Wolbachia Symbiont Infections Induce Strong
Cytoplasmic Incompatibility in the Tsetse Fly Glossina
Uzma Alam1, Jan Medlock1¤a, Corey Brelsfoard1¤b, Roshan Pais1¤c, Claudia Lohs1¤d, Se ´verine Balmand2,
Jozef Carnogursky3, Abdelaziz Heddi2, Peter Takac3, Alison Galvani1, Serap Aksoy1*
1Yale University, School of Public Health, Division of Epidemiology of Microbial Diseases, New Haven, Connecticut, United States of America, 2INSA-Lyon, UMR203 BF2I,
INRA, Biologie Fonctionnelle Insectes et Interactions, Bat. Louis-Pasteur, Villeurbanne, France, 3Institute of Zoology, Section of Molecular and Applied Zoology, Slovak
Academy of Sciences, Bratislava, Slovakia
Tsetse flies are vectors of the protozoan parasite African trypanosomes, which cause sleeping sickness disease in humans
and nagana in livestock. Although there are no effective vaccines and efficacious drugs against this parasite, vector
reduction methods have been successful in curbing the disease, especially for nagana. Potential vector control methods
that do not involve use of chemicals is a genetic modification approach where flies engineered to be parasite resistant are
allowed to replace their susceptible natural counterparts, and Sterile Insect technique (SIT) where males sterilized by
chemical means are released to suppress female fecundity. The success of genetic modification approaches requires
identification of strong drive systems to spread the desirable traits and the efficacy of SIT can be enhanced by identification
of natural mating incompatibility. One such drive mechanism results from the cytoplasmic incompatibility (CI) phenomenon
induced by the symbiont Wolbachia. CI can also be used to induce natural mating incompatibility between release males
and natural populations. Although Wolbachia infections have been reported in tsetse, it has been a challenge to understand
their functional biology as attempts to cure tsetse of Wolbachia infections by antibiotic treatment damages the obligate
mutualistic symbiont (Wigglesworthia), without which the flies are sterile. Here, we developed aposymbiotic (symbiont-free)
and fertile tsetse lines by dietary provisioning of tetracycline supplemented blood meals with yeast extract, which rescues
Wigglesworthia-induced sterility. Our results reveal that Wolbachia infections confer strong CI during embryogenesis in
Wolbachia-free (GmmApo) females when mated with Wolbachia-infected (GmmWt) males. These results are the first
demonstration of the biological significance of Wolbachia infections in tsetse. Furthermore, when incorporated into a
mathematical model, our results confirm that Wolbachia can be used successfully as a gene driver. This lays the foundation
for new disease control methods including a population replacement approach with parasite resistant flies. Alternatively,
the availability of males that are reproductively incompatible with natural populations can enhance the efficacy of the
ongoing sterile insect technique (SIT) applications by eliminating the need for chemical irradiation.
Citation: Alam U, Medlock J, Brelsfoard C, Pais R, Lohs C, et al. (2011) Wolbachia Symbiont Infections Induce Strong Cytoplasmic Incompatibility in the Tsetse Fly
Glossina morsitans. PLoS Pathog 7(12): e1002415. doi:10.1371/journal.ppat.1002415
Editor: David S. Schneider, Stanford University, United States of America
Received March 22, 2011; Accepted October 17, 2011; Published December 8, 2011
Copyright: ? 2011 Alam 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 received support from NIH AI06892, GM069449 and Ambrose Monell Foundation awards to SA. 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. The corresponding author is Editor-in-Chief of PLoS Neglected Tropical
* E-mail: Serap.Aksoy@yale.edu
¤a Current address: Department of Mathematical Sciences, Clemson University, Clemson, South Carolina, United States of America
¤b Current address: Department of Entomology, University of Kentucky, Lexington, Kentucky, United States of America
¤c Current address: Department of Immunology, University of Connecticut Health Center, Connecticut, Farmington, United States of America
¤d Current address: Max F. Perutz Laboratories, Vienna, Austria
Tsetse flies are the sole vector of Human African Trypanosomiasis
(HAT), also known as sleeping sickness, caused by the protozoan
Trypanosoma brucei spp. in sub-Saharan Africa. Recent figures released
by the World Health Organization (WHO) indicate that the
devastating HAT epidemics, which started in the early 1990s, are
coming under control and may no longer represent a major public
health crisis [1–3]. While this news is welcoming, about 60 million
people continue to live in tsetse infested areas at risk for HAT in 37
countries, and those at high risk are in remote areas where disease
control isdifficult to implement .Diseases caused bytrypanosomes
in animals continue to be rampant in Africa and result in severe
economic and nutritional losses. The ability to curb infections in
animals stands to increase both economic and nutritional status of the
Unfortunately, the disease toolbox remains very limited. To date,
no vaccines have been developed for HAT, therapeutic treatments
are expensive and have serious side effects, and diagnostic tools are
inadequate . Reduction of tsetse populations, however has
proven as an effective method for disease control . Although
effective, implementation of vector control methods in remote
PLoS Pathogens | www.plospathogens.org1December 2011 | Volume 7 | Issue 12 | e1002415
regions of Africa where the disease is rampant is difficult, expensive
and relies on extensive community participation and thus has not
been widely exercised for human disease control . During an
endemic period however, vector control can be particularly
advantageous in the absence of continued active case surveillance
. Mathematical models indicate that parasite infection preva-
lence in the tsetse host is an influential parameter for HAT
epidemiology and disease dynamics . Thus, reducing vector
populations or reducing the parasite transmission ability of flies can
be most effective in preventing disease emergence.
Advances in tsetse biology offer novel strategies, one being a
population replacement approach to modify tsetse’s parasite
transmission ability (vector competence) by expressing trypanoci-
dal molecules in the gut bacterial symbiont fauna, termed
paratransgenic transformation strategy [6–10]. For the paratrans-
genic approach to be successful, gene drive mechanisms need to be
discovered to spread parasite resistant phenotypes into natural
populations. An alternative vector control approach currently
being entertained on the continent involves a population
eradication method, through sterile male releases (SIT) .
Genetic methods that induce reproductive male sterility are
superior to the currently available SIT strategy that relies on
chemical irradiation to induce male sterility.
Tsetse flies are infected with multiple bacterial symbionts. Two
of the symbionts are enteric: the obligate Wigglesworthia glossinidia
reside within bacteriocytes in the midgut bacteriome organ as well
as in milk gland accessory tissue , while commensal Sodalis
glossinidius reside both inter- and extra-cellularly in various tissues
. A large portion of Wigglesworthia’s proteome encodes vitamin
products that may be necessary to supplement the strictly
vertebrate blood meal diet of tsetse . Without the bacteriome
population of Wigglesworthia, tsetse flies have reduced egg
development and are infecund [15–18]. The third symbiont,
Wolbachia resides mainly in the reproductive tissues [13,19,20].
Tsetse females have an unusual viviparous reproductive biology.
Females develop a single oocyte per gonotrophic cycle. The oocyte
is ovulated, fertilized and undergoes embryonic development in-
utero. The resulting larva hatches and is carried in the intrauterine
environment through three larval instars before being deposited.
During its intrauterine life, the larva receives all of its nutrients in
the form of milk secreted by the female accessory glands, milk
glands. While Wolbachia is transovarially transmitted, the enteric
symbionts are maternally transmitted into tsetse’s intrauterine
larva through mother’s milk secretions . By providing
ampicillin in the blood meal diet, it has been possible to clear
the extracellular Wigglesworthia in the milk without damaging the
intracellular Wigglesworthia in the bacteriome . Thus, such
females remain fecund but give rise to sterile progeny that lack
Wigglesworthia (both bacteriome and milk gland populations) but
retain Wolbachia and Sodalis. As a result of the obligate role of
Wigglesworthia, it has not been possible to use tetracycline treatment
to cure Wolbachia infections, and the biological significance of
Wolbachia infections in tsetse has thus remained elusive.
Wolbachia infections associated with various insects have been
shown to cause a number of reproductive modifications in their
hosts, the most common being CI [22–24]. CI occurs when a
Wolbachia infected male mates with an uninfected female, causing
developmental arrest of the embryo. In contrast, Wolbachia infected
females can mate with either an uninfected male or a male infected
with the same Wolbachia strain and produce viable Wolbachia infected
offspring. This reproductive advantage of infected females results in
the spread of Wolbachia infectionsalong with other traits that infected
insects may exhibit [25,26]. Empirical studies and previously
developed models have shown that the reproductive advantage
provided byWolbachiamaybeableto drive desiredphenotypes along
with other maternally inherited genes, organelles and/or symbionts
into natural populations [27–30]. The Wolbachia type found in the
tsetse species Glossina morsitans morsitans belongs to the Wolbachia A
super group . In a number of insect systems, Wolbachia strains
belonging to the A super group have been associated with the CI
phenotype in the different hosts they infect .
Here we investigated the possible role of Wolbachia symbionts that
can be used to drive desirable tsetse phenotypes into natural
populations, or to induce natural reproductive male sterility for field
applications. We developed a dietary supplementation method that
fauna, including obligate Wigglesworthia and Wolbachia. We report on
the fitness parameters of the engineered symbiont-free lines and on
the level of CI expression after wild type and aposymbiotic flies are
crossed. A mathematical model was also developed to ascertain
whether Wolbachia infections in tsetse could be used to drive a disease
refractory phenotype into a natural population.
Dietary Supplementation with Yeast Extract Rescues
Fecundity in the Absence of the Obligate Wigglesworthia
In many insect systems, tetracycline supplemented diet is used to
generate Wolbachia free lines to demonstrate the functional role of
Wolbachia through mating experiments. Inseminated tsetse females
maintained on tetracycline-supplemented blood meals however do
not generate any viable progeny. This is because tetracycline
treatment damages the obligate intracellular Wigglesworthia present
in the midgut bacteriome structure (Figure S1) . These results
are similar to prior reports where damage to Wigglesworthia had
been found to reduce host fecundity [17,21,32].
The fecundity of fertile females maintained on various diets was
evaluated (Figure 1A). Specifically, the diet combinations were as
follows: (a) blood only, (b) blood and ampicillin, (c) blood and
tetracycline, (d) blood and yeast, (e) blood, ampicillin and yeast,
and (f) blood, tetracycline and yeast. We monitored the number of
Infections with the parasitic bacterium Wolbachia are
widespread in insects and cause a number of reproductive
modifications, including cytoplasmic incompatibility (CI).
There is growing interest in Wolbachia, as CI may be able
to drive desired phenotypes such as disease resistance
traits, into natural populations. Although Wolbachia
infections had been reported in the medically and
agriculturally important tsetse, their functional role was
unknown. This is because attempts to cure tsetse of
Wolbachia by antibiotic treatment damages the obligate
mutualist Wigglesworthia, without which the flies are
sterile. Here we have succeeded in the development of
Wolbachia free and still fertile tsetse lines. Mating
experiments for the first time provides evidence of strong
CI in tsetse. We have incorporated our empirical data in a
mathematical model and show that Wolbachia infections
can be harnessed in tsetse to drive desirable phenotypes
into natural populations in few generations. This finding
provides additional support for the application of genetic
approaches, which aim to spread parasite resistance traits
in natural populations as a novel disease control method.
Alternatively, releasing Wolbachia infected males can
enhance Sterile Insect applications, as this will reduce
the fecundity of natural females either uninfected or
carrying a different strain of Wolbachia.
Wolbachia Induces CI in Tsetse
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larva deposited in each group over a 40-day period when females
undergo two gonotrophic cycles (defined as time required for the
development of a single progeny in-utero). Under optimum
conditions the first gonotrophic cycle takes about 20–22 days for
development from egg to parturition. In subsequent gonotrophic
cycles females produce a larva every 9 to 11 days. As we had
previously shown, ampicillin treatment does not reduce fecundity
since it does not damage Wigglesworthia resident within bacter-
iocytes in the midgut, unlike tetracycline, which clears all bacteria
including Wigglesworthia and Wolbachia and induces sterility.
Accordingly, ampicillin-receiving flies remained fecund while
tetracycline receiving flies were rendered sterile.
Figure 1. The effects of antibiotic treatment on G. m. morsitans. (A) Effect of yeast supplementation on percent larval deposition over two
gonotrophic cycles between wild type flies maintained on normal blood supplemented with antibiotics (ampicillin or tetracycline) compared to flies
maintained on yeast supplementation. The sample size (n) is above each column, and is represented as the number of females alive at the beginning
of each gonotrophic cycle. (B) PCR analysis shows the GmmWtflies are positive for Wigglesworthia (Wig Thic), Sodalis (Sod Chit) and Wolbachia (Wol
Groel). In contrast offspring resulting from tetracycline treated females (A and B) lack all three of the symbionts. The bottom panel shows gDNA
quality as measured by tsetse b-tubulin. (C) Presence of Wolbachia infections in late developing egg chambers of GmmWtfemales. Nuclei are
indicated by the blue DAPI stain and Wolbachia is shown by the red stain (D&E) Presence and absence of Wolbachia infections in early developing
egg chambers of GmmWtand GmmApofemales respectively. (F) Comparison of adult longevity between female GmmWtand GmmApoover a forty-day
period on yeast supplemented diet. Error bars are reflective of standard error. Data points are offset for clarity.
Wolbachia Induces CI in Tsetse
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Yeast extract (10% w/v) provisioning of the blood meal rescued
fecundity of the females receiving tetracycline to similar levels as
that of wild type and ampicillin receiving flies (65%, 55% and 64%
over the first gonotrophic cycle and 53%, 58% and 49% over the
second gonotrophic cycles, respectively). However, yeast provision-
ing at 10% w/v had a cost on fecundity when compared to flies
maintained on normal blood meals, (92% versus 55% over the first
gonotrophic cycle and 92% and 58% over the second gonotrophic
cycle, respectively). Nevertheless, yeast supplementation was able to
rescue the tetracycline-induced sterility to levels comparable to
those observed for GmmWtreceiving yeast or ampicillin supplement-
ed blood meals, respectively (Figure 1A). Thus yeast supplemented
dietary regiment allowed us to develop two lines to analyze the
functional role of Wolbachia symbionts in tsetse biology; one lacking
all symbionts (GmmApo) and another lacking Wigglesworthia but still
retaining Wolbachia and Sodalis (GmmWig2).
The GmmApoprogeny resulting from the first and second
depositions of tetracycline treated mothers were tested for the
presence of Sodalis, Wigglesworthia and Wolbachia by a bacterium-
specific PCR-assay. The PCR-assay demonstrated the absence of all
three symbionts as early as the first deposition in both the male and
female GmmApoadults (Figure 1B). The absence of Wolbachia from the
reproductive tissues of GmmApofemales was also verified by
Fluorescent In Situ Hybridization (FISH) analysis (Figure 1E). In
contrast, Wolbachia was present in egg chambers during both early
andlate developmental stagesin GmmWtfemales (Figure 1C &D).For
analysis of longevity, survivorship curves were compared using the
compared to that of GmmWtadults maintained on the same yeast-
supplemented blood meal over 40 days (two-gonotrophic cycles). No
difference (X2=0.71, df=1, P=0.4) was observed in survivorship
comparisons between the two groups (Figure 1F).
The second line (GmmWig2) generated from ampicillin treated
females still retain their Wolbachia and Sodalis symbionts, while
lacking both Wigglesworthia populations as evidenced by FISH and
PCR amplification analysis . When maintained on yeast-
supplemented blood, this line (similar to GmmApo) also did not
display any longevity differences from the GmmWtadults sustained
on the same diet.
No Paternal Wolbachia Effect Evidence in Aposymbiotic
Tetracycline treatment has been shown to have a negative
impact on the fertility of Drosophila simulans males . To
determine if the fertility of GmmApomales is negatively affected, we
mated GmmWtfemales with either GmmWtor GmmApomales and
maintained all flies on yeast-supplemented blood meals. Larval
deposition and eclosion rates from both crosses were compared
using arcsin(sqrrt(x)) transformed data to ensure normality. No
significant difference was observed between the crosses for two
gonotrophic cycles (P.0.05) (Table 1). The mean larval deposition
rate for GmmWtfemales crossed with GmmWtmales was 0.68 and
0.65 for the first and second gonotrophic cycles respectively, while
the mean larval deposition rate for GmmWtfemales crossed with
GmmApomales was 0.87 and 0.89 for the first and second
gonotrophic cycles, respectively (Table 1). Similarly, no difference
in eclosion rates was observed between the two groups (P.0.05)
(Table 2). Of the pupae obtained in the first gonotrophic cycle
from the GmmWtcross, 82% underwent eclosion compared to 83%
for the cross between GmmWtfemales and GmmApomales. For the
second gonotrophic cycle, we observed 89% average eclosion for
pupae from GmmWtcrosses and 93% for pupae from GmmWt
females crossed with GmmApomales (Table 2). Taken together,
these results demonstrate the preservation of reproductive fitness
in GmmApomales and rule out possible paternal effects of Wolbachia
To determine the expression of Wolbachia-induced CI, cage
population crosses were setup between GmmWtand GmmApo
individuals. Cages were the experimental units and the data were
arcsin(sqrrt(x)) transformed to ensure normality. To estimate the
possible cost of reproductive fitness due to loss of Wigglesworthia, we
made use of GmmWig2flies. Since GmmWig2flies still retained
Wolbachia infections but lacked Wigglesworthia (as described earlier
and in Figure 1A), this line served as the control for the CI cross in
order to measure potential fecundity effects due to loss of
Wigglesworthia in the GmmApoline and possible yeast-supplementa-
Although CI typically manifests itself as embryonic lethality,
given the viviparous nature of reproduction in tsetse, we measured
larval deposition rates, which are reflective of both successful
embryogenesis and larvagenesis (Table 1). Differences in larval
deposition rates (number of larva deposited per female) over the two
gonotrophic cycles for all crosses were significant by ANOVA on
arcsin(sqrrt(x)) transformed data (ANOVA; first deposition, F4,
9=20.6, P,0.0001, second deposition, F4, 10=21.9, P#0.0001).
No differences in larval deposition were observed between the
crosses GmmWt6 GmmWt, GmmWig26 GmmWig2and GmmApo6
GmmApo(Table 1). However differences were observed in compar-
Table 1. CI expression shown by average larval deposition rates in crosses between GmmApofemales mated with GmmWtmales.
Larval deposition rate 1st
Larval deposition rate 2nd
0.6860.01ab; n=1080.6560.07ab; n=89
0.8760.06a; n=590.8960.16a; n=48
0.6160.20ab; n=490.5360.18b; n=26
0.1060.02c; n=440c; n=38
0.6860.14b; n=530.5960.07ab; n=50
Larval deposition rates for each gonotrophic cycle and each cross type replicate were determined by dividing the number of larvae deposited per day by the number of
remaining females in the cage on the day of larviposition, and summing the values for each gonotrophic cycle. Mean deposition rate values with different superscripted
letters are statistically different from each other (P,0.05) using Tukey-Kramer post hoc multiple comparison tests within each gonotrophic cycle, ie., a, b, and c are
significantly different from each other, c but not a and b are different from ab). n was calculated by combining the total number of females alive when the first larva
were deposited for the three replicates of each cross type. GmmWt= Wild-type flies with all three symbionts, GmmApo= flies treated with tetracycline that lack
Wigglesworthia, Sodalis, and Wolbachia, and GmmWig2= flies treated with ampicillin that lack only Wigglesworthia.
Wolbachia Induces CI in Tsetse
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isons of the GmmApo6GmmWtcross with all other crosses for the first
and second gonotrophic cycles (Table 1). Given that the GmmWig2
females that lack Wigglesworthia are equally fecund as GmmWt, the
strong incompatibility we observed in GmmApofemales when crossed
with GmmWtmales is likely due to Wolbachia mediated reproductive
affects, and not due to nutritional effects resulting from loss of the
obligate symbiont Wigglesworthia.
We found that GmmWtfemales were compatible with all male
infection types, while GmmApofemales were only compatible with
GmmApomales. Crosses of GmmApofemales and GmmWtmales
demonstrated a pattern of unidirectional CI (Table 1). Spermathe-
cae dissections of females in incompatible crosses that did not
deposit a larva revealed the presence of sperm, suggesting females
were inseminated and that lack of deposition was the result of CI.
We also found that larval deposition rates and pupal eclosion rates
showed similar patterns to large cage experiments when measured
in single-pair crosses (Table S2). Differences were observed in larval
deposition rates (number of larva deposited per female) over the two
gonotrophic cycles for all single-pair crosses (Kruskal-Wallis; first
deposition, x2=9.3, df=3, P=0.03, second deposition, x2=9.5,
df=3, P=0.02). No differences in larval deposition were observed
in pair-wise comparisons of the crosses GmmWt6GmmWt, GmmWt6
GmmApoand GmmApo6 GmmApo(Table S2). However differences
wereobserved incomparisons of the incompatibleGmmApo6GmmWt
cross with all other crosses for the first and second gonotrophic
cycles (Table S2). These results support strong CI expression driven
by the Wolbachia infection status in female flies.
Effect of Symbiont Infection on Host Eclosion
Other than reproductive modifications, Wolbachia infections have
been shown to affect the fitness of their insect hosts [34,35]. In this
study, differences in eclosion rates (Table 2) were observed in the first
gonotrophic cycle of crosses of GmmApo, GmmWt, and GmmWig2
individuals on arcsin(sqrrt(x)) data (ANOVA, first gonotrophic cycle,
F4, 11=7.5, P=0.0036, second gonotrophic cycle, F3,
P=0.13) (Table 2). No differences in eclosion rates were observed
in single pair crosses for both gonotrophic cycles (Kruskal-Wallis; first
gonotrophic cycle, x2=0.74, df=3, P=0.86, second gonotrophic
cycle, x2=0.31, df=2, P=0.85) (Table S2). To determine if observed
differences in eclosion rates were due to Wolbachia infection we
compared the GmmWig26GmmWig2and the GmmApo6GmmApocross,
since both strains lack Wigglesworthia infection, but one (GmmWig2)
harbors Wolbachia infection. There were no significant differences
however between these crosses (P.0.05) (Table 2), suggesting no
extensive effect of Wolbachia infection on host eclosion rates.
CI in Tsetse Manifests During Early Embryogenesis
The CI phenotype was further examined by analyzing the re-
productive tract physiology of tsetse females between incompatible
and compatible crosses during embryonic development. Under
normal conditions a single oocyte undergoes and completes
oogenesis during larvagenesis. In compatible crosses (R GmmWt6=
GmmWt) we observed that the reproductive tract contains a
developing larva in the uterus and a developing or completed
oocyte in one of the two ovaries (Figure 2A). In an incompatible
cross (R GmmApo6 = GmmWt) a developing oocyte is observed in
one of the ovaries in the absence of a developing larva in the
uterus, suggesting a disruption of embryogenesis or early larval
development (Figure 2C). The observation of an incomplete
oocyte in the absence of a developing larva in the uterus suggests
the failure and abortion of either an embryo or very young larva.
These observations differ from older GmmWtvirgin females.
Typically, GmmWtvirgin females undergo oogenesis but do not
undergo ovulation, which results in the development and eventual
accumulation of two oocytes in each of the ovaries. Larvae are
never observed in the uterus as developed oocytes are never
ovulated, or fertilized in adult virgin females (Figure 2B).
Spread of Wolbachia in Tsetse Populations
From the experimental data, we estimated the impact of CI on
tsetse population biology using a Bayesian Markov chain Monte
Carlo method. The transmission failure of Wolbachia from mothers
to developing oocytes was moderate: 10.7% [0.07%, 22.7%] of
progeny produced by GmmWtmothers were Wolbachia uninfected
(Table 3). In addition, the incompatibility between GmmWtmales
and GmmApofemales was strong: 79.8% [63.0%, 90.3%] of matings
between GmmWtmales and GmmApofemales did not result in viable
larvae as measured by pupal deposition. There was a significant
fecundity (number of larval progeny deposited) benefit for
Wigglesworhia infection: GmmWt
54.2%] higher fecundity than GmmWig2females. Furthermore,
Wolbachia infection alone was estimated to give a fecundity benefit
of 19.3% [29.2%, 57.9%]. This is an estimate of the fecundity
difference between hypothetical females carrying Wigglesworthia
and Sodalis but not Wolbachia and the experimental GmmWtfemales.
Most importantly, our model demonstrates that, given a large
enough initial release, Wolbachia infected individuals will success-
fully invade a tsetse population (Table 4). The fixation prevalence
of Wolbachia is estimated to be 96.9% [85.6%, 99.8%]. There may
exist a release threshold, which an initial release must be above in
order for Wolbachia to invade: the median was no release threshold
(i.e. 0%), but the upper end of the 95% credible interval was a
release of the size of 39.6% of the native population. The median
threshold value is zero because, despite imperfect maternal
transmission, the fecundity benefit of Wolbachia is strong enough
to allow Wolbachia to invade a naı ¨ve tsetse population from any size
initial release, no matter how small. In addition, the time to reach
fixation from a release of the size of 10% of the native population
females had 28.4% [8.5%,
Table 2. Eclosion rates (%) of deposited pupae.
Cross type% Pupal Eclosion 1stgonotrophic cycle % Pupal Eclosion 2ndgonotrophic cycle
8268.0a; n=67 8965.0a; n=61
8369.0a; n=459366.0a; n=38
60618.0ab; n=3452624.0a; n=27
17628.0b; n=4 NA; n=0
8867.0a; n=33 75613a; n=25
Mean % pupal eclosion values depicted by different superscripted letters are statistically different from each other (P,0.05) using Tukey-Kramer post hoc multiple
comparison tests within each gonotrophic cycle, i.e., a and b are significantly different from each other, both not different from ab. n = the total number of pupae
Wolbachia Induces CI in Tsetse
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can be relatively short: the median value was 529 days, however
the upper end of the 95% credible interval was undefined because
in more than 2.5% of samples, 10% initial release was below the
Sensitivity analysis showed that the model results are sensitive to
both assumed and estimated parameters (supplementary material
Text S1). In particular, time to fixation had the largest sensitivity
to the time to first deposition and large elasticities to Wolbachia-
and Wigglesworthia-related parameters, suggesting that improving
the estimates of these parameters would most effectively improve
the fidelity of the estimate of time to fixation.
Here, we report for the first time on the functional role of
Wolbachia infections in tsetse, which support the expression of CI.
Microscopic analyses of the CI expressing females show that loss of
fecundity results from early embryogenic failure. Essential for our
studies we have discovered that we can maintain Wolbachia cured
tsetse lines fertile by dietary provisioning of tetracycline supple-
mented blood meals with yeast extract, despite the fact that such
flies lack the obligate mutualist Wigglesworthia, which is essential for
tsetse’s fecundity. When incorporated into a mathematical model,
our results suggest that Wolbachia can be used successfully as a gene
driver and, the time to reach fixation is relatively short given a
large enough initial release: on the order of 1 to 2 years. These
results provide a first insight into the role of Wolbachia infections in
a viviparous insect and indicate that Wolbachia mediated CI can
potentially be used to drive desirable tsetse phenotypes into natural
Our data presented here as well as previous results from other
studies indicate that in the absence of Wigglesworthia, tsetse females
are rendered sterile. Our prior studies where we maintained
inseminated flies on ampicillin supplemented blood diets resulted
in progeny deposition. This is because ampicillin treatment did not
affect the intracellular Wigglesworthia resident in the bacteriome
organ in the midgut, which provides essential nutrients to maintain
Figure 2. Wolbachia-induced CI phenotype in G. m. morsitans. Normal reproduction between GmmWtfemales and males is discernible by a
developing oocyte indicated by the white arrow and the presence of a larva in the uterus indicated by the pink arrow, following the first gonotrophic
cycle. (B) Unmated adult female tsetse. Unmated GmmWtfemales have an empty uterus and multiple developing oocytes indicated by white arrows.
Note the transparent nature of the spermatheca reflective of lack of sperm (C) Manifested CI. CI is indicated by GmmApofemales mated with GmmWt
males by the absence of a larva in the uterus and deformed embryo indicated by the blue arrow. Many of these embryos were aborted without
hatching into larva. Orange arrows indicate spermathecae in each image. Images were collected forty days (corresponding to the second gonotrophic
cycle) post mating.
Table 3. Cytoplasmic-incompatibility parameter estimates.
ParameterMedian95% Credible Interval
Fecundity Benefit of Wolbachia (sf,Wol)
Fecundity Benefit of Wigglesworthia
0.2839 [0.0854, 0.5420]
CI Strength (sh)
Transmission Failure (m)
Shown are the posterior median and 95% credible interval from Bayesian
Markov chain Monte Carlo estimation.
Table 4. Population-genetics quantity estimates: the
posterior median and 95% credible interval from Bayesian
Markov chain Monte Carlo estimation.
EstimateMedian 95% Credible Interval
Wolbachia Fixation Prevalence 0.9689[0.8559, 0.9984]
Release Threshold 0.0000[0.0000, 0.3958]
Time to Fixation (days)529 [296, ——]
Wolbachia fixation prevalence is the level at which Wolbachia is stably present
in the population. Release threshold is the number of Wolbachia-positive tsetse
that must be released into a Wolbachia-free population in order for Wolbachia
to ultimately go to fixation, relative to the size of the existing population. Time
to fixation is the number of days required to go from 10% initial Wolbachia
prevalence to 95% of the fixation prevalence; its upper 95% CI is undefined
because for more than 2.5% of samples, a release of 10% of the population is
below the release threshold so that Wolbachia is driven from the population for
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tsetse host fecundity . Antibiotic ampicillin treatment however
eliminated the extracellular Wigglesworthia population present in
the milk gland essential for symbiont transmission, and thus the
resulting progeny from such females lacked Wigglesworthia
(GmmWig2). Such progeny were reproductively sterile although
they retained the symbiont Wolbachia. The tetracycline diet
eliminated both intracellular and extracellular forms of Wiggle-
sworthia and thus we did not obtain any viable progeny from
inseminated females that were maintained on the tetracycline only
diet. Prior studies showed that tetracycline blood meals supple-
mented with vitamin B1 could partially rescue fertility , but in
our experiments vitamin supplementation could give rise to at
most one progeny deposition, which either did not hatch or did
not survive as an adult (data not shown). In sharp contrast,
supplementation of the blood meal diet with 10%(w/v) yeast-
extract reverted sterility in tetracycline treated flies to levels
comparable to GmmWtand GmmWig2females receiving the same
diet (Figure 1A). Although we have compared the fecundity of all
three lines for two gonotrophic cycles here, yeast supplemented
flies continue to deposit four to five progeny (data not shown).
Given the complex nature of the yeast extract (peptides, amino
acids, vitamins and other yeast cell components), it is difficult to
know the exact nature of the essential nutrients it provides, but we
believe that it could be working via supplementation of lipids and/
or essential vitamins that are lacking in the strict blood diet of
tsetse. However, we did observe some negative effect attributable
to the yeast diet when the fecundity of GmmWtflies receiving yeast
supplemented blood meals is compared to those receiving normal
blood diets. As such, we are further investigating the use of
different yeast supplementations and/or concentrations in an
effort to improve the diet efficiency. Nevertheless the availability of
Wolbachia-cured flies (GmmApo) allowed us to begin to understand
the functional role of this symbiosis.
In addition to Wolbachia symbiont specific PCR amplification,
we confirmed the absence of Wolbachia from the reproductive
tissues of GmmApofemales by FISH analysis. We show the presence
of Wolbachia in GmmWtfemales, isolates to a pole late in
development (Figure 1C). There are a number of studies in other
model systems that have investigated the link between Wolbachia
localization during spermatogenesis and density effects on CI
[36,37]. However, other studies have found no correlation
between Wolbachia density and CI during spermatogenesis
[38,39]. There have also been a number of studies investigating
Wolbachia localization during oogenesis [40–42]. Different Wolba-
chia strains in Drosophila embryos display posterior, anterior, or
cortical localization congruent with the classification based on the
wsp gene sequence . A positive correlation between levels of
Wolbachia at the posterior pole and CI has been suggested, but this
has yet to be examined in detail . Not withstanding, assessing
the role of Wolbachia during oogenesis is important, given that
factors promoting CI rescue are deposited in the egg cytoplasm
during oocyte development  and bacterial deposition in the
oocyte is an essential even for efficient maternal transmission.
Before we could perform crossing experiments to assess for CI,
we evaluated the effect of Wolbachia clearance on male reproduc-
tive capacity. This evaluation is important given that tetracycline
has been shown to negatively affect reproductive fitness in
Drosophila simulans . Additionally, the importance of this
finding is highlighted by a study of the mosquito A. albopictus
system in which the natural Wolbachia strains (wAlbA and wAlbB)
were cleared and transinfected with the Wolbachia strain wRi from
D. simulans . Their results showed that the wRi transinfected
males have a reduced mating capacity compared with the wild
type super infected males . In contrast, in our system, no
decrease in mating capacity was observed in GmmApomales
compared with GmmWtmales under the laboratory conditions. Our
observation agrees with the evolutionary model proposed by
Charlat et al., , where Wolbachia is exclusively maternally
transmitted therefore males may be considered an evolutionary
dead end in terms of Wolbachia infection . Consequently, no
direct selection by Wolbachia can be theoretically expected on
paternal reproductive fitness.
Loss of fecundity in the cross (R GmmApox = GmmWt) could
conceivably arise from loss of Wigglesworthia-mediated nutritional
benefits in GmmApofemales rather than to Wolbachia mediated CI.
To test this possibility, we compared the larval deposition rates in
crosses between RGmmApo6 = GmmApoand RGmmWig26 =
GmmWig2flies (Table 1). Our results show no statistically significant
differences between these crosses indicating that loss of fecundity
in the CI cross is not due to loss of Wigglesworthia.
Our empirical results were used to parameterize a population
genetic model of the spread of Wolbachia. Our model demonstrated
that GmmWtwould successfully invade an uninfected natural
population with a large enough release given CI rates. Indeed,
uninfected natural populations and natural populations with low
infection prevalence have recently been identified for multiple
tsetse species . This modeling result is consistent with the
natural spread of Wolbachia in Drosophila populations [48–50]. In
addition, the rise to the predicted fixation prevalence of between
86% and 100% is rapid. Apparently, the Wolbachia-mediated CI
has the potential to rapidly and effectively drive a desirable
phenotype into natural populations. We have previously been able
to culture and genetically transform the commensal symbiont of
tsetse, Sodalis glossinidius . It has also been possible to
reintroduce the transformed Sodalis into tsetse, called a paratrans-
genic approach [52,53]. Given that Sodalis resides in close
proximity to pathogenic trypanosomes in tsetse’s midgut, products
expressed in recSodalis can have an immediate effect on
trypanosome biology. The potential paratransgenic strategy in
tsetse could harness the Wolbachia mediated CI to drive a
recombinant Sodalis strain that would encode parasite resistance
genes into natural populations [6,10]. Our studies on the maternal
transmission dynamics of tsetse’s symbionts in the laboratory
indicated perfect transmission of both Wolbachia and Sodalis into
tsetse’s sequential progeny . This high transmission fidelity of
the two symbionts, coupled with strong nearly 100% CI caused by
Wolbachia would serve paratransgenic applications favorably.
An alternative control strategy to paratransgenic population
replacement strategy would be use CI as part of an incompatible
insect technique (IIT), which is analogous to a SIT approach
[29,55–58]. In a Wolbachia-based SIT approach female sterility is
artificially sustained by repeated releases of cytoplasmically
incompatible males. Similar to SIT, the increasing ratio of
incompatible matings over time can lead to population suppres-
sion. The benefit of an IIT strategy is that it would not require the
use of irradiation or chemosterilants to sterilize males prior to
release, which often reduces the fitness of released males, but
would rely on the naturally induced sterility of an incompatible
Wolbachia infection . A Wolbachia-based paratransgenic and
IIT control strategy for tsetse would rely upon the introduction of
a novel infection type into a population with an existing infection
that could result in bi-directional CI or the introduction of a novel
infection into an uninfected host population. Typically, in other
insect systems novel Wolbachia infections are established by
embryonic microinjections [60,61]. This would be difficult in
tsetse given their viviparous reproductive biology, in that adult
females carry and nourish their offspring for their entire larval
developmental cycle making injections of embryos difficult. Future
Wolbachia Induces CI in Tsetse
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studies however can focus on the introduction of novel infection
types via microinjection in aposymbiotic and naturally infected
adult flies . Maternal intrathoracic injections of Wolbachia
infection establishment has also been successful in Aedes aegypti .
There has been a growing interest in understanding the variety
of Wolbachia induced phenotypes in arthropods given the impact
that Wolbachia infections could potentially have on genetic
variation and host speciation impacting evolution of the species.
Our data add to this growing field, as this is the first demonstration
of the biological significance of Wolbachia infections in tsetse.
Interestingly, CI in tsetse appears to be strong in that by the
second gonotrophic cycle 0% of the females in an incompatible
cross give rise to progeny. This is an exception given that in many
insect systems incomplete CI is observed [27,64]. Future studies
with natural populations would now be important to confirm some
of the parameters we report here including maternal transmission
rates, infection prevalence and the maternal linkage efficacy
between Wolbachia and other maternally transmitted symbionts
such as Sodalis, which is being entertained for paratransgenic
Additionally, the aposymbiotic lines generated in this study are
currently being used to address the interactive role of trypanosome
transmission in tsetse. The importance of which is highlighted by
recent studies that have shown that Wolbachia infections may
impact host immune biology, limiting pathogen proliferation in
insect hosts [65–70].
Materials and Methods
The Glossina morsitans morsitans colony maintained in the
insectary at Yale University was originally established from
puparia collected in Zimbabwe. Newly emerged flies are separated
based on sex and mated at three to four days post eclosion. Flies
are maintained at 2461uC with 50 – 55% relative humidity and
fed defibrinated bovine blood (HemoStat Laboratories, CA) every
forty eight hours using an artificial membrane system .
Selective elimination of natural tsetse endosymbionts was obtained
as described below.
Wild type (GmmWt) fertile females were maintained on blood
meals supplemented with 10% (w/v) yeast extract (Becton
Dickinson) and 20 ug/ml of tetracycline. The yeast extract was
briefly boiled in water before being added the blood meal each
time. Flies were fed every 48 h using an artificial membrane
feeding system (as above) for the duration of their life span. The
resulting progeny are aposymbiotic (GmmApo) in that they lack their
natural endosymbionts, Wigglesworthia and Wolbachia. These
GmmApolines were maintained on blood meals supplemented with
10% (w/v) yeast extract without tetracycline.
fertile females were maintained on blood meals
supplemented with 50 ug/ml of ampicillin. The resulting progeny
do not have Wigglesworthia (GmmWig2), and were maintained on
blood meals supplemented with 10% (w/v) yeast extract without
Monitoring the Fecundity Cost of Yeast-extract
Newly eclosed aged matched females and males were divided
into six groups and copulation observed. Three of these groups
were provided with either normal blood meals (control) or blood
meals supplemented with ampicillin at 50 ug/ml or tetracycline at
20 ug/ml. Whereas the remaining three groups received blood
meals supplemented with 10% (w/v) yeast extract with either
ampicillin (50 ug/ml) or tetracycline (20 ug/ml). The cages were
monitored daily for pupal deposition and fly mortality over two
gonotrophic cycles (40 days). Fecundity was quantified by
determining the number of fecund females relative to total
number of females alive at the end of the gonotrophic cycle to
give an average percent of females depositing pupae. Each group
was setup with 100 females per cage.
Symbiont Prevalence Assay
Total DNA was extracted from adults eight days post eclosion
using the Qiagen Blood and Tissue extraction kit under
manufacturers conditions (Qiagen Kit #, 69506. CA). The
presence of the symbionts Sodalis, Wigglesworthia and Wolbachia
was determined by a species-specific PCR amplification assay
using the primer sets and conditions described (Table S1). For
input DNA quality control, the tsetse gene b-tubulin (GmmTub)
specific primer set was used. All PCR reactions were performed in
an MJ-Research thermocycler and the amplification products
were analyzed by electrophoresis on a 1% agarose gel and
visualized using image analysis software.
Wolbachia Infection Status by FISH
Dissected reproductive tracts from GmmWtand GmmApofemales
were fixed in 4% paraformaldehyde (PFA), embedded in paraffin,
cut into 5 mm thick sections and mounted on polyL-lysine coated
microscopy slides. After dewaxing in methylcyclohexane and
rehydration the sections were processed using the FISH protocol
previously described in Anselme et al. 2006 . Slides were
covered with a drop of 70% acetic acid and incubated at 45uC
until drop had dried, followed by dehydration and a 10 min
deproteinization step in 0.01N HCl/pepsine at 37uC. Slides were
then dehydrated again, prehybridized for 30 min at 45uC and
hybridized for 3 h at 45uC with 59 end rhodamine labeled 16S
RNA probes (59-AAT CCG GCC GAR CCG ACC C -39) and
(59-CTT CTG TGA GTA CCG TCA TTA TC -39). Microscopic
analyses were conducted using a Zeiss Axioskop2 microscope
equipped with an Infinity1 USB 2.0 camera and software
(Lumenera Corporation). Fluorescent images were taken using a
fluorescent filter set with fluorescein, rhodamine and DAPI specific
Monitoring Longevity of GmmApoand GmmWtFemales
GmmApoand GmmWtflies that emerged within a 24-hour period
(teneral) were collected, mated with GmmApomales at a ratio of 5:2
and copulation was observed. After six days males were removed
from experimental cages. Six independent cages were set-up for
both GmmApoand GmmWtgroups, comprising of a total of 169
GmmApoand 170 GmmWtfemales, respectively. Both the males and
females used represented offspring acquired from different
gonotrophic cycles (1stand 2nd). All flies were maintained on
yeast extract supplemented blood meals and fly mortality was
monitored daily over a 40-day period.
CI Mating Crosses
To determine the expression of CI, reciprocal crosses were set
up between GmmApo, GmmWtand GmmWig2flies, in triplicate. Cages
with a minimum of 15 females and 7 males each were set-up in the
following combinations: 1) R GmmWt6= GmmWt, 2) R GmmWt6
= GmmApo, 3) R GmmApo6 = GmmApo, 4) R GmmApo6 =
GmmWtand 5) R GmmWig26= GmmWig2. All flies received yeast
Wolbachia Induces CI in Tsetse
PLoS Pathogens | www.plospathogens.org8December 2011 | Volume 7 | Issue 12 | e1002415
supplemented blood meal diets. Flies were observed over two-
gonotrophic cycles with daily recording of mortality, larval
deposition dates, pupal eclosion dates and sex of emergent
progeny. Larval deposition rates for each gonotrophic cycle were
determined by dividing the number of larvae deposited per day by
the number of remaining females in the cage on the day of
larviposition and summing the values for each gonotrophic cycle.
At the conclusion of the experiment, all females were checked for
insemination by examination of dissected spermatheca for the
presence of sperm microscopically. Additionally, single line crosses
consisting of a single female and male per cage were set up (Table
S2). For the R GmmWt6= GmmWta total of 31 crosses were set up.
Also set up were 40 crosses for R GmmWt6 = GmmApo, 20 for R
GmmApo6= GmmApoand 33 for R GmmApo6= GmmWt. Both the
males and females used in these crosses represented offspring
acquired from different gonotrophic cycles to rule out batch
affects. Spermathecae of females was also dissected to confirm
Here we will briefly describe the mathematical modeling used in
this study; full details are available in the supplementary material
(Text S1). The data from mating crosses were modeled as samples
from the standard binomial random variable, with probability of
larval deposition per mated female per gonotrophic cycle, and
using a different probability for each cross. Following the empirical
findings regarding Wolbachia -mediated CI in Drosophila , the
probabilities were then defined in terms of four mechanistic
parameters: the probability of reproduction success (larval deposit)
from a cross between an GmmApofemale and an GmmApomale
(fT), the proportion of Wolbachia-free eggs of Wolbachia-carrying
mothers (m), the relative benefit to reproduction success of
Wolbachia infection to females (sf,Wol), the relative benefit to
reproduction success of Wigglesworthia infection to females (sf,Wig),
and the proportion of fertilizations of Wolbachia-free eggs by
larval-deposition probabilities in terms of these parameters are
are not viable(sh). The
where the subscripts refer to the types of the female and male,
respectively, with W for wild type (GmmWt), T for tetracycline
treated (GmmApo), and A for ampicillin treated (GmmWig2).
In addition to these mechanistic parameters, we also estimated
population-genetic quantities fundamental to the invasion of
Wolbachia into a novel tsetse population. Again following existing
models for Wolbachia-induced CI in Drosophila , a mathematical
model was developed for the temporal evolution of tsetse
abundance with and without Wolbachia infection. We incorporated
the Wolbachia-mediated CI trade-off of the fitness cost to male hosts
in reducing their mating success with uninfected females versus the
fitness benefit to female hosts in allowing them to successfully mate
with both infected and uninfected males (in addition to direct
effects of Wolbachia on fecundity and mortality).
For some values of the mechanistic parameters, these mo-
dels exhibit a threshold for Wolbachia invasion into the host
population: if, in a novel population, the proportion that is
initially Wolbachia infected is above the threshold, Wolbachia will
continue to stable fixation in the population at a high level. If the
proportion infected is below the threshold, Wolbachia will be
driven out of the population over time. This threshold level was
calculated, along with the prevalence of Wolbachia at fixation, and
the time to fixation. For the population-genetic model, several
parameters could not be estimated from the data on mating
crosses. Thus, we also performed a sensitivity analysis on these
parameters, along with the parameters estimated from the
To estimate both the mechanistic parameters for CI and the
population-genetics quantities derived from these parameters, a
Bayesian Markov chain Monte Carlo (MCMC) method was used
with uninformative prior distributions for the parameters .
morsitans. Images of bacteriome sections stained with Giemsa (A)
bacteriome organ showing bacteriocytes harboring Wigglesworthia
from a female maintained on normal bloodmeals, image taken at
10x magnification (B) bacterioctes taken 40 magnification from a
female maintained on ampicillin supplemented diet. A normal
bacteriome structure is retained on the ampicillin diet allowing for
continued fertility of such females. (C and D) Bacteriome structure
observed in the progeny of ampicillin receiving females (C) and
tetracycline and yeast extract receiving females in (D). In these
individuals, the bacteriocyctes lack Wigglesworthia and these females
are reproductively sterile, images taken at 10x magnification.
The effect of antibiotics on the bacteriome of G. m.
Symbiont PCR primers.
crosses. In three separate experiments % larval deposition and %
eclosion of the pupa deposited was determined. For each experiment,
number of larval deposited for surviving females over two
gonotrophic cycles and number of their pupae that hatched were
recorded. Larval deposition was used as a measure of CI expression.
To analyze for CI in replicate experiments of individual crosses,
to comparelarval deposition rates.Wilcoxon tests, with a Bonferrroni
correction were also conducted to compare pupal eclosion. Super-
scripted letters indicate significant differences, P,0.01.
Larval deposition and pupal eclosion data for single cage
We thank Kostas Bourtzis for the WSP antibody. We thank Jeffrey
Townsend for statistical advice. We are also grateful to FAO/IAEA
Coordinated Research Program on "Improving SIT for Tsetse Flies
through Research on their Symbionts and Pathogens’’ and to Slovak
Academy of Science, Bratislava, Slovakia for providing puparia for our
Conceived and designed the experiments: SA AG UA JM CB AH PT.
Performed the experiments: UA JM CB RP CL JC SB. Analyzed the data:
UA JM CB SA AG SB PT. Contributed reagents/materials/analysis tools:
AH PT. Wrote the paper: SA UA CB JM.
Wolbachia Induces CI in Tsetse
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1. Simarro PP, Jannin J, Cattand P (2008) Eliminating human African
trypanosomiasis: where do we stand and what comes next? PLoS Med 5: e55.
2. Cecchi G, Paone M, Franco J, Fevre E, Diarra A, et al. (2009) Towards the atlas
of human African trypanosomiasis. Int J Health Geogr 8: 15.
3. Simarro P, Diarra A, Ruiz Postigo J, Franco J, Jannin J (2011) The human
African Trypanosomiasis control and surveillance programme of the World
Health Organization 2000-2009. PLoS Negl Trop Dis 5: e1007.
4. Leak SG, Peregrine AS, Mulatu W, Rowlands GJ, D’Ieteren G (1996) Use of
insecticide-impregnated targets for the control of tsetse flies (Glossina spp.) and
trypanosomiasis occurring in cattle in an area of south-west Ethiopia with a high
prevalence of drug-resistant trypanosomes. Trop Med Int Health 1: 599–609.
5. Davis S, Aksoy S, Galvani A (2010) A global sensitivity analysis for African
sleeping sickness. Parasitology. pp 1–11.
6. Aksoy S, Weiss B, Attardo G (2008) Paratransgenesis applied for control of tsetse
transmitted sleeping sickness. Adv Exp Med Biol 627: 35–48.
7. Chen XA, Aksoy S (1999) Tissue tropism, transmission and expression of foreign
genes in vivo in midgut symbionts of tsetse flies. Insect Mol Biol 8: 125–132.
8. Weiss BL, Mouchotte R, Rio RV, Wu YN, Wu Z, et al. (2006) Interspecific
transfer of bacterial endosymbionts between tsetse fly species: infection
establishment and effect on host fitness. Appl Environ Microbiol 72: 7013–7021.
9. Welburn SC, Maudlin I, Ellis DS (1987) In vitro cultivation of rickettsia-like-
organisms from Glossina spp. Ann Trop Med Parasitol 81: 331–335.
10. Rio RV, Hu Y, Aksoy S (2004) Strategies of the home-team: symbioses exploited
for vector-borne disease control. Trends Microbiol 12: 325–336.
11. Vreysen MJ, Saleh KM, Ali MY, Abdulla AM, Zhu Z, et al. (2000) Glossina
austeni (Diptera: Glossinidae) eradicated on the Island of Unguga, Zanzibar,
using the sterile insect technique. J Econ Entomol 93: 123–135.
12. Aksoy S (1995) Wigglesworthia gen. nov. and Wigglesworthia glossinidia sp. nov., taxa
consisting of the mycetocyte-associated, primary endosymbionts of tsetse flies.
Int J Syst Bacteriol 45: 848–851.
13. Aksoy S (2000) Tsetse - A haven for microorganisms. Parasitol Today 16:
14. Attardo GM, Lohs C, Heddi A, Alam UH, Yildirim S, et al. (2008) Analysis of
milk gland structure and function in Glossina morsitans: Milk protein production,
symbiont populations and fecundity. J Insect Physiol 54: 1236–1242.
15. Nogge G (1976) Sterility in tsetse flies (Glossinia morsitans Westwood) caused by loss
of symbionts. Experientia 32: 995–996.
16. Nogge G (1978) Apos-Symbiotic tsetse flies, Glossina-Morsitans-Morsitans obtained
by feeding on rabbits immunized specifically with symbionts. J Insect Physiol 24:
17. Nogge G (1980) Elimination of symbionts of tsetse flies (Glossina m. morsitans
Westw.) by help of specific antibodies. In: Schwemmler W, Schenk H, eds.
Endocytobiology. Berlin: W. de Gruyter. pp 445–452.
18. Nogge G, Gerresheim A (1982) Experiments on the elimination of symbionts
from the tsetse-Fly, Glossina-Morsitans-Morsitans (Diptera, Glossinidae), by antibi-
otics and lysozyme. J Invertebr Pathol 40: 166–179.
19. O’Neill SL, Gooding RH, Aksoy S (1993) Phylogenetically distant symbiotic
microorganisms reside in Glossina midgut and ovary tissues. Med Vet Entomol 7:
20. Cheng Q, Ruel TD, Zhou W, Moloo SK, Majiwa P, et al. (2000) Tissue
distribution and prevalence of Wolbachia infections in tsetse flies, Glossina spp.
Med Vet Entomol 14: 44–50.
21. Pais R, Lohs C, Wu Y, Wang J, Aksoy S (2008) The obligate mutualist
Wigglesworthia glossinidia influences reproduction, digestion, and immunity
processes of its host, the tsetse fly. Appl Environ Microbiol 74: 5965–5974.
22. Werren JH (1997) Biology of Wolbachia. Annu Rev Entomol 42: 587–609.
23. Werren JH, Baldo L, Clark ME (2008) Wolbachia: master manipulators of
invertebrate biology. Nat Rev Microbiol 6: 741–751.
24. Saridaki A, Bourtzis K (2010) Wolbachia: more than just a bug in insects genitals.
Curr Opin Microbiol 13: 67–72.
25. Dobson SL, Fox C, Jiggins FM (2002) The effect of Wolbachia-induced
cytoplasmic incompatibility on host population size in natural and manipulated
systems. Proc Biol Sci 269: 437–445.
26. Hoffman AA, Hercus M, Dagher H (1998) Population Dynamics of the
Wolbachia infection causing cytoplasmic incompatibility in Drosophila melanogaster.
Genetics 148: 221–231.
27. Sinkins SP, Gould F (2006) Gene drive systems for insect disease vectors. Nat
Rev Genet 7: 427–435.
28. Rasgon J (2007) Population replacement strategies for controlling vector
populations and the use of Wolbachia pipientis for genetic drive. J Vis Exp. 225 p.
29. Brelsfoard CL, Dobson SL (2009) Wolbachia-based strategies to control insect
pests and disease vectors. Asia Pac. J. Mol. Biol. Biotechnol 17: 55–63.
30. Rasgon JL (2008) Using predictive models to optimize Wolbachia-based strategies
for vector-borne disease control. Adv Exp Med Biol 627: 114–125.
31. Van Meer MMM, Witteveldt J, Stouthamer R (1999) Phylogeny of the
arthropod endosymbiont Wolbachia based on the wsp gene. Insect Mol Biol 8:
32. Nogge G (1981) Significance of symbionts for the maintenance of an optimal
nutritional state for successful reproduction in hematophagous arthropods.
Parasitology 82: 101–104.
33. Ballard JWO, Melvin RG (2007) Tetracycline treatment influences mitochon-
drial metabolism and mtDNA density two generations after treatment in
Drosophila. Insect Mol Biol 16: 799–802.
34. Dobson SL, Rattanadechakul W, Marsland EJ (2004) Fitness advantage and
cytoplasmic incompatibility in Wolbachia single- and superinfected Aedes albopictus.
Heredity: 93: 135–142.
35. Dean M (2006) A Wolbachia-associated fitness benefit depends on genetic
backgroun in Drosophila simulans. Proc Biol Sci 273: 1415–1420.
36. Clark ME, Veneti Z, Bourtzis K, Karr TL (2002) The distribution and
proliferation of the intracellular bacteria Wolbachia during spermatogenesis in
Drosophila. Mech Dev 111: 3–15.
37. Clark ME, Veneti Z, Bourtzis K, Karr TL (2003) Wolbachia distribution and
cytoplasmic incompatibility during sperm development: the cyst as the basic
cellular unit of CI expression. Mech Dev 120: 185–198.
38. Clark ME, Bailey-Jourdain C, Ferree PM, England SJ, Sullivan W, et al. (2008)
Wolbachia modification of sperm does not always require residence within
developing sperm. Heredity 101: 420–428.
39. Veneti Z, Clark ME, Zabalou S, Karr T, Savakis C, et al. (2003) Cytoplasmic
incompatibility and sperm cyst infection in different Drosophila-Wolbachia
association. Genetics 164: 545–552.
40. Ferree PM, Frydman HM, Li JM, Cao J, Wieschaus E, et al. (2005) Wolbachia
utilizes host microtubules and dynein for anterior localization in the Drosophila
oocyte. PloS Pathog 1: 111–124(e114).
41. Serbus LR, Sullivan W (2007) A cellular basis for Wolbachia recruitment to the
host germline. PloS Pathog 3: e190.
42. Veneti Z, Clark ME, Karr TL, Savakis C, Bourtzis K (2004) Heads or tails: host-
parasite interactions in the Drosophila-Wolbachia system. Appl Environ Microbiol
43. Tram U, Fredrick K, Werren JH, Sullivan W (2006) Paternal chromosomal
segregation during the frist mitotic division determines Wolbachia-induced
cytoplasmic incompatibility phenotype. J Cell Sci 119: 3655–3663.
44. Xi Z, Khoo CC, Dobson SL (2006) Interspecific transfer of Wolbachia into the
mosquito disease vector Aedes albopictus. Proc Biol Sci 273: 1317–1322.
45. Charlat S, Hurst GD, Mercot H (2003) Evolutionary consequences of Wolbachia
infections. Trends Genet 19: 217–223.
46. Dobson SL (2004) Evolution of Wolbachia cytoplasmic incompatibility types.
Evolution 58: 2156–2166.
47. Doudoumis V, Tsiamis G, Wamwiri F, Brelsfoard C, Alam U, et al. (in press)
Detection and characterization of Wolbachia infections in laboratory and natural
populations of different species of tsetse (genus Glossina). BMC Microbiol.
48. Hoffmann AA, Turelli M, Harshman LG (1990) Factors affecting the
distribution of cytoplasmic incompatibility in Drosophila simulans. Genetics 126:
49. Hoffmann A, Hercus M, Dagher H (1998) Population dynamics of the Wolbachia
infection causing cytoplasmic incompatibility in Drosophila melanogaster. Genetics
50. Weeks A, Turelli M, Harcombe W, Reynolds K, Hoffman AA (2007) From
parasite to mutualist: rapid evolution of Wolbachia in natural populations of
Drosophila. PLoS Biol 5: e114.
51. Beard CB, O’Neill SL, Mason P, Mandelco L, Woese CR, et al. (1993) Genetic
transformation and phylogeny of bacterial symbionts from tsetse. Insect Mol Biol
52. Cheng Q, Aksoy S (1999) Tissue tropism, transmission and expression of foreign
genes in vivo in midgut symbionts of tsetse flies. Insect Mol Biol 8: 125–132.
53. Hu YJ, Aksoy S (2005) An antimicrobial peptide with trypanocidal activity
characterized from Glossina morsitans morsitans. Insect Biochem Mol Biol 35:
54. Rio RV, Wu YN, Filardo G, Aksoy S (2006) Dynamics of multiple symbiont
density regulation during host development: tsetse fly and its microbial flora.
Proc Biol Sci 273: 805–814.
55. Brelsfoard CL, Sechan Y, Dobson SL (2008) Interspecific hybridization yields
strategy for South Pacific filariasis vector elimination. PLoS Negl Trop Dis 2:
56. Zabalou S, Riegler M, Theodorakopoulou M, Stauffer C, Savakis C, et al. (2004)
Wolbachia-induced cytoplasmic incompatibility as a means for insect pest
population control. Proc Natl Acad Sci U S A 101: 15042–15045.
57. Laven H (1967) Eradication of Culex pipiens fatigans through cytoplasmic
incompatibility. Nature 216: 383–384.
58. Zabalou S, Apostolaki A, Livadaras I, Franz G, Robinson A, et al. (2009)
Incompatible insect technique: incompatible males from a Ceratitis capitata
(Diptera: Tephritidae) gentic sexing strain. Entomol Exp Appl 132: 232–240.
59. Vreysen M, Saleh K, Lancelot R, Bouyer J (2011) Factory Tsetse flies must
behave like wild flies: A prerequisite for the sterile insect technique. PLoS Negl
Trop Dis 5: e907.
60. McMeniman CJ, Lane RV, Cass BN, Fong AW, Sidhu M, et al. (2009) Stable
introduction of a life-shortening Wolbachia infection into the mosquito Aedes
aegypti. Science 323: 141–144.
61. Xi Z, Khoo CCH, Dobson SL (2005) Wolbachia establishment and invasion in an
Aedes aegypti laboratory population. Science 310: 327–310.
62. Frydman HM, Li JM, Robson DN, Wieschaus E (2006) Somatic stem cell niche
tropism in Wolbachia. Nature 441: 509–512.
Wolbachia Induces CI in Tsetse
PLoS Pathogens | www.plospathogens.org10 December 2011 | Volume 7 | Issue 12 | e1002415
63. Ruang-Areerate T, Kittayapong P (2006) Wolbachia transinfection in Aedes aegypti: Download full-text
a potential gene driver of dengue vectors. Proc Natl Acad Sci U S A 103:
64. Turelli M, Hoffman A (1999) Microbe induced cytoplasmic incompatibility as a
mechanism for introducing genes into arthropod populations. Insect Mol Biol 8:
65. Osborne SE, Leong YS, O’Neill SL, Johnson KN (2009) Variation in antiviral
protection mediated by different Wolbachia strains in Drosophila simulans. PLoS
Pathog 5: e1000656.
66. Kambris Z, Blagborough A, Pinto S, Blagrove M, Godfray H, et al. (2010)
Wolbachia stimulates immune gene expression and inhibits plasmodium develop-
ment in Anopheles gambiae. PLoS Pathog 6: e1001143.
67. Kambris Z, Cook PE, Phuc HK, Sinkins SP (2009) Immune activation by life-
shortening Wolbachia and reduced filarial competence in mosquitoes. Science
68. Moreira LA, Iturbe-Ormaetxe I, Jeffery JA, Lu G, Pyke AT, et al. (2009) A
Wolbachia symbiont in Aedes aegypti limits infection with dengue, Chikungunya
and Plasmodium. Cell 139: 1268–1278.
69. Bian G, Xu Y, Lu P, Xie Y, Xi Z (2010) The endosymbiotic bacterium Wolbachia
induces resistance to dengue virus in Aedes aegypti. PLoS Pathog 6: e1000833.
70. Teixeira L, Ferreira A, Ashburner M (2008) The bacterial symbiont Wolbachia
induces resistance to RNA viral infections in Drosophila melanogaster. PLoS Biol 6:
71. Moloo SK (1971) An artificial feeding technique for Glossina. Parasitology 63:
72. Anselme C, Vallier A, Balmand S, Fauvarque MO, Heddi A (2006) Host PGRP
gene expression and bacterial release in endosymbiosis of the weevil Sitophilus
zeamais. Appl Environ Microbiol 72: 6766–6772.
Wolbachia Induces CI in Tsetse
PLoS Pathogens | www.plospathogens.org11 December 2011 | Volume 7 | Issue 12 | e1002415