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Applied and Environmental Microbiology, April 2000, p. 1553-1558, Vol. 66, No. 4
0099-2240/00/$04.00+0
Copyright © 2000, American Society for Microbiology. All rights reserved.
A Binding Site for Bacillus
thuringiensis Cry1Ab Toxin Is Lost during Larval Development in
Two Forest Pests
Carolina
Rausell,
Amparo
Consuelo
Martínez-Ramírez,
Inmaculada
García-Robles, and
María Dolores
Real*
Departamento de Genética, Facultad de
Ciencias Biológicas, 46100-Burjassot (Valencia), Spain
Received 28 June 1999/Accepted 27 January 2000
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ABSTRACT |
The insecticidal activity and receptor binding properties of
Bacillus thuringiensis Cry1A toxins towards the forest
pests Thaumetopoea pityocampa (processionary moth) and
Lymantria monacha (nun moth) were investigated. Cry1Aa,
Cry1Ab, and Cry1Ac were highly toxic (corresponding 50% lethal
concentration values: 956, 895, and 379 pg/µl, respectively) to
first-instar T. pityocampa larvae. During larval
development, Cry1Ab and Cry1Ac toxicity decreased with increasing age,
although the loss of activity was more pronounced for Cry1Ab. Binding
assays with 125I-labelled Cry1Ab and brush border membrane
vesicles from T. pityocampa first- and last-instar larvae
detected a remarkable decrease in the overall Cry1Ab binding affinity
in last-instar larvae, although saturable Cry1Ab binding to both
instars was observed. Homologous competition experiments demonstrated
the loss of one of the two Cry1Ab high-affinity binding sites detected
in first-instar larvae. Growth inhibition assays with sublethal doses
of Cry1Aa, Cry1Ab, and Cry1Ac in L. monacha showed that all
three toxins were able to delay molting from second instar to third
instar. Specific saturable binding of Cry1Ab was detected only in
first- and second-instar larvae. Cry1Ab binding was not detected in
last-instar larvae, although specific binding of Cry1Aa and Cry1Ac was
observed. These results demonstrate a loss of Cry1Ab binding sites
during development on the midgut epithelium of T. pityocampa and L. monacha, correlating in T. pityocampa with a decrease in Cry1Ab toxicity with increasing age.
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INTRODUCTION |
The discovery of Bacillus
thuringiensis has had the greatest impact on the use of
biopesticides in forestry, as well as in crop systems and stored
products. B. thuringiensis is a bacterium that produces
proteinaceous insecticidal toxins in the form of inclusion bodies or
crystals at sporulation. Each toxin specifically recognizes and binds
to receptors in the insect gut, forming membrane pores that disrupt the
selective permeability of the cells and eventually cause lysis of
epithelial cells and insect death (5, 11, 22, 36).
Binding of toxins to midgut receptors is a key step in the mode of
action of B. thuringiensis toxins (5, 6, 14, 15, 36,
37). Specific binding of lepidopteran-active toxins has been
demonstrated by incubating midgut brush border membrane vesicles (BBMVs) (14, 15, 36, 37), tissue sections (5, 6), and BBMV protein blots (10, 26) with radiolabelled or
biotin-labelled toxins. These studies aided in the characterization of
the receptor system of many insect pests and identified several
toxin-binding proteins (7, 12, 20, 21, 23, 30, 32).
The use of B. thuringiensis as a biopesticide has proven to
be a viable alternative to chemical insecticides (25). There has been a major increase in the proportion of areas treated with B. thuringiensis in North America, particularly in Eastern
Canada (33, 34), and in Eastern Europe, at the expense of
chemical insecticides such as fenitrothion (Canada) and diflubenzuron
and pyrethroids (Eastern Europe). This trend is expected to continue in
the future, with at least 50% of forest areas being treated with
microbial insecticides, of which B. thuringiensis will
remain dominant (8). The environmental benefit of their
increased use is their specificity and the possibility to use them in
both rural and urban woods (25).
This paper analyzes the toxicity and binding of B. thuringiensis toxins in two important forest pests,
Thaumetopoea pityocampa and Lymantria monacha, in
an attempt to optimize the use of this microbial pathogen to control
the two forest pests in the field.
T. pityocampa, processionary moth, is an important pine
defoliator that represents a major endemic pest in Southern Europe. The
larvae are covered with urticating hairs that can cause serious skin
irritation in those living in close proximity to infested trees, and
there is, thus, considerable public pressure to manage this moth population.
The nun moth, L. monacha, is one of the most severe insect
pest species of European conifers, although this insect can also affect
other host genera, such as Quercus, Fagus, and
Betula, and even herbs. This is an epidemic pest distributed
throughout Europe and parts of Asia, south of 60° latitude, thus
occurring from Portugal to Japan.
We have previously demonstrated that Cry1Aa, Cry1Ab, and Cry1Ac
specifically bound to the midgut brush border membrane of second-instar
T. pityocampa larvae and that all three toxins compete for
binding (28). In the present paper, we correlate the
decrease in Cry1Ab toxicity during development with the loss of a
binding site for this toxin in last-instar larvae. Similarly, in
L. monacha we have found that binding of Cry1Ab to the
larval midgut membrane was lost during the larval development of the insect.
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MATERIALS AND METHODS |
Biological material.
T. pityocampa S. and L. monacha L. larvae were used in all experiments. T. pityocampa larvae were collected from natural populations in
Burjassot (Valencia, Spain). Larvae were fed on pine needles (Pinus sylvestris) at room temperature. L. monacha larvae were reared from egg masses collected from Orihuela
del Tremedal (Teruel, Spain). Eggs were stored at 4°C until needed
and then incubated at 25°C for hatching. At eclosion, larvae were
transferred to petri dishes and reared, until third instar, on an
artificial diet, according to the method of Grijpma et al.
(13), except that aureomycin was omitted due to
incompatibility with B. thuringiensis treatments
(27). From third instar until pupation, larvae were fed on
Pinus halepensis needles. Rearing conditions were as
follows: 25°C, 70% relative humidity, and a 16-h-8-h (light-dark) photoperiod.
Insect toxicity assays.
Bioassays were carried out to
determine the sensitivity of first-instar T. pityocampa
larvae to Cry1A-type B. thuringiensis trypsin-activated
toxins. Trypsin digestion and toxin purification were performed by the
method of Höfte et al. (16). Each bioassay consisted
of five doses of the corresponding activated crystal toxins (Cry1Aa,
Cry1Ab, and Cry1Ac) prepared in phosphate-buffered saline (PBS) (8 mM
Na2HPO4, 2 mM KH2PO4,
150 mM NaCl, pH 7.4) containing 0.1% bovine serum albumin (BSA) and a
control of PBS (pH 7.4)-0.1% BSA. Fresh P. sylvestris
needles were dipped into this suspension and allowed to air dry. Three
groups of 20 larvae (1 day old) were placed on the coated pine needles
for 4 days, after which the coated needles were replaced with fresh,
untreated needles. Mortality was recorded 4 days later. Toxicity data
were evaluated by probit analysis using the Polo PC program (LeOra Software, Berkeley, Calif.). Fifty percent lethal concentration (LC50) measurements refer to the concentration of crystal
protein which, when applied uniformly on the needles, produces 50% mortality.
Toxicities of Cry1Ab (1 and 10 ng/µl) and of Cry1Ac (0.4 and 4 ng/µl) were also tested on third-instar T. pityocampa
larvae as described above.
To study sublethal effects of
B. thuringiensis Cry1A-type
toxins on
L. monacha development, freshly prepared
artificial diet
was dispensed in wells with 2 cm
2 of
surface area (Costar 24-well cluster plate), and dilutions
of the
corresponding toxin (50 µl; prepared in PBS [pH 7.4]-0.1%
BSA)
were uniformly applied over the food surface in each well
and allowed
to dry. Single larvae in the first day of the second
instar were placed
in each well; 20 larvae were used per dilution.
Three dilutions of each
trypsin-activated toxin (Cry1Aa, Cry1Ab,
and Cry1Ac), corresponding to
2, 20, and 200 ng/cm
2, were analyzed. After five days of
toxin exposure, the percentage
of larvae reaching third instar was
scored. Experiments were duplicated,
and controls with PBS were also
included.
Binding to midgut tissue sections.
The midgut tissue
preparation, sectioning, binding, and immunocytochemical staining were
performed according to the method of Bravo et al. (5).
Experiments involving binding of Cry1Ab and Cry1Ac trypsin-activated
toxins to T. pityocampa tissue sections from first- and
third-instar larvae were carried out as follows. Tissue sections were
incubated with the appropriate toxin for 1 h by using 0.3 ml of a
mixture of 10 µg of toxin per ml of TST buffer (10 mM Tris-HCl, 100 mM NaCl, 1 mM sodium ethylmercurisalicylate, 0.1% [vol/vol] Triton
X-100, pH 7.6). Immunolocalization of toxins was performed by
incubating tissue sections overnight with 0.3 ml of the monoclonal
antibody 4D6 (Plant Genetic Systems, Ghent, Belgium) (0.1 µg/ml in
TST buffer). Unbound antibody was rinsed with TST buffer, and an
alkaline phosphatase-coupled secondary antibody was used to detect the
binding. Color development was obtained by incubation in 0.5 ml of
BCIP-nitroblue tetrazolium (NBT) solution (8 mM
5-bromo-4-chloro-3-indolyl phosphate and 9 mM 4-NBT chloride in 5 mM
MgCl2, 100 mM Tris, 100 mM NaCl, pH 9.5) for 10 min. Color
development was stopped by lowering the pH with glacial acetic
acid-water (1:10, vol/vol). Slices were dehydrated by successive
incubations in 70% and 100% ethanol and 100% xylol. Finally, tissue
sections were covered with Entellan mounting medium (Merck, Darmstadt,
Germany) and photographed.
Preparation of BBMVs.
BBMVs were prepared according to the
method of Wolfersberger et al. (40) from T. pityocampa first- and early second-instar larvae, T. pityocampa last-instar larvae, L. monacha first- and early second-instar larvae, and L. monacha last-instar
larvae. All BBMV preparations were from whole larvae of the
corresponding instar, except for L. monacha last-instar
larvae, from which dissected midguts were used. The final pellet was
resuspended in ice-cold MET buffer (0.3 M mannitol, 5 mM EGTA, 17 mM
Tris-HCl, pH 7.5), immediately frozen, and stored at 
80°C.
The protein concentration of BBMVs was measured by Bradford's
procedure (
4) with a Bio-Rad kit (Richmond, Calif.), with
BSA as the
standard.
Labelling of toxins.
Cry1Aa, Cry1Ab, and Cry1Ac
trypsin-activated toxins were iodinated by the chloramine-T method
(17) as described by Van Rie et al. (36). The
specific radioactivities of iodinated toxins, determined by a sandwich
enzyme-linked immunosorbent assay technique, were 5 mCi/mg for Cry1Aa,
36 mCi/mg for Cry1Ab, and 4 mCi/mg for Cry1Ac.
Binding assays.
Immediately before the binding assay, the
buffer of the BBMV suspension was replaced with PBS, pH 7.4, containing
0.1% BSA, by microcentrifuge centrifugation. The reaction volume was
0.1 ml, and all samples were duplicated. Other optimal assay conditions were as follows: 0.7 nM 125I-Cry1Ab, 4 µg of BBMV, and
120 min of incubation, for T. pityocampa first- and early
second-instar larvae; 1.9 nM 125I-Cry1Ab, 12 µg of BBMV,
and 60 min of incubation, for T. pityocampa last-instar
larvae; 0.8 nM 125I-Cry1Ab and 60 min of incubation for
L. monacha first- and early second-instar larvae; and
125I-labelled toxin (8.7 nM Cry1Aa or 1.3 nM Cry1Ac) and
either 60 min (Cry1Aa experiments) or 120 min (Cry1Ac experiments) of
incubation for L. monacha last-instar larvae. After
incubation, the reaction mixtures were filtered through Whatman GF/F
glass-fiber filters in a Millipore filtration manifold 1225 Unit
(Millipore Corp., Bedford, Mass.). Filters were washed with 5 ml of
ice-cold PBS, pH 7.4, containing 0.1% (wt/vol) BSA, and their
radioactivity was measured in a 1282 Compugamma CS counter (LKB).
For homologous competition experiments, the reaction mixture contained
the corresponding amount of BBMV and labelled toxin
and increasing
amounts of unlabelled toxin. Binding data were
analyzed with the LIGAND
program (
24), which estimates the binding
constants
(equilibrium dissociation constant [
Kd] and
binding
site concentration [
Rt]). A Student
t test was used to determine
whether the mean values of the
calculated binding constants were
significantly
different.
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RESULTS |
T. pityocampa toxicity assays.
Toxicity values for
Cry1Aa, Cry1Ab, and Cry1Ac toxins to first-instar T. pityocampa larvae are presented in Table
1. Cry1Ac was most toxic, while Cry1Aa
and Cry1Ab were equally toxic (95% fiducial limits overlap).
During processionary moth larval development, 1,000 pg of Cry1Ab per
µl (concentration around the Cry1Ab LC
50 value obtained
in first-instar larvae) produced only 10% mortality when applied
to
either second- or third-instar larvae. However, 400 pg of Cry1Ac
per
µl (also corresponding to the Cry1Ac LC
50 value obtained
in
first-instar larvae) caused 35% mortality in second-instar larvae
and 10% mortality in third-instar larvae. Hence, the susceptibility
of
second-instar larvae to Cry1Ab and Cry1Ac toxins is markedly
different.
A concentration of 10,000 pg of Cry1Ab per µl (a toxin
concentration
10-fold higher than the respective LC
50 values in
first-instar larvae, which produced 100% lethality in first-instar
larvae) resulted in 10% mortality in second-instar larvae. Even
a
4,000-pg/µl Cry1Ac concentration (corresponding to a 100% lethal
concentration in first-instar larvae) caused between 40 and 60%
mortality in second-instar larvae. These findings stress that
second-instar larvae display a different susceptibility to Cry1Ab
and
Cry1Ac, Cry1Ab being less toxic than
Cry1Ac.
Binding to T. pityocampa midgut brush border
membrane.
To analyze the cellular basis for the different toxin
susceptibilities of different stages of T. pityocampa larval
development, tissue sections from early first- and third-instar
T. pityocampa larvae were prepared and binding of Cry1Ab and
Cry1Ac toxins was analyzed. Cry1Ab and Cry1Ac both were able to
specifically bind to the midgut brush border membrane of first- and
third-instar larvae (Fig. 1), confirming
the existence of specific binding sites for these B. thuringiensis toxins (28). Controls without toxin or
primary antibody were also included (data not shown).
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FIG. 1.
Binding of Cry1Ab and Cry1Ac to brush border membranes
of first- and third-instar T. pityocampa larva midguts. (A)
Cry1Ab binding to first-instar larva tissue sections; (B) Cry1Ac
binding to first-instar larva tissue sections; (C) Cry1Ab binding to
third-instar larva tissue sections; and (D) Cry1Ac binding to
third-instar larva tissue sections. BM, basal membrane; MV, microvilli;
L, lumen.
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To investigate the biochemical basis of the observed developmental
variations in Cry1Ab toxicity, binding saturation experiments
with
125I-labelled Cry1Ab were performed. Cry1Ab binding to
BBMVs from
first- and last-instar
T. pityocampa larvae
followed sigmoidal
kinetics (Fig.
2).
Binding of Cry1Ab to BBMVs varied with larval
development, maximum
binding being shown to be between 80 and
150 µg of BBMV/ml, depending
on the developmental stage analyzed
(Fig.
2). The sigmoidal shape of
the curve observed with last-instar
larvae (Fig.
2B) indicated a
decrease in binding affinity when
compared with the hyperbolic curve
obtained in first-instar larvae
(Fig.
2A).
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FIG. 2.
Specific binding of 125I-labelled Cry1Ab
toxin as a function of BBMV concentration of first-instar T. pityocampa larvae (A) or last-instar T. pityocampa
larvae (B). For each point, nonspecific binding (percentage of the
total 125I-Cry1Ab counts that were bound to the BBMVs in
the presence of a 500-fold excess of unlabelled Cry1Ab) was subtracted
from total binding (percentage of the total 125I-Cry1Ab
counts that were bound to the BBMVs).
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Quantitative estimates of homologous binding competition experiments
are shown in Fig.
3. Equilibrium
dissociation constants
and binding site concentrations for Cry1Ab
binding to first- and
last-instar
T. pityocampa larvae were
as follows. For first- and
second-instar larvae, the
Kd1 (± standard deviation [SD]) was
0.29 ± 0.17 nM; the
Kd2 was 2.55 ± 1.62 nM.
The
Rt1 was 0.95 ±
0.41 pmol/mg; the
Rt2 was 2.75 ± 1.74 pmol/mg. For
last-instar
larvae, the
Kd (± SD) was 3.42 ± 0.66 nM, and the
Rt was 1.29
± 0.06 pmol/mg. In first-instar larvae, Cry1Ab exhibits one additional
high-affinity site which is absent from last-instar larvae. The
loss of
the Cry1Ab high-affinity binding site during development
is in
accordance with the decrease in affinity observed in the
saturation
curve obtained in last-instar larvae (Fig.
2B) compared
with that of
first-instar larvae (Fig.
2A). In first-instar larvae,
saturation is
reached at around 80 µg of BBMV protein/ml, whereas,
in last-instar
larvae, only 25% of maximum binding was detected
for that BBMV protein
concentration.
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FIG. 3.
Binding of 125I-labelled Cry1Ab toxin as a
function of increasing concentration of nonlabelled Cry1Ab to BBMVs of
first-instar T. pityocampa larvae (A) or last-instar
T. pityocampa larvae (B). For each panel, data points
correspond to one of at least three independent competition experiments
performed.
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The lack of Cry1Ab binding to the high-affinity site in last-instar
larvae is consistent with the decrease in toxicity and
binding to
tissue sections of this toxin during processionary
moth
development.
Larval growth inhibition assays on L. monacha.
Preliminary bioassays with L. monacha showed a low toxicity
of Cry1A toxins compared with that for other insects, such as T. pityocampa. Due to this fact, we performed larval growth
inhibition experiments with Cry1Aa, Cry1Ab, and Cry1Ac toxins instead
of dose-mortality curves. The effects of three sublethal concentrations of each toxin are shown in Fig. 4. In
these experiments, as well as in controls with nontreated larvae,
mortality never exceeded 5% and 87% of nontreated larva controls
survived until the third instar. Results showed around 100% inhibition
of molting from second to third instar when a 200-ng/cm2
concentration of Cry1A-type toxins was used. Fifty percent of the
larvae did not molt when exposed to a 2-ng/cm2
concentration of Cry1Aa, while no significant differences with controls
were found when the same Cry1Ab or Cry1Ac concentrations were applied.
A 10-fold-higher concentration of Cry1Ac (20 ng/cm2) was
needed to give rise to 50% molting inhibition, while for the same
Cry1Ab concentration no significant differences with controls were
observed.
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FIG. 4.
Percentage of L. monacha second-instar larvae
that molted to the third instar when sublethal concentrations of
Cry1Aa, Cry1Ab, and Cry1Ac were applied on an artificial diet.
Percentages were calculated over the total surviving larvae after
treatment. In all experiments (including the control, nontreated
larvae), mortality never exceeded 5%. The control, nontreated larvae,
showed 87% molting to the third instar. Symbols:  , Cry1Aa;  ,
Cry1Ab; and  , Cry1Ac.
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In conclusion, larval growth was significantly affected by toxin
concentration and the order of relative efficiency in molting
inhibition was as follows: Cry1Aa > Cry1Ac >
Cry1Ab.
Binding to L. monacha midgut BBMVs.
In order to
determine whether the observed L. monacha larval growth
inhibition can be related to the occurrence of specific binding of
toxins to BBMV, binding experiments with radiolabelled toxins were
carried out. Saturable binding of 125I-Cry1Aa and
125I-Cry1Ac to BBMVs prepared from last-instar L. monacha larvae was evident (Fig. 5A
and B). However, no binding of 125I-Cry1Ab was detected at
this developmental stage (Fig. 5C), which contrasted with the observed
effect of Cry1Ab on molting inhibition. BBMVs from first- and
second-instar larvae were then prepared, and binding saturation
experiments with 125I-Cry1Ab were performed. As in the case
of T. pityocampa with Cry1Ab, results showed saturable
binding of Cry1Ab to BBMVs of early stages of L. monacha
(Fig. 5D).
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FIG. 5.
(A) Specific binding of 125I-Cry1Aa to
increasing concentrations of BBMVs from last-instar L. monacha larvae. (B) Specific binding of 125I-Cry1Ac to
increasing concentrations of BBMVs from last-instar L. monacha larvae. (C) Specific binding of 125I-Cry1Ab to
increasing concentrations of BBMVs from last-instar L. monacha larvae. (D) Specific binding of 125I-Cry1Ab to
increasing concentrations of BBMVs from first- and second-instar
L. monacha larvae. In all cases, the reaction volume was 0.1 ml and for each point nonspecific binding was subtracted from total
binding (see the legend to Fig. 2).
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DISCUSSION |
In this paper, the insecticidal activity of Cry1A-type B. thuringiensis toxins on T. pityocampa, and thus their
suitability for processionary moth control, was demonstrated.
In addition, we demonstrated that the toxicity of Cry1Ab and Cry1Ac
during larval development decreased with increasing age. A loss of
toxicity in later instars is well documented and has been found for
several insect pests in relation to B. thuringiensis (1-3, 9, 18, 19, 22, 29, 31, 35, 38), showing that the
toxicity was mainly determined by the developmental stage of the
larvae, i.e., it was significantly higher for earlier stages when
compared with later stages. The biochemical basis of the reduced
susceptibility among instars has so far been investigated only in
Spodoptera spp. (19, 22), in which a variation of toxin receptor density during larval development has been proposed to
account for the decrease in the capacity of Cry1C and Cry1D toxins to
induce permeability changes on BBMVs in late instars (22).
In addition, experimental data indicated a loss or affinity reduction
of one of the two Cry1Ab binding sites during development, which may
account for the reduced Cry1Ab susceptibility with increasing T. pityocampa larval age. To our knowledge, T. pityocampa
is the first insect pest in which changes in B. thuringiensis toxin binding sites during development have been
reported. The occurrence of these changes may have a significant
influence on the final efficacy of the formulations used against this
insect, because larvae in the field are not synchronized and different
instars coexist in infested areas. Thus, according to our results,
formulations having a higher content of toxins whose activity is not
readily lost during development would be more appropriate than those
enriched in other toxins whose toxic effect decreases during larval
development, as in the case of Cry1Ab in this insect.
On the other hand, in L. monacha, growth inhibition assays
with sublethal doses of Cry1Aa, Cry1Ab, and Cry1Ac have shown that all
three toxins were able to delay development from second to third
instar. We have also demonstrated that Cry1Aa and Cry1Ac toxins
specifically bound to last-instar L. monacha larva BBMVs whereas Cry1Ab did not, although Cry1Ab saturable binding to the brush
border membrane of first-second instar L. monacha larvae was
indeed observed. Therefore, the decrease in Cry1Ab toxicity during
development due to the loss of binding sites in later instars could be
a more general phenomenon, present not only in T. pityocampa but also in other insects.
A similar rationale could be applied to Lymantria dispar. In
this insect, an inverse correlation has been reported between toxicity
and binding properties of Cry1Ab and Cry1Ac to BBMV (39), Cry1Ab being significantly more toxic than Cry1Ac albeit binding with
lower affinity. Surprisingly, when we reexamined the experiments of
Wolfersberger (39) we realized that toxicity assays were performed on first-instar larvae whereas homologous competition experiments were done with BBMVs from last-instar larvae. Again, a
decrease in or loss of Cry1Ab affinity for its binding sites during
development could explain this disagreement between toxicity and
binding data.
The finding of the variation of Cry1Ab binding sites during development
can be explained by a differential expression pattern of the binding
molecule in various larval stages. Alternatively, specific
posttranslation modifications during development could also be
responsible for the decrease in or loss of Cry1Ab binding. The overall
effect of either of these changes could have important implications in
the characterization of the toxin binding molecules as well as in the
design and application of B. thuringiensis formulations appropriate for each insect pest.
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ACKNOWLEDGMENTS |
We are indebted to Gema Pérez-Guerra and Axel Gruppe for
their technical support on the rearing of L. monacha. We
also thank Plant Genetic Systems (Ghent, Belgium), Eduardo Obama,
Laboratorio de Sanidad Vegetal de Mora de Rubielos (Teruel), and
Servicio de Protección Vegetal de Silla (Valencia) for their
expert technical assistance.
This work was supported by the Spanish Ministerio de Agricultura Pesca
y Alimentación (Project No. AGF93-1171) and the Spanish Ministerio de Educación y Ciencia (Project No. PB95-1090).
Carolina Rausell and Inmaculada García-Robles were supported by
grants from the Spanish Ministerio de Educación y Ciencia and the
Conselleria Valenciana d'Educació i Ciència, respectively.
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FOOTNOTES |
*
Corresponding author. Mailing address: Departamento de
Genética (Facultad de Ciencias Biológicas), Doctor Moliner,
50, 46100-Burjassot (Valencia), Spain. Phone: 34 96 398 30 28. Fax: 34 96 398 30 29. E-mail: realmd{at}uv.es.
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REFERENCES |
| 1.
|
Ali, A., and S. Y. Young.
1996.
Activity of Bacillus thuringiensis Berliner against different ages and stages of Helicoperva zea (Lepidoptera: Noctuidae) on cotton.
J. Econ. Entomol.
31:1-8.
|
| 2.
|
Bauer, L. S.
1990.
Response of the cottonwood leaf beetle (Coleoptera: Chrysomelidae) to Bacillus thuringiensis var. san diego.
Environ. Entomol.
19:428-431.
|
| 3.
|
Bauer, L. S.
1992.
Response of the imported willow leaf beetle to Bacillus thuringiensis var. san diego on poplar and willow.
J. Invertebr. Pathol.
59:330-331[CrossRef].
|
| 4.
|
Bradford, M. M.
1976.
A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem.
72:248-254[CrossRef][Medline].
|
| 5.
|
Bravo, A.,
K. Hendrickx,
S. Jansens, and M. Peferoen.
1992.
Immunocytochemical analysis of specific binding of Bacillus thuringiensis insecticidal crystal proteins to lepidopteran and coleopteran midgut membranes.
J. Invertebr. Pathol.
60:247-253[CrossRef].
|
| 6.
|
Denolf, P.,
S. Jansens,
J. Van Rie,
D. Degheele, and M. Peferoen.
1993.
Biotinylation of Bacillus thuringiensis insecticidal crystal proteins.
Appl. Environ. Microbiol.
59:1821-1827[Abstract/Free Full Text].
|
| 7.
|
Denolf, P.,
K. Hendrickx,
J. Seurinck,
J. Vandamme,
S. Jansens,
M. Peferoen, and J. Van Rie.
1997.
Cloning and characterization of Manduca sexta and Plutella xylostella midgut aminopeptidase N related to Bacillus thuringiensis toxin-binding proteins.
Eur. J. Biochem.
248:748-761[Medline].
|
| 8.
|
Evans, H. F.
1997.
The role of microbial insecticides in forest pest management, p. 29-40.
In
British Crop Protection Council (ed.), Microbial insecticides: novelty or necessity? Symposium proceedings, no. 68. Major Desing & Production Ltd., Nottingham, United Kingdom.
|
| 9.
|
Ferro, D. N., and S. M. Lyon.
1991.
Colorado potato beetle (Coleoptera: Chrysomelidae) larval mortality: operative effects of Bacillus thuringiensis subsp. san diego.
J. Econ. Entomol.
84:806-809.
|
| 10.
|
Garczynski, S. F.,
S. W. Crim, and M. J. Adang.
1991.
Identification of putative insect brush border membrane-binding molecules specific to Bacillus thuringiensis  -endotoxin by protein blot analysis.
Appl. Environ. Microbiol.
57:2816-2820[Abstract/Free Full Text].
|
| 11.
|
Gill, S. S.,
E. A. Cowles, and P. V. Pietrantonio.
1992.
The mode of action of Bacillus thuringiensis endotoxins.
Annu. Rev. Entomol.
37:615-636[CrossRef][Medline].
|
| 12.
|
Gill, S.,
E. Cowles, and V. Francis.
1995.
Identification, isolation and cloning of a Bacillus thuringiensis CryIA(c) toxin-binding protein from the midgut of the lepidopteran insect Heliothis virescens.
J. Biol. Chem.
270:20309-20315[Abstract/Free Full Text].
|
| 13.
|
Grijpma, P.,
J. J. M. Belde, and D. C. van der Werf.
1987.
Artificial diets and rearing of the nun moth, Lymantria monacha.
Entomol. Exp. Appl.
45:219-225[CrossRef].
|
| 14.
|
Hofmann, C.,
P. Lüthy,
R. Hütter, and V. Pliska.
1988.
Binding of the delta-endotoxin from Bacillus thuringiensis to brush-border membrane vesicles of the cabbage butterfly (Pieris brassicae).
Eur. J. Biochem.
173:85-91[Medline].
|
| 15.
|
Hofmann, C.,
H. Vanderbruggen,
H. Höfte,
J. Van Rie,
S. Jansen, and H. Van Mellaert.
1988.
Specificity of Bacillus thuringiensis -endotoxins is correlated with the presence of high affinity binding sites in the brush border membrane of target insect midguts.
Proc. Natl. Acad. Sci. USA
85:7844-7848[Abstract/Free Full Text].
|
| 16.
|
Höfte, H.,
H. De Greve,
J. Seurinck,
S. Jansens,
J. Mahillon,
C. Ampe,
J. Vanderkerckhove,
H. Vanderbruggen,
M. Van Montagu,
M. Zabeau, and M. Vaeck.
1986.
Structural and functional analysis of a cloned delta endotoxin of Bacillus thuringiensis Berliner 1715.
Eur. J. Biochem.
161:273-280[Medline].
|
| 17.
|
Hunter, W. M., and F. C. Greenwood.
1962.
Preparation of iodine-131 labelled human growth hormone of high specific activity.
Nature (London)
194:495-496[CrossRef][Medline].
|
| 18.
|
James, R. R.,
B. A. Croft, and S. H. Strauss.
1999.
Susceptibility of the cottonwood leaf beetle (Coleoptera: Chrysomelidae) to different strains and transgenic toxins of Bacillus thuringiensis.
Environ. Entomol.
28:108-115.
|
| 19.
|
Keller, M.,
B. Sneh,
N. Strizhov,
E. Prudovsky,
A. Regev,
C. Koncz,
J. Schell, and A. Zilberstein.
1996.
Digestion of delta-endotoxin by gut proteases may explain reduced sensitivity of advanced instar larvae of Spodoptera littoralis to Cry1C.
Insect Biochem. Mol. Biol.
26:365-373[CrossRef][Medline].
|
| 20.
|
Knight, P.,
B. H. Knowles, and D. J. Ellar.
1995.
Molecular cloning of an insect aminopeptidase N that serves as a receptor for Bacillus thuringiensis CryIAc toxin.
J. Biol. Chem.
270:17765-17770[Abstract/Free Full Text].
|
| 21.
|
Lee, M.,
T. You,
B. Young,
J. Cotrill,
A. Valaitis, and D. Dean.
1996.
Aminopeptidase N purified from gypsy moth brush border membrane vesicles is a specific receptor for Bacillus thuringiensis Cry1Ac toxin.
Appl. Environ. Microbiol.
62:2845-2849[Abstract].
|
| 22.
|
Lorence, A.,
A. Darszon,
C. Díaz,
A. Liévano,
R. Quintero, and A. Bravo.
1995.
-endotoxins induce cation channels in Spodoptera frugiperda brush border membrane vesicles in suspension and in planar lipid bilayers.
FEBS Lett.
360:217-222[CrossRef][Medline].
|
| 23.
|
Luo, K.,
Y.-J. Lu, and M. J. Adang.
1996.
A 106 kDa form of aminopeptidase is a receptor for Bacillus thuringiensis CryIC -endotoxin in the brush border membrane of Manduca sexta.
Insect Biochem. Mol. Biol.
26:783-791[CrossRef].
|
| 24.
|
Munson, P. J., and D. Rodbard.
1980.
LIGAND: a versatile computerized approach for characterization of ligand-binding systems.
Anal. Biochem.
107:220-239[CrossRef][Medline].
|
| 25.
|
Navon, A.
1993.
Control of lepidopteran pests with Bacillus thuringiensis, p. 125-146.
In
P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs (ed.), Bacillus thuringiensis, an environmental biopesticide: theory and practice. John Wiley and Sons, Chichester, England.
|
| 26.
|
Oddou, P.,
H. Hartmann, and M. Geiser.
1991.
Identification and characterization of Heliothis virescens midgut membrane proteins binding Bacillus thuringiensis -endotoxins.
Eur. J. Biochem.
202:673-680[Medline].
|
| 27.
|
Pérez-Guerra, G.
1995.
Einflu der Nahrung auf die Empfindlichkeit von Nonnenlarven (Lymantria monacha Linné, 1758) (Lepidoptera: Lymantriidae) gegenüber Bacillus thuringiensis (Berliner, 1911).
Mitt. Dtsch. Ges. Allg. Angew. Entomol.
10:147-150.
|
| 28.
|
Rausell, C.,
A. C. Martínez-Ramírez,
I. García-Robles, and M. D. Real.
1999.
The toxicity and physiological effects of Bacillus thuringiensis toxins and formulations to T. pityocampa, the processionary caterpillar.
Pestic. Biochem. Physiol.
65:44-54[CrossRef].
|
| 29.
|
Salama, H. S.,
M. Ragaei, and M. Sabbour.
1995.
Larvae of Phthorimaea operculella (Zell) as affected by various strains of Bacillus thuringiensis.
J. Appl. Entomol.
119:241-243.
|
| 30.
|
Schnepf, E.,
N. Crickmore,
J. Van Rie,
D. Lereclus,
J. Baum,
J. Feitelson,
D. R. Zeigler, and D. H. Dean.
1998.
Bacillus thuringiensis and its pesticidal crystal proteins.
Microbiol. Mol. Biol. Rev.
62:775-806[Abstract/Free Full Text].
|
| 31.
|
Seyoum, A., and D. Abate.
1997.
Larvicidal efficacy of Bacillus thuringiensis var. israelensis and Bacillus sphaericus on Anopheles arabiensis in Ethiopia.
World J. Microbiol. Biotechnol.
13:21-24.
|
| 32.
|
Vadlamudi, R. K.,
E. Weber,
I. Ji,
T. H. Ji, and L. A. Bulla, Jr.
1995.
Cloning and expression of a receptor for an insecticidal toxin of Bacillus thuringiensis.
J. Biol. Chem.
270:5490-5494[Abstract/Free Full Text].
|
| 33.
|
Van Frankenhuyzen, K.
1990.
Development and current status of Bacillus thuringiensis for control of defoliating forest insects.
For. Chron.
56:498-507.
|
| 34.
|
Van Frankenhuyzen, K.
1993.
The challenge of Bacillus thuringiensis, p. 1-35.
In
P. F. Entwistle, J. S. Cory, M. J. Bailey, and S. Higgs (ed.), Bacillus thuringiensis, an environmental biopesticide: theory and practice. John Wiley and Sons, Chichester, England.
|
| 35.
|
Van Frankenhuyzen, K.,
J. L. Gringorten,
R. E. Milne,
D. Gauthier,
M. Pusztai,
B. Brousseau, and L. Masson.
1991.
Specificity of activated CryIA proteins from Bacillus thuringiensis subsp. kurstaki HD-1 for defoliating forest lepidoptera.
Appl. Environ. Microbiol.
57:1650-1655[Abstract/Free Full Text].
|
| 36.
|
Van Rie, J.,
S. Jansens,
H. Höfte,
D. Degheele, and H. Van Mellaert.
1989.
Specificity of Bacillus thuringiensis -endotoxins: importance of specific receptors on the brush border membrane of the midgut of target insects.
Eur. J. Biochem.
186:239-247[Medline].
|
| 37.
|
Van Rie, J.,
S. Jansens,
H. Höfte,
D. Degheele, and H. Van Mellaert.
1990.
Receptors on the brush border membrane of the insect midgut as determinants of the specificity of Bacillus thuringiensis delta-endotoxins.
Appl. Environ. Microbiol.
56:1378-1385[Abstract/Free Full Text].
|
| 38.
|
Wierenga, J. M.,
D. L. Norris, and M. E. Whalon.
1996.
Stage-specific mortality of Colorado potato beetle (Coleoptera: Chrysomelidae) feeding on transgenic potatoes.
J. Econ. Entomol.
89:1047-1052.
|
| 39.
|
Wolfersberger, M. G.
1990.
The toxicity of two Bacillus thuringiensis delta-endotoxins to gypsy moth larvae is inversely related to the affinity of binding sites on midgut brush border membranes for the toxins.
Experientia
46:475-477[CrossRef][Medline].
|
| 40.
|
Wolfersberger, M. G.,
P. Luthy,
A. Maure,
P. Pareti,
F. V. Sacchi,
B. Giordana, and G. M. Hanozet.
1987.
Preparation and partial characterization of aminoacid transporting brush border membrane vesicles from the larval midgut of the cabbage butterfly (Pieris brassicae).
Comp. Biochem. Physiol.
86:301-308[CrossRef].
|
Applied and Environmental Microbiology, April 2000, p. 1553-1558, Vol. 66, No. 4
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Copyright © 2000, American Society for Microbiology. All rights reserved.
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Abstract
Introduction
