Content uploaded by Cyril Vidau
Author content
All content in this area was uploaded by Cyril Vidau
Content may be subject to copyright.
Available via license: CC BY 4.0
Content may be subject to copyright.
Exposure to Sublethal Doses of Fipronil and Thiacloprid
Highly Increases Mortality of Honeybees Previously
Infected by
Nosema ceranae
Cyril Vidau
1,2
, Marie Diogon
1,2
, Julie Aufauvre
1,2
,Re
´gis Fontbonne
1,2
, Bernard Vigue
`s
1,2
, Jean-Luc
Brunet
3
, Catherine Texier
2
, David G. Biron
1,2
, Nicolas Blot
1,2
, Hicham El Alaoui
1,2
, Luc P. Belzunces
3
,
Fre
´de
´ric Delbac
1,2
*
1Clermont Universite
´, Universite
´Blaise Pascal, Laboratoire Microorganismes: Ge
´nome et Environnement, BP 10448, Clermont-Ferrand, France, 2CNRS, UMR 6023, LMGE,
Aubie
`re, France, 3INRA, UMR 406 Abeilles & Environnement, Laboratoire de Toxicologie Environnementale, Site Agroparc, Avignon, France
Abstract
Background:
The honeybee, Apis mellifera, is undergoing a worldwide decline whose origin is still in debate. Studies
performed for twenty years suggest that this decline may involve both infectious diseases and exposure to pesticides. Joint
action of pathogens and chemicals are known to threaten several organisms but the combined effects of these stressors
were poorly investigated in honeybees. Our study was designed to explore the effect of Nosema ceranae infection on
honeybee sensitivity to sublethal doses of the insecticides fipronil and thiacloprid.
Methodology/Finding:
Five days after their emergence, honeybees were divided in 6 experimental groups: (i) uninfected
controls, (ii) infected with N. ceranae, (iii) uninfected and exposed to fipronil, (iv) uninfected and exposed to thiacloprid, (v)
infected with N. ceranae and exposed 10 days post-infection (p.i.) to fipronil, and (vi) infected with N. ceranae and exposed
10 days p.i. to thiacloprid. Honeybee mortality and insecticide consumption were analyzed daily and the intestinal spore
content was evaluated 20 days after infection. A significant increase in honeybee mortality was observed when N. ceranae-
infected honeybees were exposed to sublethal doses of insecticides. Surprisingly, exposures to fipronil and thiacloprid had
opposite effects on microsporidian spore production. Analysis of the honeybee detoxification system 10 days p.i. showed
that N. ceranae infection induced an increase in glutathione-S-transferase activity in midgut and fat body but not in 7-
ethoxycoumarin-O-deethylase activity.
Conclusions/Significance:
After exposure to sublethal doses of fipronil or thiacloprid a higher mortality was observed in N.
ceranae-infected honeybees than in uninfected ones. The synergistic effect of N. ceranae and insecticide on honeybee
mortality, however, did not appear strongly linked to a decrease of the insect detoxification system. These data support the
hypothesis that the combination of the increasing prevalence of N. ceranae with high pesticide content in beehives may
contribute to colony depopulation.
Citation: Vidau C, Diogon M, Aufauvre J, Fontbonne R, Vigue
`s B, et al. (2011) Exposure to Sublethal Doses of Fipronil and Thiacloprid Highly Increases Mortality of
Honeybees Previously Infected by Nosema ceranae. PLoS ONE 6(6): e21550. doi:10.1371/journal.pone.0021550
Editor: Elizabeth Didier, Tulane University School of Public Health and Tropical Medicine, United States of America
Received March 16, 2011; Accepted June 1, 2011; Published June 28, 2011
Copyright: ß2011 Vidau et al. This is an open-access article distributed under the terms of the Creative Commons Attribution License, which permits
unrestricted use, distribution, and reproduction in any medium, provided the original author and source are credited.
Funding: This work was funded by a grant from the Centre National de la Recherche Scientifique (CNRS, MIE: Maladies Infectieuses et Environnement). C.V.
acknowledges the support of a Fellowship from the CNRS. J.A. and R.F. were supported by grants from the ‘‘Ministe
`re de l’Education Nationale de l’Enseignement
Supe
´rieur et de la Recherche’’. The funders had no role in study design, data collection and analysis, decision to publish, or preparation of the manuscript.
Competing Interests: The authors have declared that no competing interests exist.
* E-mail: frederic.delbac@univ-bpclermont.fr
Introduction
Honeybee pollination contributes to agriculture productivity
and biodiversity. One-third of food consumed in the world is
linked to the pollination activity of honeybees, representing a
global economic worth of 153 billion euros in 2005 [1]. A
significant and poorly understood decrease in honeybee popula-
tions, however, has been reported worldwide by beekeepers and
scientists. Several factors have been proposed to explain the
honeybee decline including nutrition, queen quality, intoxication
by pesticides and parasitic diseases [2].
The honeybee, Apis mellifera (Hymenoptera, Apoidea), may be
exposed to a wide range of pesticides when foraging or consuming
contaminated food (pollen and honey) stocked into the hive [3].
Two classes of systemic pesticides, neonicotinoids and phenylpyr-
azoles, are mainly suspected for negative effects on honeybee health.
There are intense debates about the use and the eventual restriction
of these pesticides. In many studies, the lack of knowledge about
their toxicological profile has prevented drawing conclusions about
a causal link between exposure to insecticides and the honeybee
decline [4]. This is partly due to the fact that the assessment of the
risk posed by pesticides is mainly based on the determination of
acute toxicity using LD
50
as the critical toxicological value [5]. This
approach is contested because it cannot account for chronic toxicity
and sublethal effects that are highly important elements of
neonicotinoid and phenylpyrazole toxicity in honeybees [6–8].
PLoS ONE | www.plosone.org 1 June 2011 | Volume 6 | Issue 6 | e21550
Indeed, low doses of neonicotinoids and phenylpyrazoles induce a
broad range of sublethal effects such as behavioral or physiological
alterations in honeybees and other beneficial arthropods [9].
The adverse effects usually induced by pesticides are limited by
the action of a large set of metabolic enzymes. Although
honeybees have fewer genes involved in detoxification than other
insects [10], they are not necessarily more sensitive to pesticides
[11]. In honeybees, detoxification processes occur mainly in both
midgut and fat body [12], very similar to those of mammals [13].
Induction of microsomal monooxygenases and glutathione-S-
transferase is one of the key mechanisms of insect sensitivity to
pesticides [13,14]. The role of detoxification enzymes, however, is
not limited to the protection of insect against the deleterious effects
of pesticides. These enzymes are also involved in the metabolism
of endogenous compounds such as hormones and pheromones
[10]. Therefore, changes in the activity of the detoxification system
can lead to variations in honeybee sensitivity to pesticides and
more generally to alteration of their physiological homeostasis.
Parasites may also impact insect homeostasis to promote their
development. This usually induces changes in insect development,
behavior, reproduction and parasite tolerance. Physiological
changes induced by parasitism can render insects more susceptible
to environmental stressors such as pollutants and may cause a
reduction of insect fitness [15]. This trend is particularly exploited
in the concept of integrated pest management (IPM) where
entomopathogenic parasites are used in association with insecti-
cides at low doses [16–18]. Honeybees are also victim to such joint
effects between parasites and insecticides. Potential interactions
between Nosema and pesticides have been firstly described by Ladas
in 1972 [19]. More recently, Alaux et al. demonstrated that co-
exposure to microsporidian parasites and imidacloprid weakens
honeybee [20]. This result corroborates the hypothesis of a multi-
factorial cause for the massive colony losses observed worldwide.
Two microsporidian species, Nosema apis and Nosema ceranae, are
the agents of two major diseases known as nosemoses A and C,
respectively [21]. Both species are obligate intracellular parasites of
adult honeybees. N. ceranae increases energetic demand in
honeybees [20,22] and decreases hemolymph sugar level [23].
Furthermore, N. ceranae infection significantly suppresses the
honeybee immune response [20,24] and increases ethyl -oleate
content (a primer pheromone which regulates worker behavioral
maturation) [25]. Finally, N. ceranae-infected honeybees have
shorter life-spans than uninfected honeybees [20,26].
There are no data related to the effect of N. ceranae on the
detoxification system of honeybees. Therefore, it is not possible to
consider the link between N. ceranae infection, detoxification capacity
of infected honeybees and their sensitivity to pesticides. However, we
hypothesized that detoxification system could be modified by Nosema
infection given that detoxification mainly occurs in the gut and that
this tissue is the site of N. ceranae proliferation. In this study, we
assessed the impact of N. ceranae infection on detoxification activity of
honeybees as well as their sensitivity to fipronil (phenylpyrazole) and
thiacloprid (neonicotinoid), two pesticides found at high levels in hives
[3]. Based on their oral LD
50
values, fipronil (LD
50
: 4.17 ng/bee) and
thiacloprid (LD
50
:17mg/bee) are considered highly and slightly
toxic, respectively, to honeybees [27]. We demonstrate, however, that
a daily exposure 1/100
th
concentration of the LD
50
significantly
affects the mortality rate of N. ceranae-infected honeybees.
Materials and Methods
Experimental procedures and artificial rearing
All experiments were performed with a mixture of honeybees
from three Buckfast colonies (crossed with the Apis mellifera mellifera
honeybee). We used 3 colonies to get sufficient emergent bees for
all the experiments (,2000 bees). We confirmed that these
colonies were free of Nosema by PCR using primers previously
described [26]. Two frames of sealed brood were taken in each
colony and placed in an incubator in the dark at 35uC with 80%
relative humidity. Emerging honeybees were collected, confined to
laboratory cages (Pain type) in groups of 50, and maintained in the
incubator for five days. During this time, the caged honeybees
were fed with candy (ApifondaH) and water ad libitum and were
supplied with pollen. To mimic the hive environment as much as
possible, a little piece of wax and a BeeboostH(Pherotech, Delta,
BC, Canada) releasing a queen’s mandibular pheromone, were
placed in each cage. After five days of feeding, six experimental
groups of individuals were created: (i) uninfected controls, (ii)
infected with N. ceranae, (iii) uninfected and chronically exposed to
fipronil, (iv) uninfected and chronically exposed to thiacloprid, (v)
infected with N. ceranae and chronically exposed 10 days post-
infection (p.i.) to fipronil, and (vi) infected with N. ceranae and
chronically exposed 10 days p.i. to thiacloprid.
Honeybees were first individually infected (see below honeybee
infection) and fed during 10 days with 50% (w/v) sugar syrup
supplemented with 1% (w/v) proteins (Provita’bee, Biove´
laboratory) 10 h per day and thereafter (14 h per day) were fed
with candy and water ad libitum. Each day, feeders were replaced
and the daily sucrose consumption was quantified. Ten days after
infection, honeybees were then exposed to fipronil or thiacloprid
by ingesting insecticide-containing sugar syrup ad libitum (see below
exposure to insecticides). Honeybees not exposed to insecticides
were fed ad libitum with 0.1% DMSO-containing sugar syrup. The
feeders were replaced and the daily insecticide consumption was
quantified. Throughout the experiment, each cage was checked
every morning and any dead honeybees removed and counted.
Honeybee infection
Spores of N. ceranae were obtained from honeybees infected
experimentally in our laboratory. After sacrifice, the intestinal tract
of infected honeybees was dissected and homogenized in PBS
using a manual tissue grinder. The suspension was filtered through
No. 1 Whatman mesh and the resulting suspension was cleaned by
centrifugation and resuspended in PBS. The spore concentration
was determined by counting with a hematocytometer chamber.
Nosema species was confirmed by PCR according the procedure
described by Higes et al. [26].
At 5 days post-emergence, caged bees were starved for 3 h, CO
2
anaesthetized and spread individually in ‘‘infection boxes’’
consisting of 40 ventilated compartments (3.56462 cm). Each
compartment was supplied with a tip containing 125,000 spores of
Nosema ceranae diluted in 3 mL of water. ‘‘Infection boxes’’ were
placed in the incubator and 1 h later, bees that have consumed the
total spore solution were again encaged (50 bees per cage). Non-
infected bees were similarly treated without N. ceranae spores in the
water.
Exposure to insecticides
At 10 days p.i., honeybees were exposed ad libitum to fipronil or
thiacloprid by ingesting insecticide-containing sugar syrup (50%
sucrose solution, w/v) supplemented with 1% protein (Provita’bee,
Biove´ laboratory). Stock solutions of fipronil (1 g/L) and
thiacloprid (5.1 g/L) were prepared in DMSO and diluted in
sugar syrup to obtain a final concentration of 50% sucrose, 1%
protein, 0.1% DMSO and 1 mg/L fipronil or 5.1 mg/L
thiacloprid. To expose honeybees to sublethal doses of insecticides,
the final concentrations were determined so that honeybees
absorbed daily an insecticide quantity corresponding to about 1/
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 2 June 2011 | Volume 6 | Issue 6 | e21550
100
th
of the LD
50
. The actual insecticide consumption was
quantified by measuring the daily amount of insecticide-containing
sugar syrup consumed per bee.
Preparation of microsomal and cytosolic fractions
Enzyme extraction was performed 10 days p.i. for control and
N. ceranae-infected honeybee groups. Honeybees were CO
2
-
anaesthetized and sacrificed by decapitation, the intestinal tract
was dissected, and the midgut was separated from the rectum. Ten
abdomens devoid of intestinal tract (containing the fat body) and
50 midguts were pooled in ice-cooled tubes and frozen at 280uC
until homogenization. They were homogenized twice at 4uC,
(TissuLyser
TM
; Qiagen; 5610 s at 30 MHz) in extraction buffer
(1% KCl, 1 mM EDTA, 1 mM PMSF, 0.1 mg/mL soybean
inhibitor and 50 mM Tris-HCl pH 7.6). The homogenates were
then centrifuged at 12,0006g for 20 min at 4uC and the resulting
supernatants centrifuged at 105,0006g for 60 min at 4uC. The
final supernatants (containing GST activity) were frozen at 280uC
and the microsomal pellets (containing ECOD activity) were
resuspended in 20% glycerol (v/v), 1 mM EDTA, 1 mM PMSF
and 100 mM sodium phosphate pH 7.4 and frozen at 280uC
until analysis. Protein concentration was estimated for each
preparation by the technique of Lowry et al. [28] using a standard
curve generated from known amounts (5–35 mg) of BSA.
7-ethoxycoumarin-O-deethylase (ECOD) assay
The measurement of 7-ethoxycoumarin-O-deethylase (ECOD)
activity was done as described by de Souza et al. [29]. Briefly,
ECOD activity was measured in midgut and abdomen devoid of
intestinal tract (i.e. the fat body) by adding 1 mg of microsomal
proteins to the reaction mixture containing, 0.5 mM NADP,
5 mM G6P, 10 mM MgCl
2
, 1 U G6PD, 200 mM 7-ethoxycou-
marin and 100 mM Tris-HCl pH 7.4. After 30 min of incubation,
the reaction was stopped with TCA 60% (p/v) then adjusted to
pH 9.0 with 1.6 M glycine-NaOH. The medium was centrifuged
at 30006g for 7 min and the production of 7-hydroxycoumarin
was quantified by recording the fluorescence (l
ex
380 nm and l
em
455 nm) with a fluorimeter (SFM 25, Kontron instruments)
apparatus. The amount of 7-hydroxycoumarin formed by the
ECOD activity assay was determined with a standard curve
generated from known amounts (0–100 nM) of 7-hydroxycou-
marin.
Glutathione-S-Transferase (GST) assay
Glutathione-S-Transferase (GST) activity was spectrophotomet-
rically assayed by measuring the conjugation of GSH to 1-chloro-
2,4-dinitrobenzene using a method adapted from Habig et al. [30].
GST activity was measured in midgut and abdomen by adding
enzymatic extract to the reaction mixture containing 1 mM
EDTA, 2.5 mM GSH, 1 mM 1-chloro-2,4-dinitrobenzene and
100 mM Na/K-phosphate pH 7.4. GST activity was quantified
by recording the appearance of conjugated product at 340 nm
during 5 min. GST activity was calculated using Beer Lambert
law with e
340
= 9,6 mM
21
?cm
21
.
Statistical analysis
Statistica 7.0 (StatSoft inc., Tulsa, USA) was used for the
statistical analysis. The significance thresholds were deemed as
significant (p#0.05), highly significant (p#0.01), or very highly
significant (p#0.001). Mann-Whitney U Test, a non-parametric
test, was used to compare the sucrose and insecticide consumptions
at 10 days p.i. Detoxification enzyme activities in both midgut and
fat body from control and N. ceranae-infected groups were compared
by using the Mann-Whitney U Test. The effect of N. ceranae
infection on honeybee sensitivity from exposure to insecticides was
analyzed with a survival analysis taking into consideration all groups
followed by a Cox-Mantel test (Life Tables) to determine the
significant difference between each group. The Wald-Wolfowitz
Runs Test (W-W Runs test) was used to compare insecticide uptake
in fipronil only, thiacloprid only, N. ceranae-fipronil and N. ceranae-
thiacloprid groups. A Kruskal-Wallis test combined with a multiple
comparison of mean ranks for all groups was used to determine the
effect of exposure to insecticides on N. ceranae spore production in
the digestive tract (i.e. midgut and rectum).
Results
Honeybee infection
The success of N. ceranae infection was monitored by measuring
sucrose consumption of honeybees and counting spores present in
their digestive tract 10 days p.i. Light microscopy analysis revealed
that numerous foci of N. ceranae were present at 10 days p.i. in the
epithelium cells of infected-bees (Fig. 1A). A mean of
18.4.10
6
60.4.10
6
spores per honeybee was measured in infected
honeybees whereas no spore was observed in uninfected honeybees.
PCR analysis confirmed the infection by N. ceranae for the infected
honeybee group (data not shown). Energetic stress was the main
symptom of Nosema infection. Thus, the sucrose consumption was
compared between infected and uninfected honeybees. For each
group (i.e. control and infected by N. ceranae), the amount of sucrose
consumed daily increased with time but differed between these two
groups (i.e. treatment) (Fig. 1B). Indeed, at 10 days p.i., honeybees
infected by N. ceranae consumed much more sucrose than uninfected
honeybees (M-W U test = 2.0, p = 0.0007).
Changes in detoxification enzyme activity in both midgut
and fat body
ECOD and GST are considered representative of phase I and
phase II activities of the detoxification system. The effect of N. ceranae
on ECOD activity was studied in fat body and mid-gut at 10 days p.i.
Infection induced no significant changes in ECOD activity both in
midgut (M-W U test = 9.0, p = 0.150) and fat body (M-W U
test = 16.0, p = 0.749) (Fig. 2A). Conversely, GST activity was highly
significantly increased i n midgut (M-W U test = 0, p = 0.003 948) and
fat body (M-W U test = 0.0, p = 0.003 948) of infected honeybees
(Fig. 2B). GST activity measured in midgut and fat body of infected
honeybees was increased 1.6-fold and 1.7-fold, respectively, com-
pared to the GST activity measured in uninfected honeybees.
Sublethal doses of fipronil and thiacloprid increases
mortality of Nosema ceranae-infected honeybees
The effect of exposure to insecticides on honeybee mortality was
assessed in both uninfected and N. ceranae-infected honeybees.
Survival analysis indicated that N. ceranae infection induced a very
highly significant increase in honeybee mortality compared to the
uninfected control group of honeybees (Cox-Mantel test, U = 31,78,
p,10
25
)(Fig. 3A). Exposures to fipronil (Cox-Mantel test,
U = 1.87, p = 0.517 68) and thiacloprid (Cox-Mantel test,
U = 0.96, p = 0.733 95) had no effect on the mortality of uninfected
honeybees compared to uninfected and untreated control group
over the duration of our experiments (Fig. 3A). Interestingly,
honeybee exposure to fipronil (Cox-Mantel test U = 38,41, p,10
25
)
and thiacloprid (Cox-Mantel test U = 28,26, p = p,10
25
) influ-
enced very highly significantly the amplitude and the time course of
N. ceranae-induced honeybee mortality (Fig. 3A). Indeed, honeybees
infected with N. ceranae and then exposed to insecticides died earlier
than bees only infected. In addition, at the end of the experiment (20
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 3 June 2011 | Volume 6 | Issue 6 | e21550
days p.i.), while mortality of only infected honeybees reached a
maximum of 47%, the mortalities observed in infected honeybees
exposed to fipronil and thiacloprid were not significantly different
from each other (Cox-Mantel test U= 210,4864, p = 0.132 65) and
reached a maximum of 82 and 71%, respectively (Fig. 3A).
Comparison of insecticide uptake in fipronil, thiacloprid, N. ceranae-
fipronil and N. ceranae-thiacloprid groups revealed that total
consumption of each insecticide at 20 days p.i. was not different
in non-infected and N. ceranae-infected honeybees for both fipronil
(W-W Runs test = 0.46, p = 0,648 08) and thiacloprid (W-W Runs
test = 1.37, p = 0,170 90). Also, the daily insecticide uptake was not
different in non-infected and N. ceranae-infected honeybees for both
fipronil (Fig. 3B) and thiacloprid (Fig. 3C). After exposure to
insecticides, uninfected honeybees did not display any signs of
intoxication. By contrast, at this level of exposure, insecticides
triggered aggressiveness and tremors in infected honeybees during
the first days of exposure, and later exhibited ataxia.
Effect of insecticide exposure on N. ceranae spore
production
Spore production was monitored at 20 days p.i. in all groups
(Fig. 4). No N. ceranae spore was observed in control uninfected
bees and insecticide-exposed uninfected honeybees (data not
shown). In N. ceranae-infected honeybees, a mean of 112.1610
6
(616.7) spores/bee was counted. Interestingly, statistical analysis
(Kruskal-Wallis test combined to a multiple comparison of mean
ranks for all groups) revealed that exposure to fipronil reduced
significantly (p = 0.0117 63) the spore production in infected bees
to 74.8610
6
(612.0) whereas exposure to thiacloprid enhanced
highly significantly (p = 0.0035, 99) the spore production up to
156.9610
6
(613.3) spores/bee.
Discussion
In this study, we showed that sublethal doses of a neonicotinoid
(thiacloprid) and of a phenylpyrazole (fipronil) highly increased
mortality of honeybees previously infected by the microsporidian
parasite N. ceranae. Although the exact mechanism involved in this
synergistic effect remains unclear, our data suggest that the
sensitization process is not strongly linked to a decrease of
detoxification capacity in infected bees or necessarily by an
enhancement of N. ceranae proliferation after exposure to insecticides.
During our experiments, no mortality was observed at 10 days
p.i. in infected honeybees. Numerous foci were visible in their
Figure 1. Infection monitoring. (A) Semi-thin sections of midgut epithelium of control (left) and infected (right) honeybees stained with toluidin
blue. Arrows indicate N. ceranae foci. Bar = 25 mm. (B) Effect of N. ceranae infection on sucrose consumption. Data represent the mean of cumulative
sucrose consumption (mg/bee +/2standard deviation, sd) from 9 replicates, each containing 50 honeybees.
doi:10.1371/journal.pone.0021550.g001
Figure 2. Effect of
N. ceranae
infection on ECOD (A) and GST (B) activities at 10 days p.i. in midgut and fat body of honeybees. Data
represent the mean of specific activity (nmoles/min/mg of proteins +/2standard deviation, sd) from 6 replicates (45 honeybees/replicate) of control
and infected honeybees. Asterisks indicate the level of significance at p,0.01 (**). ECOD: 7-ethoxycoumarin-O-deethylase; GST: Glutathione-S-
Transferase.
doi:10.1371/journal.pone.0021550.g002
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 4 June 2011 | Volume 6 | Issue 6 | e21550
epithelial cells and a mean of 18.4.10
6
(60.4.10
6
) spores/honeybee
was measured in their digestive tract. Our results contrast with
previous data by Higes et al. [26] who described 100% of
honeybee mortality at 8 days p.i. with N. ceranae, but are
comparable to mortality rate and spore production observed by
Paxton et al. [31]. In addition, the very highly significant
enhanced sucrose consumption by infected honeybees is consistent
with the energetic stress recently described by Mayack and Naugh
[22]. Thus, at 10 days p.i., we can consider that the infected
honeybees of our study displayed a level of N. ceranae invasion seen
in forager honeybees [32].
The first evidence of a synergistic interaction between Nosema
infection and insecticide exposure in honeybees was described by
Alaux et al. [20]. These authors demonstrated that Nosema spp
treatment combined with exposure to imidacloprid, another
neonicotinoid, resulted in a higher mortality of honeybees. Based
on these results, we hypothesized that N. ceranae infection could
alter the functioning of detoxification system. We assessed the
ECOD and GST activities to test this hypothesis. In insects,
ECOD and GST activities have often served as convenient
measures of overall phases I and II metabolizing enzyme activities.
In addition, levels of ECOD and GST activities have been
associated with sensitivity to insecticides [33,34]. Our results
showed that ECOD activity remained unchanged at 10 days p.i, in
fat body and midgut whereas GST activity increased significantly
in both tissues. Therefore, these data indicated that the higher
mortality observed after insecticide exposure in N. ceranae-infected
honeybees was not strongly linked to a decrease in detoxification
capacity. However, we cannot exclude that infection by N. ceranae
could modify other enzymes involved in detoxification of these
insecticides. Despite this observation, exposure to sublethal doses
of fipronil and thiacloprid increased the mortality rate in N. ceranae-
Figure 3. Effect of
N. ceranae
infection on honeybee sensitivity to insecticides. (A) Percentage of honeybee mortality of (i) uninfected
control (light green square), (ii) N. ceranae-infected (dark green square), (iii) uninfected and chronically exposed to fipronil (light red circle), (iv)
uninfected and chronically exposed to thiacloprid (light blue triangle), (v) N. ceranae-infected then chronically exposed to fipronil (dark red circle) and
(vi) N. ceranae-infected then chronically exposed to thiacloprid (dark blue triangle). The arrow indicates the time of exposure to insecticides (10 days
p.i.). Data represent the percentage of cumulative mortality calculated from 3 cages, each containing 50 honeybees. The means of fipronil (B) and
thiacloprid (C) consumptions (pg or ng/bee/day +/2standard deviation, sd) were daily-monitored until 20 days p.i. for both infected (dark red or
blue) and uninfected (light red or blue) honeybees.
doi:10.1371/journal.pone.0021550.g003
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 5 June 2011 | Volume 6 | Issue 6 | e21550
infected honeybees. Indeed, in our experimental procedure, at 10
day p.i. honeybees were chronically exposed to very low doses of
insecticides during 10 days. Fipronil consumption was not different
in infected and uninfected honeybees. The mean of daily fipronil
consumption corresponded to an exposure equivalent to the
LD
50
/158 (25.364.8 pg/bee) for uninfected honeybees and
LD
50
/148 (26.960.8 pg/bee) for infected honeybees. A compa-
rable exposure level has been previously considered as sublethal by
Aliouane et al. [35]. In the same way, honeybees exposed to
thiacloprid consumed a similar daily quantity of insecticides of
LD
50
/151 (112.164.4 ng/bee) and LD
50
/112 (152.868.7 ng/
bee) for uninfected and infected honeybees, respectively. As
suspected, these levels of fipronil and thiacloprid exposure had no
effect on the mortality of uninfected honeybees and on their
behavior. Surprisingly, the same levels of exposure caused
symptoms of poisoning in infected honeybees and influenced the
mortality rate.
Because this metabolic hypothesis failed to explain the
sensitization process observed with mortality data, we assessed
the effect of exposure to insecticides on spore production. Our
results indicated that exposure to fipronil and thiacloprid had
antagonist effects on spore production. Indeed, in comparison to
infected honeybees not exposed to insecticides, the spore
production decreased by about 33% during exposure to fipronil
whereas the spore production increased by 40% with thiacloprid
exposure. These results then, do not explain the mortality increase
observed in the presence of insecticides. First, exposures to fipronil
and thiacloprid induced an increase in mortality among infected
honeybees but had opposite effects on spore production. Second,
in the case of thiacloprid, the spore overproduction did not seem
sufficient to explain the enhancement of honeybees’ mortality.
The interactive effect seen between N. ceranae and insecticides on
honeybee mortality was consistent with the observations in
honeybees infected with Nosema sp and exposed to imidacloprid
[20]. While the synergistic effect observed by Alaux et al. [20]
seemed to be linked to an increased consumption of imidacloprid
by infected honeybees, the synergistic effect observed in our study,
however, was not due to increased food intake following infection.
These new data on the synergistic action of Nosema and insecticides
highlight that such interactions are not restricted to neonicotinoids
(imidacloprid, thiacloprid) but extend to other classes of insecti-
cides including phenylpyrazoles.
A further generalization of this phenomenon in honeybees
exposed to other insecticides would not be surprising. Several
classes of chemical insecticides have already shown potency to
interact synergistically with entomopathogenic fungi in other
insect species. These kinds of combination are commonly used in
integrated pest management because they counteract resistance to
insecticides of many insects [36] and allow reducing insecticide
doses spread in the environment [16,37,38]. For instance,
organophosphorus compounds (oxydemeton methyl) and pyre-
thrinoı
¨d (permethrin) insecticides used in combination with
Beauvaria bassiana, induced a higher impact on Spilarctia obliqua
[39] and Anopheles gambiae [36] survival, respectively, than the use
of these control agents alone. In general, the synergistic effect of
these combinations appears at insecticide doses considered
sublethal to the target insect [17,40]. As is our study, the major
pattern observed with these combinations included an increase in
insecticide toxicity (decrease in LD50 or LC50) and a decreased
time to onset of insect mortality. This suggests that susceptibility of
insects to pesticides is a more complex phenomenon than
previously thought. The influence of parasitism in the ecosystem
must be considered in toxicological studies. As shown in our study,
the use of the LD
50
as an indicator of systemic insecticide toxicity
leads to an underestimation of the deleterious effects induced in
infected honeybees. Indeed, we demonstrated that sublethal doses
of insecticides highly impacted Nosema-infected honeybee mortal-
ity. This precaution is important since N. ceranae spreads rapidly
and can affect more than 80% of honeybee colonies [41].
Numerous examples of interactions between chemicals and
pathogens that affect the insect lifespan have been described
[15,17,39,42–44]. Unfortunately, physiological mechanisms involved
in these interactions remain poorly understood and may even appear
to be contradictory. One of the current hypotheses explaining the
synergistic effect of such combinations suggests that pathogen
metabolites may interfere with the detoxification process [36,45].
Figure 4. Effect of exposure to insecticides on
N. ceranae
spore production. The spore production in the digestive tract (midgut and rectum)
was assessed at 20 days p.i. Data represent the mean numbers of spores/honeybee (610
6
+/2standard deviation, sd) from 15 honeybees, 20 days p.i.
Asterisks indicate the level of significance at p,0.05 (*) and p,0.01 (**).
doi:10.1371/journal.pone.0021550.g004
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 6 June 2011 | Volume 6 | Issue 6 | e21550
Reallocation of insecticide-detoxifying enzymes to counteract
parasitic infections possibly reduces the quantity of enzymes available
to target insecticides resulting in changes of insecticide toxicokinetics.
Thus, it is possible that the synergistic effect results in an effective
increase in sensitivity to insecticides in the presence of a proliferating
parasite infection. Ironically, the few published data about the effect
of parasitism on metabolizing enzymes of insects showed that a large
set of parasitic infections could activate several proteins implicated in
insect detoxification (e.g. CYP’s, GST, esterases) [45–50]. Consistent-
ly, we showed in our study, that 10 days after infection by N. ceranae,
the GST activity was enhanced in midgut and fat body of honeybees,
in agreement with the increase of the antioxidant activity recently
described in Nosema-infected queens [51]. This result contrasts with
the enhancement of infected-honeybees susceptibility to insecticides,
suggesting that GST would not be involved in detoxification process
of both fipronil and thiacloprid. However, the production of
microsomal monooxygenases is an inducible process [52] and it
remains possible that induction of detoxification genes in response to
exposure to insecticides was prevented by Nosema infection. Thus,
uninfected honeybees would be able to respond to insecticides by
enhancing detoxification process whereas infected honeybees may
not. This could explain the symptoms of intoxication observed in
infected honeybees.
Curiously, in most studies reporting synergistic effect of fungus/
insecticide combination, the impact of exposure to insecticides on
parasite virulence was not investigated [36,39,40,43]. In the rare
studies addressing this aspect, the parasite virulence was not
enhanced by the insecticides. Instead, despite the synergistic effect
on insect mortality, it appears that exposure to insecticides tends to
decrease germination or proliferation of the fungus [44]. Indeed,
insecticides have potential to affect the various developmental stages
of entomopathogenic fungi [18,53,54] to further justify why studies
of compatibility between parasites and insecticides are important for
developing IPM applications. In our study, fipronil and thiacloprid
have antagonist effect on N. ceranae proliferation whereby fipronil
decreases slightly spore production in honeybees. This effect can be
attributed either to the cytotoxic effect of fipronil on the intestinal
epithelium [55,56] or to its pro-oxidant action [57] that may affect
the reproduction cycle of N. ceranae, but this assertion should be
confirmed by other experiments. In contrast, thiacloprid increased
spore production in our study. This result was not consistent with
the observations done by Alaux et al. [20] who showed that
imidacloprid decreases slightly spore production in honeybees.
Thus, in our studies, the synergistic effect of N. ceranae infection and
exposure to insecticide did not appear to be linked to enhancement
of N. ceranae virulence by insecticides.
To conclude, our study confirms that interactions between N.
ceranae and insecticides constitute a significant risk for honeybee
health. The increasing prevalence of N. ceranae in European apiary
combined with the constant toxic pressure undergone by
honeybees, appears to contribute to the honeybee colony
depopulation. A better understanding of physiological effects
induced both by low doses of pesticides and Nosema infection seems
essential to elucidate the synergistic effects observed on honeybee
mortality. The discovery of molecular and cellular mechanisms
involved in the adverse effects induced by pathogens and pesticides
would confirm the influence of these stressors on honeybee health.
In addition, these data provide additional information that will
allow a better assessment of risk associated with these stressors and
highlight the urgent need of veterinary products for treating
nosemosis.
Author Contributions
Conceived and designed the experiments: CV NB HEA LPB FD.
Performed the experiments: CV MD JA RF BV J-LB. Analyzed the data:
CV DGB CT LPB FD. Contributed reagents/materials/analysis tools:
MD JA RF BV DGB. Wrote the paper: CV DGB LPB FD.
References
1. Gallai N, Salles J-M, Settele J, Vaissie`re BE (2009) Economic valuation of the
vulnerability of world agriculture confronted with pollinator decline. Ecological
Economics 68: 810–821.
2. Ratnieks FL, Carreck NL (2010) Ecology. Clarity on honey bee collapse?
Science 327: 152–153.
3. Mullin CA, Frazier M, Frazier JL, Ashcraft S, Simonds R, et al. (2010) High
levels of miticides and agrochemicals in North American apiaries: implications
for honey bee health. PLoS One 5: e9754.
4. Maxim L, van der Sluijs JP (2007) Uncerta inty: Cause or effect of stakeholders’
debates?: Analysis of a case study: The risk for honeybees of the insecticide
GauchoH. Science of The Total Environment 376: 1–17.
5. OEPP/EPPO (2003) Environmental risk assessment scheme for plant protection
products. . Bull OEPP/EPPO 33: 141–145.
6. Halm MP, Rortais A, Arnold G, Tasei JN, Rault S (2006) New risk assessment
approach for systemic insecticides: the case of honey bees and imidacloprid
(Gaucho). Environ Sci Technol 40: 2448–2454.
7. Rortais A, Arnold G, Halm M-P, Touffet-Briens F (2005) Modes of honeybees
exposure to systemic insecticides: estimated amounts of contaminated pollen and
nectar consumed by different categories of bees. Apidologie 36: 71–83.
8. Suchail S, Guez D, Belzunces LP (2001) Discrepancy between acute and chronic
toxicity induced by imidacloprid and its metabolites in Apis mellifera. Environ
Toxicol Chem 20: 2482–2486.
9. Desneux N, Decourtye A, Delpuech JM (2007) The sublethal effec ts of pesticides
on beneficial arthropods. Annu Rev Entomol 52: 81–106.
10. Claudianos C, Ranson H, Johnson RM, Biswas S, Schuler MA, et al. (2006) A
deficit of detoxification enzymes: pestici de sensitivity and envir onmental
response in the honeybee. Insect Mol Biol 15: 615–636.
11. Hardstone MC, Scott JG (2010) Is Apis mellifera more sensitive to insecticides
than other insects? Pest Manag Sci 66: 1171–1180.
12. Yu SJ, Robinson FA, Nation JL (1984) Detoxication capacity in the honey bee,
Apis mellifera L. Pesticide Biochemistry and Physiology 22: 360–368.
13. Terriere LC (1984) Induction of Detoxication Enzymes in Insects. Annual
Review of Entomology 29: 71–88.
14. Johnson RM, Wen Z, Schuler MA, Berenbaum MR (2006) Mediation of
pyrethroid insecticide toxicity to honey bees (Hymenoptera: Apidae) by
cytochrome P450 monooxygenases. J Econ Entomol 99: 1046–1050.
15. Holmstrup M, Bindesbol AM, Oostingh GJ, Duschl A, Scheil V, et al. (2010)
Interactions between effects of environmental chemicals and natural stressors: a
review. Sci Total Environ 408: 3746–3762.
16. Cameron PJ, Walker GP, Hodson AJ, Kale AJ, Herman TJB (2009) Trends in
IPM and insecticide use in processing tomatoes in New Zealand. Crop
Protection 28: 421–427.
17. Ericsson JD, Kabaluk JT, Goettel MS, Myers JH (2007) Spinosad interacts
synergistically with the insect pathogen Metarhizium anisopliae against the
exotic wireworms Agriotes lineatus and Agriotes obscurus (Coleoptera:
Elateridae). J Econ Entomol 100: 31–38.
18. Mohamed AK, Pratt JP, Nelson FR (1987) Compatibility of Metarhizium
anisopliae var. anisopliae with chemical pesticides. Mycopathologia 99: 99–105.
19. Ladas A (1972) Der einfluss verschiedener konstitutions- und umweltfaktoren auf
die anfa˜ ’’lligkeit der honigbiene (Apis mellifica l.)gegena˜œber zwei insektiziden
pflanzenschutzmitteln. Apidologie 3: 55–78.
20. Alaux C, Brunet JL, Dussaubat C, Mondet F, Tchamitchan S, et al. (2009)
Interactions between Nosema microspores and a neonicotinoid weaken
honeybees (Apis mellifera). Environ Microbiol 12: 774–782.
21. Higes M, Martin-Hernandez R, Meana A (2010) Nosema ceranae in Europe: an
emergent type C nosemosis*. Apidologie 41: 375–392.
22. Mayack C, Naug D (2009) Energetic stress in the honeybee Apis mellifera from
Nosema ceranae infection. J Invertebr Pathol 100: 185–188.
23. Mayack C, Naug D (2010) Parasitic infection leads to decline in hemolymph
sugar levels in honeybee foragers. J Insect Physiol 56: 1572–1575.
24. Antunez K, Martin-Hernandez R, Prieto L, Meana A, Zunino P, et al. (2009)
Immune suppression in the honey bee (Apis mellifera) following infection by
Nosema ceranae (Microsporidia). Environ Microbiol 11: 2284–2290.
25. Dussaubat C, Maisonnasse A, Alaux C, Tchamitchan S, Brunet JL, et al. (2010)
Nosema spp. infection alters pheromone production in honey bees (Apis
mellifera). J Chem Ecol 36: 522–525.
26. Higes M, Martin R, Meana A (2006) Nosema ceranae, a new microsporidian
parasite in honeybees in Europe. J Invertebr Pathol 92: 93–95.
27. Kievits JB, E (2009) Neurotoxiques syste´miques, un risque pour les abeilles?
Pesticides news. 76 p.
28. Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement
with the Folin phenol reagent. J Biol Chem 193: 265–275.
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 7 June 2011 | Volume 6 | Issue 6 | e21550
29. de Sousa G, Cuany A, Brun A, Amichot M, Rahmani R, et al. (1995) A
microfluorometric method for measuring ethoxycoumarin-O-deethylase activity
on individual Drosophila melanogast er abdomens: inter est for screening
resistance in insect populations. Anal Biochem 229: 86–91.
30. Habig WH, Pabst MJ, Jakoby WB (1974) Glutathione S-transferases. The first
enzymatic step in mercapturic acid formation. J Biol Chem 249: 7130–7139.
31. Paxton RJ, Klee J, Korpela S, Fries I (2007) Nosema ceranae has infected Apis
mellifera in Europe since at least 1998 and may be more virulent than Nosema
apis. Apidologie 38: 558–565.
32. Higes M, Martin-Hernandez R, Botias C, Bailon EG, Gonzalez-Porto AV, et al.
(2008) How natural infection by Nosema ceranae causes honeybee colony
collapse. Environ Microbiol 10: 2659–2669.
33. Bride JM, Cuany A, Amichot M, Brun A, Babault M, et al. (1997) Cytochrome
P-450 field insecticide tolerance and development of laboratory resistance in
grape vine populations of Drosophila melanogaster (Diptera: Drosophilidae).
J Econ Entomol 90: 1514–1520.
34. Enayati AA, Ranson H, Hemingway J (2005) Insect glutathione transferases and
insecticide resistance. Insect Mol Biol 14: 3–8.
35. Aliouane Y, El Hassani AK, Gary V, Armengaud C, Lambin M, et al. (2009)
Subchronic exposure of honeybees to sublethal doses of pesticides: effects on
behavior. Environ Toxicol Chem 28: 113–122.
36. Farenhorst M, Mouatcho JC, Kikankie CK, Brooke BD, Hunt RH, et al. (2009)
Fungal infection counters insecticide resistance in African malaria mosquitoes.
Proc Natl Acad Sci U S A 106: 17443–17447.
37. Mariyono J (2008) Direct and indirect impacts of integrated pest management
on pesticide use: a case of rice agriculture in Java, Indonesia. Pest Manag Sci 64:
1069–1073.
38. Weddle PW, Welter SC, Thomson D (2009) History of IPM in California pe ars–
50 years of pesticide use and the transition to biologically intensive IPM. Pest
Manag Sci 65: 1287–1292.
39. Purwar JP, Sachan GC (2006) Synergistic effect of entomogenous fungi on some
insecticides against Bihar hairy caterpillar Spilarctia obliqua (Lepidoptera:
Arctiidae). Microbiol Res 161: 38–42.
40. Pachamuthu P, Kamble ST (2000) In vivo study on combined toxicity of
Metarhizium anisopliae (Deuteromycotina: Hyphomycetes) strain ESC-1 with
sublethal doses of chlorpyrifos, propetamphos, and cyfluthrin against German
cockroach (Dictyoptera: Blattellidae). J Econ Entomol 93: 60–70.
41. Cox-Foster DL, Conlan S, Holmes EC, Palacios G, Evans JD, et al. (2007) A
metagenomic survey of microbes in honey bee colony collapse disorder. Science
318: 283–287.
42. Al Mazraawi MS (2007) Interaction effects between Beauveria bassiana and
imidacloprid against Thrips tabaci (Thysanoptera: Thripidae). Commun Agric
Appl Biol Sci 72: 549–555.
43. Farenhorst M, Knols BG, Thomas MB, Howard AF, Takken W, et al. (2010)
Synergy in efficacy of fungal entomopathogens and permethrin against West
African insecticide-resistant Anopheles gambiae mosquitoes. PLoS One 5:
e12081.
44. Furlong MJ, Groden E (2001) Evaluation of synergistic interactions between the
Colorado potato be etle (Coleoptera: Chrysomelidae ) pathogen Beauveria
bassiana and the insecticides, imidacloprid, and cyromazine. J Econ Entomol
94: 344–356.
45. Serebrov VV, Gerber ON, Maliarchuk AA, Martem’ianov VV, Alekseev AA,
et al. (2006) [The effect of ent omopathogenic fungi on the activity of
detoxicating enzymes of larvae of bee pyralid Galleria mellonella L.
(Lepidoptera, Pyralidee) and the role of detoxicating enzymes in the formation
of the insect’ resistance to entomopathogenic fungi]. Izv Akad Nauk Ser Biol. pp
712–718.
46. Felix RC, Muller P, Ribeiro V, Ranson H, Silveira H (2010) Plasmodium
infection alters Anopheles gambiae detoxification gene expression. BMC
Genomics 11: 312.
47. Gui Z, Hou C, Liu T, Qin G, Li M, et al. (2009) Effects of insect viruses and
pesticides on glutathione S-transferase activity and gene expression in Bombyx
mori. J Econ Entomol 102: 1591–1598.
48. Shiotsuki T, Kato Y (1999) Induction of carboxylesterase isozymes in Bombyx
mori by E. coli infection. Insect Biochem Mol Biol 29: 731–736.
49. Tchankouo-Nguetcheu S, Khun H, Pincet L, Roux P, Bahut M, et al. (2010)
Differential protein modulation in midguts of Aedes aegypti infected with
chikungunya and dengue 2 viruses. PLoS One 5.
50. Zibaee A, Bandani AR, Tork M (2009) Effect of the entomopathogenic fungus,
Beauveria bassiana, and its secondary metabolite on detoxifying enzyme activities
and acetylcholinesterase (AChE) of the Sunn pest, Eurygaster integriceps
(Heteroptera: Scutellaridae). Biocontrol Science and Technology 19: 485–498.
51. Alaux C, Folschweiller M, McDonnell C, Beslay D, Cousin M, et al. (2011)
Pathological effects of the microsporidium Nosema ceranae on honey bee queen
physiology (Apis mellifera). Journal of Invertebrate Pathology 106: 380–385.
52. Le Goff G, Hilliou F, Siegfried BD, Boundy S, Wajnberg E, et al. (2006)
Xenobiotic response in Drosophila melanogaster: sex dependence of P450 and
GST gene induction. Insect Biochem Mol Biol 36: 674–682.
53. Alizadeh A, Samih MA, Izadi H (2007) Compatibility of Verticillium lecani
(Zimm.) with several pesticides. Commun Agric Appl Biol Sci 72: 1011–1015.
54. Li DP, Holdom DG (1994) Effects of Pesticides on Growth and Sporulation of
Metarhizium anisopliae (Deuteromycotina: Hyphomycetes). Journal of Inverte-
brate Pathology 63: 209–211.
55. da Silva Cruz A, da Silva-Zacarin EC, Bueno OC, Malaspina O (2010)
Morphological alterations induced by boric acid and fipronil in the midgut of
worker honeybee (Apis mellifera L.) larvae: Morphological alterations in the
midgut of A. mellifera. Cell Biol Toxicol 26: 165–176.
56. Vidau C, Brunet JL, Badiou A, Belzunces LP (2009) Phenylpyrazole insecticides
induce cytotoxicity by altering mechanisms involved in cellular energy supply in
the human epithelial cell model Caco-2. Toxicol In Vitro 23: 589–597.
57. Lassiter TL, MacKillop EA, Ryde IT, Seidler FJ, Slotkin TA (2009) Is fipronil
safer than chlorpyrifos? Comparative developmental neurotoxicity modeled in
PC12 cells. Brain Res Bull 78: 313–322.
Toxico-Pathological Interactions Weaken Honeybees
PLoS ONE | www.plosone.org 8 June 2011 | Volume 6 | Issue 6 | e21550
