a surface area that can take at least 1μl insecticide in acetone. Results are generally erratic
with topical application bioassays on small insects such as 1st instar larvae
of many lepidopteran insects and small homopteran nymphs and adults.
Protocol for topical application:
1. Sort out the correct stage insects to treat. For
example with H. armigera, the third instar was identified as appropriate for
resistance detection. Although the most accurate method of sorting larvae is
based on the head-capsule width, this method can be very cumbersome and time
consuming. Hence, larvae in a weight range of 30 – 40
mg, which are third instars, are sorted out based on weight and assigned for
topical application treatments.
2. Place the larvae on fresh diet.
3. Open the glass vial containing insecticide
solution. Hold it firmly and aspirate 50 μl solution
into the 50 μl Hamilton syringe attached to the
microapplicator. Close the glass vial, seal it with tape and start dispensing
the toxin.
4. If synergist bioassays are to be carried out,
treat the larvae first with the synergist in acetone at the recommended dose
and then treat with insecticides 30 min later. Similarly, studies on joint
toxic action can also be conducted by applying one insecticide after another
with a 30 min spacing.
5. Gently depress the button to dispense 1 μl of the solution that forms a drop at the end of the
blunt needle. Do not squirt the insecticide. The drop of acetone is carefully
smeared on the prothoracic region of dorsal side of the third instar larva.
Ensure that the acetone does not drip to the lateral sides of the larva. Once
all the larvae in the tray are treated, close the lid and label on upper and
lower sides of the tray.
6. Transfer the cups to bioassay chambers or to BOD
incubators at 25 + 1oC, 70 + 5 %RH.
7. Change the diet once every three days.
8. Record mortality for seven days, and individual
weights of surviving larvae on the seventh day.
Diet incorporation: Diet incorporation or surface-coating tests, were developed for oral
toxicant bioassays. The tests are fairly simple, but depend on several factors
that include the availability of large amounts of toxin, thermal stability and
a consistent bioactivity under bioassay conditions.
Procedure :
1.
Insects used in diet
incorporation assays are reared to the late second or third larval instar.
2.
Serial dilutions of
formulated insecticide are added to 200 ml of diet at a ratio of 20:1 and incorporated
by vigorous shaking by hand for 30 s to produce a homogenous mixture.
3.
Diet is dispensed into
45-well bioassay trays (Tacca Plastics, Sydney, Australia) with each well
containing 1.5 ml of diet.
4.
Larvae are introduced to
trays (one larva per well) and covered.
5.
Each bioassay was performed
on three cohorts of insects. Untreated diet was used as the control.
Case study:
1. Variability in Susceptibility: Field populations
of H. armigera from eastern Australia showed significant variability in
susceptibility to cyantraniliprole.
2. Testing Methods: Baseline susceptibility was
determined through both topical and ingestion assays.
3. Intraspecific Variation: In topical bioassays,
intraspecific variation in susceptibility was 9.3-fold among 23 strains, while
in ingestion bioassays, it was 2.6-fold among 31 strains.
4. Toxicity Comparison: Cyantraniliprolewas over 400
times more toxic when administered orally compared to contact exposure.
5. Discriminating Dose:A discriminating dose of 1.5
mg/liter of diet was proposed based on diet incorporation bioassays for
resistance management of cyantraniliprolein Australia.
Immersion method:
Another form of topical application
specifically developed for simple toxicological evaluation of insecticides in
field conditions or for extension and field workers, is the larval dip method.
Larval dips for lepidopteran insects, or whole insect immersion methods for
mites, and homopteran insects, using diluted solutions of formulated
insecticides, were recommended for small sized insects. The methods appeared to
be promising for lepidopteran larvae when first proposed in the early 80s, in
terms of being rapid and practical for direct determination of resistance under
field conditions by extension workers and farmers. The methods are simple and
are somewhat closer to field application of insecticides. However, many
extraneous factors and practical problems make the assays unreliable under some
conditions. For example, it is not very easy to collect adequate sample sizes
of 100-200, healthy third instar larvae from fields in a short period of time
unless there is a very heavy infestation that goes beyond economic threshold
levels (ETL). And when pest populations are at ETL stage, farmers rarely wait
for bioassay results before making pest management decisions. Field collected
third instar H. armigera larvae are rarely healthy. Most of them harbor
diseases and parasitoids. Third instar H. armigera larvae are cannibalistic and
hurt one another when in proximity. They must be collected and kept in separate
cells. When they have to be dipped, it is important to ensure that they are not
dipped in groups. In a group they get entangled, cling together and start
biting each other. Therefore, larvae have to be dipped one at a time, placed on
blotting paper to remove excess insecticide and then placed on diet in
individual wells of multi-cell trays.
The following protocol is useful for H. armigera.
1. Collect at least 150 third instar H. armigera
larvae directly from fields and place larvae singly in individual cups.
2. Sort out 30-40 mg larvae. Discard underweight and
overweight larvae.
3. Ideally, the recommended field application rate
should set a proper guideline for the assay. For example, if endosulfan 35 EC
is recommended for field application at a concentration of 0.07 %, then the
calculations is as follows
In this example:
i.e. 1 ml endosulfan 35 EC made up to 500 ml with
water.
4. Dip the larvae one by one and place them on a
blotting paper for a few seconds. Treat at least 100 larvae with the
recommended dose. Keep 20-30 untreated larvae as controls.
5. Replace the larvae on the diet singly in
individual wells of multi-cell trays. Pre-soaked grains of Kabuli-gram (hybrid
chickpea) can be used instead of semi-synthetic diet. Change the diet daily.
6. Record observations every day for 4 days.
Case study:
1.
The experiment was carried
out at Werer Agricultural Research Center under the laboratory condition using
larva immersion and square dip methods.
2.
The selected insecticides
were tested in seven dilutions levels. In each dilution 30 larvae of 3rd
instars, H. armigera were treated.
3.
A low level of resistance
was detected for all tested locations to alphacypermethrin and a high
resistance ratio to lambda-cyhalothrin and deltamethrin for Gewane and Werer
populations.
4.
Alplhacypermethrin was the
most toxic insecticide and its LC50 was low compared to other tested synthetic
pyrethroids, Whereas, deltamethrin was the least toxic insecticide with high
LC50.
5.
The study concluded that
Helicoverpa armigera might have resistant to deltamethrin in Werer and Gewane
populations
Insecticide surface coating assay:
Commonly called residual tests, the technique
involves coating a thin film of diluted solutions of formulated insecticides on
to leaf, paper or plastic surfaces by immersion. Glass vials are coated with a
thin film of insecticide solution in acetone, by evaporating the solvent
through continuous rolling of the vials. Insects are released on to the treated
surface and thus get exposed to the insecticide. The leaf residue assays
closely simulate field exposure conditions, and have been used to monitor
insecticide resistance in H. armigera, whiteflies, aphids and mites.
Early second instar larvae are used in leaf-dip assays in Pakistan (Ahmad et
al., 1997). The method closely simulates field conditions, but tends to show
variable results because of variation in the age. of the leaf; stage of the
plant; variety; environmental stress to plants and poor leaf feeding capability
of H. armigera, in addition to the risk of avoidance of the
treated surface.
The assay protocol is described below:
1. Glass scintillation vials are used in the assay.
Rinse the vials in acetone and oven dry at 120o c.
2. Label the vials. Start dispensing the serial
dilutions of the insecticides in acetone, beginning with the lower
concentrations.
3. Pipette out 500 ul of the toxin solution into
each 25 ml glass vial. Lay the vials carefully on their sides and roll the
vials on a motorized roller or simply on a bench-top surface until the solvent
evaporates completely. Ensure uniform and complete spread of the solution over
the inner surface of the vial.
4. Coat control vials with acetone.
5. The method is ideal for moths or flies as the
test stage. Feed one-day old moths with 10 % sugar for 2-3 h and release them
at the rate of one per vial, 3 h after feeding. Close the vials with cotton or
glasswool stoppers.
6. Transfer the vials to the insectary at
temperature of 25 + 1oC and 70 + 5 % R.H or into BOD incubators.
7. Change the diet (cotton swabs with 5% sucrose+ 5%
honey in water) every days and record mortality daily for three days.
Case
Study-
Study Objective: The study
aimed to assess the resistance of Thrips tabaci Lindeman (onion thrips)
collected from commercial onion fields in Ontario to three insecticides:
lambda-cyhalothrin, deltamethrin, and diazinon.
1.
Resistance in 2001:
i.
Six out of eight adult populations were
resistant to lambda-cyhalothrin. - Resistance ratios (RR) ranged from 2 to 13.1
for lambda-cyhalothrin.
ii.
Four of these populations were also resistant
to deltamethrin, with RR ranging from 19.3 to 120.
iii.
Three out of four adult populations were
resistant to diazinon, with RR ranging from 2.5 to 165.8.
2.
Resistance in 2002:
i.
Four out of seven nymphal populations and three
out of six adult populations were resistant to deltamethrin. RR ranged from 4.3
to 72.5 for nymphal populations and from 9.4 to 839.2 for adult populations.
ii.
Limited resistance to diazinon was observed,
with only one nymphal population and one adult population showing resistance
(RR of 5.6 and 2.3, respectively).
3.
Resistance in 2003:
i.
In diagnostic dose bioassays, 15 out of 16
onion thrips populations were resistant to lambda-cyhalothrin.
ii.
All populations were resistant to deltamethrin.
iii.
Eight out of 16 populations were resistant to
diazinon.
5.
Conclusion: The results indicate widespread insecticide resistance among onion
thrips populations in commercial onion fields in Ontario. This suggests the
need for alternative pest management strategies to address this issue
effectively.
Sticky
card assay:
The sticky card test (Prabhaker, et al., 1988)
has been extensively used to monitor insecticide resistance in whiteflies. The
test is simple and elegant and has the advantage of being easily used by field
workers.
1. Yellow cards (7.5 x
12.5 cm) are sprayed with a thin layer of sticky adhesive using aerosol can.
2. Serial dilutions of
formulated insecticides are sprayed on the cards using Potter’s tower.
3. Controls are sprayed
with water.
4. Treated cards are
carried to the field in cool boxes and exposed to whiteflies for a fixed period
of time, generally one or two minutes.
5. The cards are placed
on styrofoam slabs at room temperature in humid conditions.
6. Mortality
observations are recorded 24 h later.
The test has advantages
over the vial method. Insecticide used in vial tests were found to degrade more
rapidly in vials compared to on the sticky cards. Unlike the vial tests, the
sticky card test does not impose a fumigation effect and does not provide
untreated areas for insects to seek refuge.
IRAC
Method:
|
Pest |
Irac method |
Remarks |
Pest |
IRAC method |
Remarks |
|
Leaf hopper |
5 |
Dip methods for adults or nymphs |
Aphid |
1 |
Dip method for all growth stages |
|
Weevil |
6 |
Filter paper methods for all growth stages |
19 |
Dip method for adults and nymphs |
|
|
Helicoverpa , spodoptera |
7 |
Dip method for adults and larvae |
23 |
Feeding method for nymphs |
|
|
20 |
Diets method for larvae |
24 |
Feeding method for nymph and adult |
||
|
Housefly |
26 |
Feeding method for adults |
Psyllids spp. |
2 |
Dip method for all growth stages |
|
Diamond black moth |
18 |
Dip method for larvae |
Mites |
3 |
Dip methods for egg only |
|
4 |
Dip methods for adults |
||||
|
12 |
Petri dish method for adults |
||||
|
13 |
dip method for adults |
Bioassays
with transgenic plants
Transgenic Bt-cottons currently express Cry
toxins. The toxins are effective in causing mortality to a wide range of cotton
pests. However, resistance development in target pests can impair the toxic
effects. Bioassays with transgenic plants help in
1. Evaluating efficacy
of the plants on target pests.
2. Determining the
expression levels of the Cry toxins.
3. Confirming
resistance when it occurs in target pests
Bioefficacy
of Bt-transgenic plant parts
1. The Bt-cotton plant
parts to be tested are excised along with their petiole from the node, and
brought to the lab in cool boxes. The plant parts are rinsed under tap water
and sandwiched gently in two layers of blotting paper to remove the water.
2. The bioassay is
carried out in cups having a 3 mm diameter hole at the bottom, through which
the petiole is passed and the distal end dipped in 0.5 % agar, containing
anti-mould solution and Murashige-Skoog medium (optional), present in a cup
held beneath the upper cup. The hole is plugged with wax around the petiole.
Alternatively, the plant parts are placed in plastic cups directly or on a
moist layer of blotting paper. However, in this case, the parts will have to be
changed everyday, to avoid larval mortality or growth reduction due to tissue
deterioration.
3. Ensure that the same
method of bioassay is followed to generate standard curves with plant parts
from non-Bt plants and to test the efficacy of the plant parts from Bt-cotton.
4. Release five, first
instar larvae on the plant part in each cup. Close the cups with finely
perforated lids and transfer the cups to bioassay chambers or to BOD incubators
at 25 + 1oC, 70 + 5 % R. H. Change the plant parts every alternate day.
5. If the larvae moult
to the second instar, transfer each larva into a single cup, to avoid
cannibalism.
6. Record mortality for
seven days, and individual weights of surviving larvae on the seventh day.
Whole-plant
efficacy assessment
Bt-transgenic plants can be tested for their
efficacy in no-choice bioassays by confining larvae with plant parts.
1.
The simplest of these tests is to release 10 first instar larvae on each branch
(sympodia of cotton plants) and cover them with two layers of fine-perforated
plastic bags.
2.
The bags are sealed at the base of the branch tightly with rubber bands and
tape. The bags must be transparent and allow air, but not permit larval escape.
Two layers of the bags normally prevent escapes.
3.
The method can be used for potted and field grown plants.
4.
Observations for the presence or escape of larvae must be made everyday.
5.
Control plants comprise isogenic non-Bt plants on which larva are released and
confined in perforated bags, identical to that on the Bt-plants.
6.
Final observations are recorded on the seventh day.
7.
The reduction in weight of larvae surviving on Bt-plants can be calculated
relative to the average weight of larvae on control plants.
8.
Mortality observations can be used to calculate the % mortality on transgenic
plants, in comparison with that on non-Bt plants.
Case
study :
v Approval
and Cultivation: Bt maize producing the Cry1Ab toxin was approved for
cultivation in the EU in 1998 to manage Sesamia nonagrioides and Ostrinia
nubilalis.
v Limited
Planting: Spain is the sole country in the EU where Bt maize has been planted
consistently since approval. In 2021, approximately 100,000 hectares of
Cry1Ab-producing Bt maize were cultivated in the EU, with Spain accounting for
96% of the area and Portugal 4%.
❖ Insect Resistance Management:
The EU has implemented insect resistance management strategies based on the
high-dose/refuge strategy since 1998
❖ Monitoring for Resistance:
Regular monitoring through laboratory bioassays has been conducted to detect
any decrease in susceptibility to Cry1Ab. As of 2021, no reduction in
susceptibility to Cry1Ab has been observed in either targeted pest species.
❖ Effectiveness Confirmation:
No instances of control failures have been reported, indicating the continued
efficacy of Cry1Ab-producing Bt maize against Sesamia nonagrioides and Ostrinia
nubilalis.
❖ Future Challenges: EU
regulations limiting the approval of new genetically modified crops,
particularly those producing multiple Bt toxins targeting corn borers, may pose
challenges to future resistance management strategies.
The
study focuses on insecticide resistance in five major insect pests of cotton in
India: Helicoverpa armigera, Pectinophora gossypiella, Spodoptera litura,
Earias vittella, and Bemisia tabaci.
Cypermethrin
Resistance: Helicoverpa armigera showed widespread resistance to cypermethrin,
with resistance levels ranging from 23 to 8022-fold in field strains.
Endosulfan
and Chlorpyriphos Resistance: Resistance to endosulfan and chlorpyriphos in H.
armigera was low to moderate.
Pectinophora
gossypiella: Overall resistance to pyrethroids was low, but high resistance
levels (23-57-fold) to endosulfan were recorded in some areas of Central India.
Spodoptera
litura: Majority of strains from South India exhibited high resistance levels
(61-148-fold) to cypermethrin and chlorpyriphos.
Earias
vittella: Resistance was low to moderate in strains from North India.
Bemisia
tabaci: Exhibited moderately high levels of resistance to cypermethrin, while
resistance to endosulfan and chlorpyriphos was negligible in tested field
strains.
Case
study:
Ø Between
2002 and 2009, Bt-cotton effectively controlled pink bollworm (PBW) in India by
producing Cry1Ac and Cry2Ab toxins.
Ø However,
since 2009, studies have reported increased PBW survival on Bt-cotton,
indicating reduced susceptibility to Cry toxins.
Ø A
recent study in Andhra Pradesh aimed to estimate the frequency of resistance
alleles to Cry1Ac and Cry2Ab toxins in PBW populations using F2 screen
methodology.
Ø The
study found a Cry1Ac resistance allele frequency of 0.082 and a Cry2Ab
resistance allele frequency of 0.054, with a high detection probability.
Ø It
revealed that high PBW survival and damage on Bt-cotton expressing Cry1Ac and Cry2Ab, with over 30% flower
damage, over 90% green boll damage, and over 80% locule damage.
Statistical
analysis of bioassay data:
Log dose probit analysis is carried out to
obtain a regression equation that enables the calculation of the dose /
concentration required for any particular % mortality that they cause in the
test population. The analysis can also be done for biological responses other
than mortality, such as weight reduction, moult inhibition etc. For the
regression analysis, it is necessary to assess the biological response of the
organism against a series of serially diluted concentrations. Once the bioassay
results are found to confirm to a graded response depending on the
concentration of the toxicant, they are then subjected to probit analysis
through a series of manual calculations or on computer-aided programs such as
POLO, MLP, MSTAT, GENSTAT etc. The details of probit analysis are not being
dealt with here. Generally the median lethal dose (commonly called the LD50, a
dose which kills 50% of the test population) is calculated to compare responses
of test populations. If control mortality exceeds 5% discard the replicate.
1.
Use Abbott's formula to correct control mortality
2.
Plot percentage mortality on a probit scale against log insecticide dose. Read
the LD50 and LD90 values from the graph. Alternatively software programs such
as POLO-PC, MLP, MSTAT, GLIM or GENSTAT may be used for probit analysis.
Resistance
Detection Kit:
Two immunochromatographic dip-stick-format kits
were developed to detect resistance to carbamates (methomyl) and
organophosphates (quinalphos, chlorpyriphos and profenophos). The strips are
based on polyclonal antisera raised against resistance associated esterase
isozymes isolated form H. armigera. The use of 20-40 strips would be adequate
to determine the resistance frequencies in a region within a radius of about 20
km. The strips are expected to be modestly priced (equals the manufacturing
cost). The strips are simple to use and were specifically designed for use of
illiterate farmers. Each of the immunochromatographic strip is a 6x 0.4 cm
strip that contains an assembly of nitrocellulose membrane on a plastic
backing, overlaid by small filter pads and conjugate release pads, that enable
the uptake of the test insecticide by capillary flow so that the nitrocellulose
strip gets saturated.The test takes 10 minutes for the results to appear.
The
basic steps in the test procedure are outlined as below.
Step
1. Place a one cm sized larva in a plastic vial.
Step
2. Pour 0.5 ml buffer (provided with the kit)
Step
3. Crush the larva in buffer with a pestle.
Step
4. Place the dip-stick into the homogenate as per the instructions provided.
Step
5. Wait for 10 minutes until the strip is saturated with the capillary flow of
the solution.
Step
6. Two clear purple band (as shown in figure) represent a resistant larva. Only
one purple band at the upper portion indicates susceptible larva.
Step
7. Calculate results from 20-40 strips to determine resistance frequencies in
the region.
Applications
of resistance detection and monitoring are as follows:
1. Help to document
geographical and temporal variability in population responses to insecticide
selection pressures.
2. Helps to keep track
of the precise changes in resistant phenotype frequencies occurring in field
populations.
3. Resistance detection
bioassays determine the relative efficacy of insecticides for a given field
population
4. The bioassays
diagnose and confirm the causes of pest control failure by specific
insecticides under field conditions.
5. helps to evaluate
the impact of resistance management strategies, which have been implemented.
Conclusion:
Monitoring results can
provide an early warning of resistance evolution, advance the understanding of
factors that drive resistance evolution, document the effectiveness of IRM
strategies, and provide relevant information to guide implementation of
effective pest management practices.
Reference:-
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