1 uninfected mosquito bites confer protection against...
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Uninfected Mosquito Bites Confer Protection Against Infection 1
with Malaria Parasites. 2
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RUNNING TITLE: Mosquito Bites Confer Protection Against Plasmodium 4
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Michael J. Donovan1, Andrew S. Messmore2, Deborah A. Scrafford1, David 6
L. Sacks2, Shaden Kamhawi2, and Mary Ann McDowell1* 7
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1Center for Global Health and Infectious Diseases, Department of Biological Sciences, 9
University of Notre Dame, Notre Dame, IN 46556 10
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2Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, 12
National Institutes of Health, Bethesda, Maryland 20892 13
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* Corresponding author. Mailing address: 215 Galvin Life Sciences, Department 21
of Biological Science, University of Notre Dame, Notre Dame, IN 46656. Phone: 22
(574) 631-9771. Fax: (574) 631-7413. E-mail: [email protected] 23
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Copyright © 2007, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.Infect. Immun. doi:10.1128/IAI.01928-06 IAI Accepts, published online ahead of print on 5 March 2007
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Abstract 24
Despite decades of research and multiple initiatives, malaria continues to be one 25
of the world’s most debilitating infectious diseases. New insights for malaria 26
control and vaccine development will be essential to thwart the staggering 27
worldwide impact of this disease (7); ultimately successful vaccine strategies will 28
undoubtedly be multi-factorial, incorporating multiple antigens and targeting 29
diverse aspects of the parasite’s biology (23). Using a murine model of malaria 30
infection we show here that exposure to uninfected mosquito bites prior to 31
Plasmodium yoelii infection influences the local and systemic immune response 32
and limits parasite development within the host. In hosts pre-exposed to 33
uninfected mosquito bites reduced parasite burdens were detected early in the 34
liver and remained lower during the blood stage of the life-cycle as compared to 35
hosts that only received mosquito bites at the time of infection. Repeated 36
exposure to uninfected mosquito bites skewed the immune response towards a 37
T-helper 1 (Th1) phenotype as indicated by increased levels of interleukin-12 (IL-38
12), interferon-gamma (IFN-g) and inducible nitric oxide synthase (iNOS). These 39
data suggest that the addition of mosquito salivary components to anti-malaria 40
vaccines may be a viable strategy for creating a Th1 biased environment known 41
to be effective against malaria infection. Furthermore, this strategy may be 42
important for the development of vaccines to combat other mosquito-transmitted 43
pathogens. 44
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Introduction 47
Malaria continues to be a major public health threat and has an enormous 48
economic impact, resulting in nearly 3 million deaths annually and ranking eighth 49
as a contributor to loss of global disability adjusted life years (7). Emerging drug 50
resistance in the Plasmodium parasites that cause malaria and insecticide 51
resistance in mosquito vectors that transmit these pathogens emphasizes the 52
urgent need for developing an effective malaria vaccine to control the devastating 53
burden of malarial disease. The complex biology of malaria parasites coupled 54
with antigenic polymorphism, poor antigen immunogenecity, and parasite-55
induced immuno-suppression distinguishes the quest for a malaria vaccine as 56
extraordinarily daunting. 57
58
Malaria is transmitted to humans via the bite of its insect vector, a female 59
anopheline mosquito. During blood feeding mosquitoes inject infective 60
Plasmodium sporozoites into the avascular skin tissue of its host where they 61
eventually migrate into the circulation (53); simultaneously, a plethora of 62
pharmacologically active compounds in mosquito saliva are introduced into the 63
host. These compounds have substantial anti-haemostatic, anti-inflammatory, 64
and immunosuppressive activities that aid the mosquito in the blood feeding 65
process (40). Furthermore, many of the salivary components are immunogenic 66
and elicit strong immune responses, evidenced by the swelling and itching that 67
accompanies a mosquito bite (38). This substantial effect of immune activation 68
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by saliva creates an inflammatory context for further responses to co-injected 69
pathogens. 70
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A role for arthropod saliva in modifying the outcome of infection is not novel to 72
mosquitoes and malaria parasites; increased pathogen infectivity has been 73
described for ticks, sand flies and mosquitoes [for review see (50)]. While 74
studies primarily have focused on the enhancement of transmission and disease 75
when pathogens are introduced in the presence of vector saliva, some studies 76
have explored the effect of repeated exposure of vector saliva on the outcome of 77
infection. Although the mechanism has yet to be completely elucidated, repeated 78
infestation with pathogen-free Ixodes scapularis ticks induces resistance to 79
Borrelia burgdorferi transmission (56). The most striking host-parasite-vector 80
system that has been studied is Leishmania infection by the bites of 81
phlebotomine sand flies. Interestingly, multiple exposures to uninfected sand fly 82
bites prior to infection confers resistance to L. major, due to increases in the 83
cytokines responsible for cell-mediated immunity (24). Mosquito bites also have 84
been shown to influence immunity and potentiate viral disease in mouse models 85
(18, 26, 45, 46), possibly through modulation of host systemic cytokine 86
responses (46, 58). 87
88
Mosquito bites induce immediate, delayed and systemic hypersensitivity 89
reactions in hosts (38); consequently, we hypothesized that the local tissue and 90
systemic environment when ‘immunized’ by mosquito salivary components can 91
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enhance malaria immunity. We tested this hypothesis using the prototypic 92
murine model of malaria infection: sporozoite infection in mice via P. yoelii 93
infected Anopheles stephensi mosquitoes. 94
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Materials and Methods 96
Mice Balb/c mice aged 6-8 weeks were bred at the Friemann Life Sciences 97
Center at the University of Notre Dame under approved Institutional Animal Care 98
and Use Committee (IACUC) protocols. All mice were female and age-matched 99
for all experiments. Interferon gamma deficient (IFN-g KO) and Balb/c wild type 100
(WT) counterparts were purchased (Jackson Labs, Bar Harbor, ME) and used at 101
6-8 weeks of age. 102
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Pre-sensitization Mice were anesthetized, and their ears were exposed for 20 104
min. to a screened vial containing 15-20 fully-matured female adults every two 105
weeks for six weeks (pre-sensitized). A control group of age-matched mice were 106
only anesthetized at each time point (naïve). Mice were challenged 2 weeks 107
following the last pre-exposure. Twenty-four (cytokine responses) or 40 108
(parasite quantification) hours after the final exposure, ears, liver, and spleen 109
samples were harvested and stored in RNAlater (Qiagen, Valencia, CA). 110
111
Mosquito Infections P. yoelii (17XNL) parasites were maintained by alternating 112
passage of parasites through A. stephensi and Balb/c mice. Murine parasitemia 113
was observed through thin-layered blood smears. Smears were fixed in 100% 114
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methanol and stained with Geimsa. Once gametocytes were present, infected 115
animals were anesthetized and A. stephensi were allowed to feed. Four to eight 116
days after infection, A. stephensi were anesthetized and midguts were dissected 117
out, and stained with mercurochrome (Fisher Scientific, Chicago, IL), and the 118
number of oocysts per midgut were counted. Average infection rates were 119
between 75 and 100 percent. After the parasite matured to the salivary gland-120
sporozoite stage (14 days post-infection), appropriate groups were exposed to P. 121
yoelii-infected A. stephensi. Mice were exposed to 10 infected mosquitoes for 15 122
minutes on the right ear, the same vial of mosquitoes was transferred to the left 123
ear for an additional 15 minutes. Feeding success was assessed by visual 124
observation, looking for blood in the midgut; no obvious differences between 125
feeding on naïve versus pre-sensitized mice were detected. Sporozoites were 126
isolated as previously described (17). 127
128
Blood Stage Quantification Mice were pre-sensitized as described above. 129
Two weeks following 3 exposures to uninfected mosquitoes, mice were exposed 130
to bites of P. yoelii-infected A. stephensi. 24 hours post-exposure, blood 131
samples were taken, thin-layer blood smears were made, and were stained with 132
Geimsa. Subsequently, samples were taken and quantified each day until 133
mouse euthanasia on day 7 post-infection. For determination of parasitemia, 134
1000 cells were counted from each sample. 135
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RNA Isolation and Generation of cDNA RNA was isolated using an RNeasy 137
Mini kit (Qiagen, Valencia, CA) for ears and Trizol Reagent (Invitrogen, Carlsbad, 138
CA) for spleens and livers. The entire harvested organ was homogenized in the 139
respective lysis buffers. Contaminating DNA was removed from 1µg of RNA via 140
DNase I treatment (Invitrogen, Carlsbad, CA), using 1 Unit of DNase I, and a final 141
concentration of DNAse buffer containing 20mM Tris-HCl (ph 8.4), 2mM MgCl, 142
and 50mM KCl. DNA-free RNA was used to generate cDNA with oligo dT (for 143
cytokine analysis) (Invitrogen, Carlsbad, CA) or Random Primers (for infection 144
studies) (Invitrogen, Carlsbad, CA) at a final concentration of 10ng and 15ng, 145
respectively, in addition to 500nM dNTPs (Invitrogen, Carlsbad, CA), 200 units of 146
Superscript III Reverse Transcriptase (Invitrogen, Carlsbad, CA), 40 units RNase 147
Out (Invitrogen, Carlsbad, CA), and 5mM DTT. 148
149
Quantitative PCR and Analysis cDNA was used for quantitative real time PCR 150
analysis using the 2x SYBR Green Kit (Applied Biosystems, Foster City, CA). 151
Reactions were run on the ABI 7700 Sequence Detector machine. The SYBR 152
Green kit was used at a 1x proportion, along with 300 nM of forward and reverse 153
primers for each reaction. Primers used (IDT Coralville, IA) were HPRT 5’-GTT 154
GGA AGG CCA GAC TTT GTT-3’ and 5’-GAT TCA ACT TGC GCT CAT CTT 155
AGG C-3’; IFN-g 5’-AGA GCC AGA TTA TCT CTT TCT ACC TCA-3’ and 5’-CCT 156
TTT TCG CCT TGC TGT TG-3’; IL-4 5’-ACG AGG TCA CAG GAG AAG GA-3’ 157
and 5’-AGC CCT ACA GAC GAG CTC ACT C-3; IL-12p40 5’ AACCAT CTC CTG 158
GTT TGC CA-3’and 5’-CGG GAG TCC AGT CCA CCT C-3’;and iNOS primers 159
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as previously described (39). Cytokine primers were designed to overlap introns 160
to help assure no DNA amplification occurred. In order to be assured of proper 161
amplification, a melting curve analysis was performed on each product. We 162
employed the comparative threshold cycle method to determine relative 163
differences in parasite burdens. After generation of Ct values (the cycle number 164
at which the reaction crosses the threshold), relative copy number was 165
determined according to the following equation: number of copies = 2-∆∆ct, 166
where ∆∆ct = ∆ct(sample) – ∆ct(calibrator), ∆ct = ∆ct(sample) – ∆ct(HPRT), ct = 167
cycle at which a statistically significant increase in the emission intensity over the 168
background, and ∆ct(calibrator) = the mean ∆ct for the naïve control. Parasite 169
liver loads were determined using primers for P. yoelii 18s rRNA 40 hours after 170
infection, as previously described (9). The amount of RNA of the different 171
samples was normalized based on the measurement of the mRNA levels from 172
the mouse housekeeping gene HPRT as described above. Cytokine graphs are 173
expressed as the mean value of induction levels that have been normalized to 174
the mean values from naïve tissues. Infection levels were expressed in 40-∆∆ct 175
as previously described (51). Significance levels were determined using a 176
Student’s T Test with a confidence level of 95%. 177
178
ELISA analysis Serum was collected from each mouse at the time of organ 179
harvest and IFN-g was quantified by ELISA (Pierce Biotechnology, Rockford, IL) 180
according to the manufacturer’s instruction. 181
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Results and Discussion 183
In some areas a single individual can receive nearly 200 mosquito bites/day (52) 184
or over 10,000 bites/year (22). Malaria infection rates in these vectors range 185
from below 0.1% to 10% (22, 28), indicating that even in the highest areas of 186
transmission a single individual is exposed to drastically more mosquito saliva 187
than malaria parasites. To assess the impact of previous exposure to uninfected 188
mosquito bites on Plasmodium development, we compared P. yoelii burdens in 189
mice pre-exposed to uninfected A. stephensi bites (pre-sensitized) to those of 190
unexposed (naïve) mice. We utilized a system where infected and uninfected 191
mosquito bites were limited to the ears so that local and systemic responses 192
could be separated easily. Forty hours post-infection, after parasite 193
differentiation and amplification but prior to parasite release into the circulation, 194
pre-sensitized mice exhibited significantly reduced liver burdens of P. yoelii 195
compared to naïve mice (Fig. 1a), corresponding to a 9-fold reduction in parasite 196
numbers following the natural infection (Fig. 1b). These reduced parasite 197
burdens also were evident when blood parasitemias were assessed (Fig. 1c). 198
While the pre-patent period (i.e. the time it takes to visually detect blood 199
parasites) was not effected by pre-sensitization, the parasitemia levels were 200
reproducibly lower in mice pre-exposed to uninfected mosquito bites. 201
202
The hepatic stage is the most vulnerable of the Plasmodium life-cycle for 203
intervention and substantial research on liver stage immunity, primarily using 204
murine models, exists. The predominant effector mechanism mediating this pre-205
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erythrocytic immunity is the production of IFN-g that inhibits parasite 206
development within hepatocytes via nitric oxide (16). To investigate the effect 207
that repeated exposure of mosquito saliva has on cytokine profiles, both local 208
tissue and systemic IFN-g and interleukin-4 (IL-4) mRNA levels were assessed. 209
Local IFN-g expression was relatively low in naïve animals and mice that were 210
only exposed a single time (naïve bit), while pre-sensitized animals readily 211
produced IFN-g in response to A. stephensi bites (Fig. 2a). Furthermore, pre-212
sensitized mice express reduced levels of IL-4 as compared to naive animals 213
(Fig. 2b). The pre-sensitized mice are Th1 biased, illustrated by cytokine ratio 214
(IFN-g:IL-4) with much more IFN-g being expressed than IL-4 (Fig. 2c). 215
Therefore, A. stephensi saliva significantly changes the local cytokine 216
environment at the tissue site where parasites are introduced; Th1 bias following 217
mosquito pre-sensitization also occurs systemically in the liver (Fig. 2d-f), spleen 218
(Fig. 2g-i), and serum (data not shown), indicating a systemic cytokine shift to a 219
Th1 profile. These results suggest that up-regulation of IFN-g is part of the 220
protective phenotype against P. yoelii infection associated with mosquito saliva 221
pre-sensitization. 222
223
To further evaluate the role of IFN-g, WT and IFN-g KO BALB/c mice were pre-224
sensitized to uninfected mosquito bites, or left naïve, prior to natural infection 225
with P. yoelii. Pre-sensitization-associated protection against P. yoelii infection 226
was abrogated in the absence of IFN-g (Fig. 3a), indicating that IFN-g is essential 227
for the protective response. Because nitric oxide is required for immunity to P. 228
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yoelii liver infection (5), we measured inducible nitric oxide synthase (iNOS) 229
mRNA levels in the livers of naïve and pre-sensitized WT and IFN-g KO mice 230
(Fig. 3b). The 10-fold up-regulation of iNOS levels in response to A. stephensi 231
pre-exposure was not observed in IFN-g KO mice, suggesting that the IFN-g 232
induced following pre-sensitization to mosquito bites leads to NO-induced killing 233
of malarial parasites. Interestingly iNOS mRNA was also increased by pre-234
sensitization in the local ear environment as early as 5 hours post infection (Fig. 235
4). 236
237
As interleukin-12 (IL-12) is necessary for immunity against P. yoelii (17) and has 238
been proposed as a potential adjuvant for anti-malaria vaccines (44, 47), we 239
evaluated IL-12p40 mRNA levels in the livers (Fig. 5a) and spleens (Fig. 5b) of 240
pre-sensitized animals. We detected significantly higher levels of IL-12p40 241
mRNA in the organs of pre-sensitized animals as compared to naïve mice. 242
243
We show that the immunity induced following A. stephensi pre-sensitization 244
involves local and systemic up-regulation of IFN-g and iNOS (Figs. 2-4, data not 245
shown). These results raise the question of what tissue is primarily affected by 246
this protective mechanism and the timing of this response. The majority of 247
infectious malaria sporozoites released during mosquito blood feeding do not 248
immediately enter the circulation, instead sporozoites are deposited into the skin 249
where they eventually move into dermal vessels (53). Although the timing of 250
vascular entry remains a matter of debate, it is clear that some sporozoites 251
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remain in the skin and may take substantially longer to enter the blood stream 252
(53). This delayed timing coupled with the suggestion that sporozoites are 253
directly susceptible to NO-mediated killing (29) invokes a model of reduction in P. 254
yoelii burdens in the liver due to lower numbers of parasites entering the 255
circulation. To determine the chronology and location of parasite killing in pre-256
sensitized mice, we evaluated the relative parasite burdens in both the ear and 257
the liver at various time points (5-30 hours) following natural infection in naïve 258
and pre-sensitized mice (Fig. 6a & b). By 10 hours post infection, parasite 259
quantification in the ear returned to background levels in both groups of mice, 260
indicating that the same numbers of sporozoites leave the bite site (Fig. 6a) and 261
enter the liver (Fig. 6b) in naïve and pre-sensitized mice. The difference in liver 262
parasite burdens is not evident until 20 hours post-infection. As increases in 263
parasite burden during the first 5 – 40 hours of infection is primarily due to 264
parasite multiplication our data suggests that the protective mechanism due to 265
pre-sensitization is operating in the liver. To conclusively address whether liver 266
protection could be occurring, we exposed naïve and pre-sensitized mice to 267
uninfected mosquito bites and immediately challenged intravenously in the tail 268
vein, rather than exposing mice to infected mosquitoes. Forty hours post-i.v. 269
challenge, P. yoelii liver burdens remained significantly lower in pre-sensitized 270
mice, demonstrating that the protection is due to, in part, the response in the liver 271
(Fig. 6c). 272
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The substantial effort towards the production of an efficacious malaria vaccine 274
has yet to yield a suitable product. Following the initial report that vaccination 275
with radiation-attenuated sporozoites can protect against malaria challenge (36), 276
extensive research has elucidated the immunological parameters that confer 277
such protection. The protective mechanisms identified for malaria sporozoite and 278
liver stages are remarkably similar in rodent models and human volunteers (30). 279
It is possible that residual salivary components remaining from the isolation of 280
sporozoites can partially explain the protection observed in the studies involving 281
radiation-attenuated sporozoites. One original study investigating irradiated 282
sporozoites as a vaccine used repeated vaccination with large amounts of 283
mosquito salivary gland homogenate (70 glands) as a control and demonstrated 284
that this procedure conferred partial protection to P. berghei infection in mice (1, 285
2). More recently it has been demonstrated that P. gallinaceum parasitemias are 286
increased in the presence of Aedes fluviatilis saliva in a chicken malaria model; a 287
response that was reversed with prior exposure to mosquito saliva (43). 288
289
It is well established that naïve travelers and children are at increased risk for 290
severe malarial disease as compared to adult endemic populations and that 291
these adults do not exhibit sterile immunity; rather, this degree of natural 292
immunity results in asymptomatic infections with lower parasite burdens in the 293
circulation (42). Historically this phenomenon has been attributed to the gradual 294
onset of immunity due to repeated parasite infections (48). We propose that the 295
extensive exposure to mosquito saliva that accrues over time in malaria endemic 296
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regions results in a Th1 climate that influences pathogen establishment. 297
Coupled with specific immunity to malaria parasites this inhospitable environment 298
may contribute to lower malarial burdens in endemic adults. Here we utilize a 299
model whereby we infect with 10 mosquitoes with infection rates ranging from 300
75-90% with oocyst counts of 24-39 per mosquito to ensure adequate parasite 301
numbers for detection. The majority of mosquitoes in the field harbor only one 302
oocyst (12) with infection rates never greater than 10% (22, 28). We did not 303
observe sterile immunity in our model system; however, it is possible that the 304
cytokine shift that we detected may be adequate to control the small numbers of 305
parasites an individual encounters in a natural setting. 306
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The IFN-g up-regulation due to pre-sensitization that we detected diminishes 308
over time as is evident by the lower levels of IFN-g mRNA that are present in 309
ears and liver in pre-sensitized mice that did not receive one final bite prior to 310
analysis. This observation is consistent with the typical rapid loss of anti-malarial 311
immunity when an individual leaves an endemic region (48). We suggest that 312
this loss of immunity can partially be explained by an absence, or drastically 313
lower level, of mosquito boosting. In our study mosquito exposure ceased after 314
the infection was initiated. It is intriguing to postulate that increased resistance 315
may be observed if hosts continued to be exposed to mosquito bites throughout 316
the infection as would be the case in an endemic region. IFN-g production is 317
associated with primary immune responses to blood stage Plasmodium 318
infections in mice (13, 34) and humans (31) and appears to be crucial for the 319
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development of protective immunity (15, 49). Although in this study we did not 320
explore blood stage immunity, the increased levels of IFN-g in the serum induced 321
by repeated mosquito exposures suggest that blood stage immunity may be 322
influenced and continued mosquito boosting may increase this response. 323
324
The systemic shift in cytokine balance that we detected is not unique as bite-325
induced systemic responses have been documented with Rhipicephalus 326
sanguineus (tick) (19), Culex pipiens and Aedes aegypti (58). The majority of 327
studies assessing arthropod modulation of host immune responses have focused 328
on one-time exposures to bites or salivary gland components [for review see (6, 329
8, 50)]. Combined these studies engender a model where initial exposure to 330
arthropod saliva induces a Th2 immune response, potentiating infectivity of a 331
variety of vector-borne pathogens. For mosquito transmitted pathogens, 332
infectivity of P. berghei (54), Cache Valley virus (18), La Crosse virus (37), and 333
vesicular stomatitis virus (VSV) (26, 27) is enhanced in the presence of saliva. 334
For VSV infection, increased viral loads are associated with a SGH dependent 335
decrease of type I interferons in vitro (26). Feeding of both Culex pipiens and 336
Aedes aegypti mosquitoes on mice induces increased levels of systemic IL-4 and 337
IL-10 with a concomitant decrease in IFN-g production (58). Similarly, 338
inoculation of Ae. Aegypti SGH with Sindbis virus results in higher levels of Th2 339
cytokines and reduced expression of interferons (46). 340
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While we detected increased Th2 responses in naïve mice versus pre-sensitized 342
hosts, we did not observe an increase in IL-4 upon one exposure to A. stephensi. 343
Our results are in agreement with previous studies investigating skin and 344
lymphnode cytokine production in response to A. stephensi bites (14). In 345
contrast to our observations, studies exploring splenic cytokine production in 346
response to exposure to Ae. Aegypti or C. pipiens mosquitoes detected an 347
increase in Th2 cytokine production in C3H/HeJ mice; however, in a congenic 348
host strain (C3H/RV) Th1 cytokines predominated (58). In vitro studies suggest 349
that mosquito species may differ in their ability to modulate host immune 350
responses (55); the discrepancies between our results and previous work 351
investigating in vivo immune modulation by mosquito exposure may be attributed 352
to variation in the experimental approaches or may reflect actual differences 353
between mosquito species. 354
355
Data exploring the effect of multiple, repeated exposures to arthropod saliva are 356
scant. While single exposures to sand fly bites (24) or SGH (4, 32, 35) are 357
associated with increased levels of Th2 cytokines, repeated exposure leads to a 358
switch to Th1 immunity (4, 24). This switch to primarily IFN-g production at the 359
bite site induces resistance to L. major infection. In contrast, repeated 360
exposures to Ae. aegypti bites results in elevated production of antigen-specific 361
IL-4 production in cultured spleen cells, although this response is not detected in 362
response to ConA stimulation (10). Multiple infestations of ticks are generally 363
thought to lead to heightened levels of Th2 cytokines. As the literature conflicts 364
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as to whether tick infestation blocks B. borgdorferi transmission (40, 55) and 365
what type of cytokine response is favorable or detrimental for spirochete 366
transmission (24, 56), the role that the IL-4 induced by repeated tick infestation 367
plays in resistance to Lyme disease is unclear. In contrast to sand flies and 368
mosquitoes, ticks take several days to complete feeding. The modulation of host 369
immunity towards Th2 cytokine expression may provide an evolutionary 370
advantage to ticks to avoid host sensitization to tick feeding; rapid feeding 371
arthropods may not require such immunomodulation. Interestingly, animal 372
species that have acquired immunity to tick feeding express cutaneous basophil 373
hypersensitivity reactions (CBH) at attachment sites (3), a reaction mediated by 374
Th1 responses (20). Furthermore, even though the immune response of mice to 375
repeated tick infestations is predominated by IL-4, IFN-g (21) and IgG2a (11) 376
levels increase with multiple exposures. In conjunction with our observations, a 377
model is beginning to emerge indicating that repeated exposure to rapid feeding 378
arthropods induces Th1 profiles that lead to increased resistance to pathogen 379
transmission. 380
381
It previously has been demonstrated that treatment of BALB/c mice with 382
recombinant IL-12 prior to sporozoite challenge protects against P. yoelii 383
infection (47), suggesting that short-term prophylaxis with rIL-12 could be used to 384
combat malaria. The increased level of IL-12p40 mRNA detected in pre-385
sensitized animals suggests that it is possible that ‘vaccination’ with uninfected 386
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mosquitoes may be an efficient method to induce IL-12 and avoid the toxicity 387
associated with treatment with recombinant proteins. 388
389
It is thought that individuals living in mosquito intense areas naturally become 390
desensitized to mosquito bites. The particular mechanism of this tolerance 391
remains to be defined, but is known to be associated with a loss of the wheal and 392
flare reactions of type I hypersensitivity as well as delayed reactions (38). 393
Although cytokine responses have not been evaluated in desensitized 394
individuals, the hypersensitivity reactions in response to mosquito saliva are Th2 395
mediated (i.e. IgE-mediated for Type I and eosinophil mediated for Type IV). 396
Therefore it is possible that desensitization may cause a reduction in Th2 397
cytokines, thus promoting an even greater Th1 environment than we detected in 398
our model system. 399
400
New vaccine targets and novel strategies will be essential for the ultimate 401
success of malaria vaccine development and data suggests that any measure 402
that limits parasite densities in the liver will reduce the morbidity and mortality 403
associated with malaria infection (33). Our findings imply that mosquito salivary 404
constituents could be effective components in such a vaccine. In this context, 405
saliva can be thought of as a non-specific potentiator; as long as vaccinated 406
individuals encounter malaria together with mosquito saliva the potentiator will be 407
effective at inducing a Th1 biased environment that is known to be effective 408
against malaria infection. 409
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Acknowledgements 410
411
We thank Freimann Life Science Center for excellent animal care, and Dr. John 412
Adams for helpful discussions concerning murine malaria. We also are grateful 413
to Dr. Tom McCutchan for advice on the Plasmodium 18S RNA assays and to 414
Ursula Krzych for her insightful suggestions. This work was supported by a grant 415
from the Defense Advanced Research Projects Agency of the Department of 416
Defense (#W911NF-04-1-0380).417
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References 418
1. Alger, N. E., and J. Harant. 1976. Plasmodium berghei: sporozoite challenge, 419
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Figure Legends 606
Figure 1. Lower Parasite Burden in mice pre-sensitized to A. stephensi 607
saliva. Mice in naïve and pre-sensitized groups were exposed to the bites of P. 608
yoelii-infected A. stephensi. (A) Parasite Burden in the liver was detected by 609
quantitative real time RT-PCR and infection levels were normalized to mouse 610
HPRT and expressed as 40-∆∆ct. (B) Naïve (open squares) and Pre-sensitized 611
(open triangles) RNA isolated from the liver also was analyzed using a standard 612
curve generated using RNA harvested from known numbers of salivary gland 613
sporozoites (closed squares). (C) Blood stage infection was monitored each day 614
for 7 days via blood smear from both naïve (solid bars) and pre-sensitized (lined 615
bars) mice. This data is representative of three independent experiments (n=4-5 616
per experiment). 617
* p<0.05 and **p<0.10 by student’s T Test. 618
619
Figure 2. Pre-sensitization skews response towards Th1 Phenotype. 620
Cytokine levels were quantified in naïve (Naïve), pre-sensitized (Pres) and in 621
naïve (Naïve Bit) and pre-sensitized (Pres Bit) mice that received one final 622
exposure to A. stephensi bites 24 hours prior to analysis. Local (ear) (A-C) and 623
systemic (liver (D-F) and spleen (G-I)) tissue IFN-g (A,D,G) and IL-4 (B,E,H) 624
mRNA levels were quantified by quantitative real time PCR. (C,F,I) IFN-g and IL-625
4 expression levels were used to create a cytokine ratio (IFN-g:IL-4). Data is 626
representative of four independent experiments (n=5 per experiment). Error bars 627
representative of S.E.M. * p<0.05 and ** p<0.001 by Student’s T Test. 628
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629
Figure 3. Lower hepatic parasitemia is IFN-g-dependent. Balb/c WT (closed) 630
and IFN-g KO (open) animals in both Naïve (squares) and Pres (triangles) 631
groups were exposed to the bites of P. yoelii-infected A. stephensi. (A) parasites 632
and (B) iNOS mRNA were quantified by quantitative real time RT-PCR 40 hours 633
post infection. Infection levels were normalized to mouse HPRT and expressed 634
as relative parasite burden (40-∆∆ct). Data is representative of two independent 635
experiments. (n=5 per experiment * p< 0.05). Error bars are representative of 636
S.E.M.. 637
638
Figure 4. Local induction of iNOS occurs 5 hours after exposure to 639
infected A. stephensi. Balb/c animals in both Naïve and Pres groups were 640
exposed to the bites of P. yoelii-infected A. stephensi and iNOS mRNA was 641
quantified by quantitative real time RT-PCR 5 hours post infection. Error bars 642
are representative of S.E.M. *p< 0.05 by Student’s T Test. Data is representative 643
of two independent experiments. (n=5 per experiment). 644
645
Figure 5. IL-12p40 is induced by Pre-sensitization. Balb/c animals in both 646
Naïve and Pres groups were exposed to the bites of P. yoelii-infected A. 647
stephensi and IL-12p40 mRNA was quantified in liver (A) and spleen (B) 40 648
hours post infection by quantitative real time RT-PCR. Error bars are 649
representative of S.E.M. *p< 0.05 by Student’s T Test. Data is representative of 650
two independent experiments. (n=4 per experiment). 651
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652
Figure 6. Protective phenomenon associated with pre-sensitization to 653
mosquito saliva evident at 20 hours post-infection and is localized to the 654
liver. Pre-sensitized (lined bars) and naïve (solid bars) animals were subjected 655
to the bites of P. yoelii 17xNL-infected A. stephensi. Animals were sacrificed at 656
5, 10, 20, and 30 hours post-infection and 18s rRNA levels were quantified in 657
ears (A) and livers (B). Pre-sensitized and naïve animals were infected with 1000 658
P. yoelii 17xNL sporozoites intravenously through the tail vein immediately 659
following a 4th pre-sensitization to uninfected bites, and euthanized 40 hours 660
post-infection, subsequently parasite 18s rRNA levels were quantified by 661
quantitative real time RT-PCR (C). Infection levels were normalized to mouse 662
HPRT and expressed as relative parasite burden (40-∆∆ct). Data is 663
representative of two independent experiments (n=4). Error bars represent the 664
S.E.M. *p<0.05 by Student’s T Test. 665 ACCEPTED
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