Sand fly saliva compounds are able to elicit specific immune responses that have a significant role in Leishmania parasite establishment and disease outcome. Characterizing anti-saliva immune responses in individuals living in well defined leishmaniasis endemic areas would provide valuable insights regarding their effect on parasite transmission and establishment in humans.
We explored the cellular and humoral immune responses to Phlebotomus (P.) papatasi salivary gland extracts (SGE) in individuals living in cutaneous leishmaniasis (CL) old or emerging foci (OF, EF). OF was characterized by a higher infection prevalence as assessed by higher proportions of leishmanin skin test (LST) positive individuals compared to EF. Subjects were further subdivided into healed, asymptomatic or naïve groups. We showed anti-SGE proliferation in less than 30% of the individuals, regardless of the immune status, in both foci. IFN-γ production was higher in OF and only observed in immune individuals from OF and naïve subjects from EF. Although IL-10 was not detected, addition of anti-human IL-10 antibodies revealed an increase in proliferation and IFN-γ production only in individuals from OF. The percentage of seropositive individuals was similar in immune and naïves groups but was significantly higher in OF. No correlation was observed between anti-saliva immune responses and LST response. High anti-SGE-IgG responses were associated with an increased risk of developing ZCL. No differences were observed for anti-SGE humoral or cellular responses among naïve individuals who converted or not their LST response or developed or not ZCL after the transmission season.
These data suggest that individuals living in an old focus characterized by a frequent exposure to sand fly bites and a high prevalence of infection, develop higher anti-saliva IgG responses and IFN-γ levels and a skew towards a Th2-type cellular response, probably in favor of parasite establishment, compared to those living in an emerging focus.
During murine experimental leishmaniasis sand fly saliva components modulate the host immune response and facilitate infection while pre-exposition to uninfected sand fly bites is associated with a protective cellular response against subsequent infection. Human anti-saliva immune responses are not well defined in leishmaniasis endemic areas. Here, we report an analysis of anti P. papatasi saliva cellular and humoral responses in individuals residing in endemic foci showing different prevalence rates of L. major infection. Individuals were further subdivided based on LST response and presence of typical CL scars. We showed higher anti-saliva cellular and humoral responses and a skew towards a Th2 response in the old focus characterized by the highest prevalence of infection. No correlation was observed between LST and anti-saliva cellular or humoral response. We showed that high anti-saliva IgG responses constituted a risk factor for the development of CL. Our findings suggest that the anti-P. papatasi saliva cellular and humoral response profiles vary with the level of sand fly exposure and the prevalence of infection in CL endemic areas. Such studies in humans from highly endemic areas could contribute to a better understanding of the immune response to sand fly saliva and its role in leishmaniasis outcome.
Citation: Kammoun-Rebai W, Bahi-Jaber N, Naouar I, Toumi A, Ben Salah A, Louzir H, et al. (2017) Human cellular and humoral immune responses to Phlebotomus papatasi salivary gland antigens in endemic areas differing in prevalence of Leishmania major infection. PLoS Negl Trop Dis 11(10): e0005905. https://doi.org/10.1371/journal.pntd.0005905
Editor: Martin Olivier, McGill University, CANADA
Received: April 17, 2017; Accepted: August 24, 2017; Published: October 12, 2017
Copyright: © 2017 Kammoun-Rebai 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.
Data Availability: All relevant data are within the paper.
Funding: This work was supported by the National Institutes of Health/National Institute of Allergy and Infectious Diseases (NIH/NIAID) to ABS and HL. Grant number: SP50AI074178. 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.
Leishmaniasis caused by protozoan parasites of the genus Leishmania transmitted by phlebotomine sand fly vectors has one of the largest diseases burden among the neglected tropical diseases [1,2]. These infections cause a broad clinical spectrum including cutaneous, mucocutaneous or visceral forms with variable severity, depending on the parasite species and the host immune status [3,4]. Leishmaniasis can also be asymptomatic in humans [4–6]. Disease control is mainly based on surveillance of incident cases and treatment, which is expensive, toxic and often associated with the emergence of drug-resistant strains . In Tunisia, zoonotic cutaneous leishmaniasis (ZCL) due to Leishmania (L.) major is the most frequent clinical form. Thousands of cases are reported every year since its first emergence as an epidemic in central Tunisia in 1982. The disease has spread in many parts of the country, with the emergence of several new foci and constitutes a public health problem [8–10]. L. major is transmitted by the bite of Phlebotomus (P.) papatasi [11,12]. Sand fly bite is a critical event in Leishmania transmission and saliva of this vector is a determining factor in infection. It has been shown that co-inoculation of Leishmania parasites with saliva enhances disease progression [13–16]. These exacerbating effects have been attributed to salivary proteins such as maxadilan, a vasodilator in Lutzomyia (Lu.) longipalpis saliva, and associated with immuno-modulatory activities, including inhibition of IFN-γ ability to activate macrophages to kill the intracellular parasite, up-regulation of Th2 cytokines production, down-regulation of some molecules important in parasite destruction such as nitric oxide and alterations in dendritic cell phenotype and function [15–22]. However, it has also been demonstrated that repeated exposure to uninfected sand fly bites elicits saliva-specific Th1-mediated delayed-type hypersensitivity (DTH) responses that were associated with protection against Leishmania infection in mice [15,23,24]. Immunization with individual salivary proteins was shown to induce distinct immune profiles that correlated with resistance or susceptibility to L. major infection in animal models [25–29]. DTH to sand fly bites was also described in humans [24,30] and more recently in a cutaneous leishmaniasis (CL) endemic area where it was shown to be Th1-mediated . However, effects of sand fly saliva during natural exposure of individuals in leishmaniasis endemic areas, on modulation of parasite-specific immune response and outcome of infection, are not clearly understood. Field studies have demonstrated the presence of anti-saliva antibodies in humans naturally exposed to sand fly bites [32–35]. The use of antibodies against sand fly saliva or against salivary recombinant proteins as potential markers of exposure was suggested in endemic areas of leishmaniasis [34–39]. Antibodies to saliva might also be used as markers for the risk of Leishmania infection [34,35,40]. Few studies have assessed human cellular immune responses to sand fly saliva and its effects on parasite establishment. PBMC from human volunteers experimentally exposed to Lu. longipalpis bites displayed an increased frequency of CD4+ and CD8+ T cells as well as an increase in IFN-γ and IL-10 production upon stimulation with saliva . Analysis of cellular immune responses against P. papatasi saliva in PBMC from individuals naturally exposed to L. major infection, showed low proliferation, absence of IFN-γ production but significant IL-10 levels, which could favor establishment of infection .
To better understand the impact of host immune response to P. papatasi saliva on L. major infection development and to determine potential correlations between anti-saliva immune response and leishmaniasis outcome during natural exposure, we have investigated here the humoral and proliferative responses, as well as cytokine production to P. papatasi salivary gland extracts, in a large cohort of individuals living in CL old or emerging foci differing in prevalence of L. major infection, in Tunisia.
Materials and methods
This research was conducted with the approval of the local ethical Committee of the Pasteur Institute of Tunis (protocol number 07–0018). Written informed consent was obtained from all individuals and from parents or legal guardians in case of minors before enrollment.
Study area and target population
Our cohort consisted of 790 individuals aged from 6 to 20 years (mean age 12.5+/-3.5) randomly selected from 5 zoonotic foci of CL due to L. major situated in central Tunisia. During this prospective study, a specific questionnaire to collect informations on past history of ZCL, clinical examination for the presence of typical scars leishmanin skin test (LST) (for detection of exposure to Leishmania parasites) and peripheral blood sampling were performed for all individuals, in April, prior to the transmission season of CL, which occur between June and October  (Fig 1).
790 participants living in endemic areas of ZCL were followed up over 1 year throughout one season of L. major transmission. Parameters such as LST and the presence of typical scars were monitored at the beginning of the study and after the transmission season and the triggering of new cases. Peripheral blood samples were obtained from each donor at the beginning of the study.
The LST was considered positive if the mean of the 2 diameters of induration was five mm or more [42,43]. A clinical follow up of one year throughout one L. major transmission season and a second LST (LST2) as well as the detection of new CL cases, were carried out. 29 new active CL cases were detected during the subsequent transmission season. CL diagnosis was based on clinical criteria and the demonstration of Leishmania parasites in Giemsa-stained dermal smears by microscopy.
All individuals taking part in our study come from a larger cohort of individuals living in the 5 foci of CL that have been recently analyzed during an LST-epidemiological study attempting to estimate the prevalence of L. major infection based on LST reactivity . Mnara was an old-focus (OF) whereas, Mbarkia, Dhouibet, Msaâdia and Ksour were considered as emerging foci (EF), on the basis of case notification data in the district epidemiological surveillance system. There were no significant differences between the 5 foci regarding demographic and socioeconomic characteristics. Bettaieb et al. demonstrated a significantly higher prevalence of infection in the old focus whereas no differences regarding this parameter were observed between the 4 emerging foci . For our cohort, we further compared the clinical and immunological parameters related to exposure and infection by Leishmania, namely the median size of the LST and LST2, the percentage of individuals with positive LST and LST2, the median number of scars and lesions per individual, the percentage of individuals with scars, the percentage of healed (LST+/Scar+), asymptomatic (LST+/Scar-) and naïve (LST-/Scar-) individuals and the number of active ZCL cases, between the four emerging foci, and showed no significant differences (Table 1).
However, these parameters were significantly different between the old focus and combined emerging foci, especially for LST size reaction (median/IQR: 12/4 and 0/9, respectively) and percentage of individuals with positive LST (98.3% and 41.2%, respectively) (Table 2). Taking these data into account, individuals from the old focus (OF) and those from emerging combined foci (EF) were considered separately.
Human groups were defined based on the following inclusion criteria (i) living in endemic area of L. major infection, information on past history of ZCL based on a specific questionnaire, positive LST reaction and presence of typical CL scars (LST+Scar+) for healed individuals (ii) living in endemic area of L. major infection, positive LST reaction and absence of typical CL scars (LST+Scar-) for individuals with a probable asymptomatic infection and (iii) living in endemic area of L. major infection, negative LST reaction and absence of typical CL scars (LST-Scar-) for naïve individuals.
We first analyzed anti-SGE humoral and cellular responses in all individuals from OF (n = 184, n = 82; respectively) and EF (n = 606, n = 312; respectively), regardless of their response to LST or the presence of scars (Table 3). In a second step we performed this analysis in LST+/Scar+ (humoral/cellular responses performed in 74/33 individuals), LST+/Scar- (109/49) groups from OF and LST+/Scar+ (101/45), LST+/Scar- (159/79) and LST-/Scar- (331/173) groups from EF. Luminex assay was performed in a subgroup of 58 EF individuals subdivided into 17 LST+/Scar+, 23 LST+/Scar- and 18 LST-/Scar- (Table 3).
Salivary gland extracts preparation
Sand fly salivary glands were kindly provided by E. Zhioua and S. Cherni (Laboratory of Vector Ecology, Pasteur Institute of Tunis). They were dissected out from a colony of the Tunisian P. papatasi vector of L. major and stored in groups of 20 pairs in 20μl phosphate buffered saline (PBS). Immediately before use, salivary glands were disrupted by 5 freezing/thawing cycles. After centrifugation, the supernatants were stored at –80°C with 10% glycerol. Just before use, SGE was prepared by dilution in cell culture medium added with gentamycin (Invitrogen).
Peripheral Blood Mononuclear cells (PBMC) were isolated from blood by density centrifugation through Ficoll-Hypaque (Pharmacia, Uppsala, Sweden). Cells were resuspended in RPMI 1640 culture medium supplemented with 1% Hepes 1M, sodium Pyruvate 1mM, 1X non essential amino acids, 0.1% β mercapto-ethanol 50mM, 10% heat inactivated AB human serum, 2mM L-glutamine, 100 IU/ml penicillin and 100 mg/ml streptomycin. Purified blocking anti-human IL-10 antibody (BD Biosciences, Le Pont de Claix, France) was used in some cell culture conditions. PBMC were cultured at 105 cells/well in triplicates in 96 well plates in a final volume of 200 μL and incubated with SGE (1gland/ml) with or without an anti-IL10 antibody (500ng/ml), at 37°C, in a 5%CO2 humidified atmosphere, for 5 days. Purified Protein Derivative (PPD) (5μg/ml) was used as a positive control. Control experiments were also performed on non-stimulated PBMC. During the last 10 hours of incubation, 0.4μCi/well of 3H-thymidine (Amersham) was added to cultures. The cells were harvested on glass-fiber filters using a multichannel cell harvester (PHD cell harvester, Cambridge technology) and 3H-thymidine incorporation was measured by liquid scintillation counter (Rack Beta; LKB wallace). Results were expressed as stimulation index (SI) obtained by dividing the mean counts of triplicates in antigen-stimulated cultures by the mean counts of triplicates in non-stimulated cultures. Lymphoproliferation was considered positive when SI was superior or equal to 2.
Cytokine detection by ELISA
IL-10 or IFN-γ levels were measured by Enzyme-linked immunosorbent assay (ELISA) in cell culture supernatants collected after 48h or 72h, respectively, centrifuged and stored at -80°C until use. Human IL-10 or IFN-γ ELISA Sets (BD Biosciences) were used according to manufacturer’s instructions. For each cytokine determination, the results were interpolated from a standard curve using recombinant cytokines and expressed in pg/mL.
Analysis of antibody response against SGE
Specific anti-saliva IgG antibodies were measured by ELISA. The wells (NUNC, Maxisorp Roskilde, Denmark) were coated with SGE (0.5 glands/well) in 0.1 M carbonate-bicarbonate buffer (pH 9.6) overnight at 4°C. After three washes with PBS-0.05% Tween, free binding sites in the plates were blocked for 1 hour at 37°C with PBS containing 0.1% Tween 20 and 0.5% gelatine (PBS-T-G). Human sera were diluted (1:200) in PBS-T-G and incubated for 2 hours at 37°C. After washing, wells were incubated with peroxidase-conjugated anti-human IgG antibodies (Sigma) at a 1:5000 dilution in PBS-T-G, for 1 hour at 37°C. Tetramethylbenzidine (TMB) substrate (Sigma) was added for 10 min at room temperature to visualize antibody-antigen complexes. The absorbance was measured using an automated ELISA reader (Thermo Labsystems Multiskan Ex) at 450 nm to 620 nm wavelengths. All sera were tested in duplicate and the mean value was recorded. Results were expressed as relative OD (ROD) defined as the ratio of sample OD/mean OD of sera from 20 negative controls (+ two standard deviations). ROD superior or equal to 2 was considered positive. Negative sera were obtained from healthy controls living outside Tunisia in sand fly-free regions.
Cytokines and chemokines measurements by Luminex assay
Seventeen cytokines and chemokines (IFN-γ, IL-1β, IL-2, IL-4, IL-5, IL-6, IL-10, IL-12p70, IL-13, GM-CSF, TNFα, IL-18, MIP- 1α, IL-8, IL-17A, MCP-1, M-CSF) were analysed using the Luminex multiplex bead-based technology, in culture supernatants from cells stimulated for 48h with SGE, according to the manufacturer’s instructions (Affymetrix, eBioscience).
Data analysis was performed with Stata statistical software (StataCorp. 2009. Stata Statistical Software: Release 11. College Station, TX: StataCorp LP.). Results were expressed as medians (interquartile range, IQR as variances). P values less than 0.05 were considered statistically significant. We used Wilcoxon signed-rank or Mann-Whitney test to determine intragroup differences of cytokine median levels (i.e., differences between stimulated and non-stimulated cultures). Kruskal-Wallis rank test was used for intergroup analysis on normalized data after deducting the non-stimulated value. Proportions for categorical variables were compared using chi-square test. Correlations were estimated using the Spearman rank (rs) correlation coefficient.
Logistic regression was applied to identify risk factors of ZCL cases. Univariate analysis for the following variables was used: age, gender, LST, presence of scars, foci, proliferative responses to (SI <2 and > = 2), IFN-γ responses to SGE, and anti-SGE IgG responses (ROD <2 and > = 2). Variables in univariate analysis with p values less than 0.25 were included in binary multivariate logistic regression. The final model was obtained by backward selection using a significance level of 5%.
Cell proliferation to P. papatasi SGE in individuals living in old versus emerging foci for ZCL
Proliferative responses to SGE were evaluated in 82 individuals living in OF and 312 individuals living in EF. Positive proliferative responses (IS≥2) were observed in 20.7% (17/82) and 28.8% (90/312) of individuals with median SI/IQR of 3.3/2.7 and 3.1/2.5 in OF and EF, respectively (Fig 2A). No significant differences were observed between both foci. In OF, where only asymptomatic (n = 49) and healed individuals (n = 33) were present, the percentage of positive SGE responders and median SI were similar in both groups (20.4% (10/49); 3.9/2.1 and 21.2% (7/33); 2.8/4.5, respectively) (Fig 2B). In EF, where naïves (n = 173), asymptomatics (n = 79) and healed (n = 45) were present, higher percentage of SGE positive responders was observed in naïve group (32.3%: 56/173) compared to asymptomatic (26.5%: 21/79) and healed (15.6%: 7/45) groups. Among the immune subjects, the proportion of positive responders was higher in asymptomatic individuals. However, differences were not statistically significant and median SI was similar in the three groups (3/2.5 in naïves; 3.2/3.6 in asymptomatics and 3.4/0.7 in healed individuals) (Fig 2B). Furthermore, no significant correlation was found between proliferative responses to SGE and LST in both foci. We further analyzed proliferation results, in EF, by grouping the immune individuals into a single group, based only on the LST response (n = 124 for LST+ individuals). We showed again higher but not significant percentage of SGE positive responders in naïve individuals (32.3%) compared to immune subjects (22.5%) (Fig 2C).
PBMC were stimulated for 5 days with P. papatasi salivary gland extracts (1gland/ml). Proliferation was analyzed in individuals from donors residing in an old (OF) or an emerging focus (EF) (a), LST+/Scar+ (healed), LST+/Scar- (asymptomatics) and LST-/Scar- (naïve) groups (b) and LST+, LST- groups (c), in each focus. Results were expressed as stimulation index (SI) obtained by dividing the mean counts of triplicates in antigen-stimulated cultures by the mean counts of triplicates in non-stimulated cultures. The proliferative response was considered positive when SI was superior or equal to 2. Statistical significance was assigned to a value of p<0,05. Horizontal lines represent median SI values and dotted lines represent cut-off level. The percentages of positive responders are presented for groups and foci.
Cytokine responses to P. papatasi SGE in individuals living in old versus emerging foci for ZCL
IFN-γ and IL-10 responses to SGE were analyzed by ELISA on the same individuals that were evaluated for proliferative responses to saliva. Significant IFN-γ levels were observed in response to SGE stimulation when compared to non-stimulated cultures, in OF (median/IQR: 1136/2307; 775/2016 pg/ml, respectively) and EF (12/39; 9/26 pg/ml) (p = 0.01), (Fig 3A). These levels were significantly higher in OF compared to EF (p = 0.03). IFN-γ levels were only significant in individuals with positive proliferation, in both foci (OF (stimulated, non-stimulated cultures): 1744/2363, 665/1308 pg/ml, p = 0.0009; EF: 20/84.7, 9/28 pg/ml, p = 0.0000). A positive correlation was found between SGE proliferative response and IFN-γ production in both OF (r = 0.22; p = 0.047) and EF (r = 0.26; p<0.0001). Interestingly, analysis of IFN-γ responses to SGE according to LST response and presence of scars revealed significant IFN-γ levels only in naïves individuals from EF (13/41, 8/25 pg/ml, in stimulated and non-stimulated cultures, p = 0.007) (Fig 3B). We did not detect IFN-γ response in asymptomatics and healed individuals in both foci. However, when we analyzed again IFN-γ response in groups subdivided only on the basis of the LST response, we observed a significant IFN-γ production in LST+ (1152/2307 and 778/2017; p = 0.01) group, only in OF (Fig 3C). The difference between this result and the one observed in groups subdivided according to LST and scars could be explained by a lower number of individuals in the latter groups. No correlation was found between anti-SGE IFN-γ response and LST in both EF and OF.
IFN-γ was quantified by ELISA, in the culture supernatants of PBMC stimulated with P. papatasi salivary gland extracts (1 gland/ml) during 72h. Analysis of IFN-γ production was performed in individuals from OF and EF (a), LST+/Scar+ (healed), LST+/Scar- (asymptomatics) and LST-/Scar- (naïve) groups (b) and LST+, LST- groups (c), in each focus. Statistical significance was assigned to a value of p<0,05. Statistically significant differences between stimulated and non-stimulated cultures and between groups are showed. Horizontal lines represent median IFN-γ values.
In vitro SGE stimulation did not cause a significant change in IL-10 production, when compared to non-stimulated cultures in OF and EF. Furthermore, no significant IL-10 levels were observed in response to SGE in healed, asymptomatic and naïves individuals.
In order to extend the analysis of cytokine response to P. papatasi saliva, additional cytokines and chemokines (IFN-γ, IL-10, IL-1β, IL-2, IL-4, IL-5, IL-6, IL-12p70, IL-13, GM-CSF, TNFα, IL-18, MIP-1α, IL-8, IL-17A, MCP-1, M-CSF) were evaluated, using Luminex, in culture supernatants from cells issued from 17 healed, 23 asymptomatic and 18 naïve individuals (randomly selected among the EF subjects) and stimulated for 48h with SGE. None of these cytokines or chemokines was detected at significant levels, in response to SGE stimulation. The detection of significant levels of IFN-γ by ELISA but not by Luminex, could be explained by the lower number of individuals analyzed using the Luminex assay.
Cellular responses to SGE in the presence of an anti-IL10 antibody
Although we have demonstrated the presence of a cellular response against P. papatasi saliva in individuals living in endemic areas of CL, both in terms of proliferation and IFN-γ production, this response was generally low. It was shown that P. papatasi saliva was able to induce the activation of IL-10 producing T cells in naturally exposed individuals . Although we did not detect any significant IL-10 production in response to SGE, we further analyzed proliferation and IFN-γ production to SGE in the presence of an anti-human IL-10 antibody, in order to determine whether the observed SGE-specific cellular responses are down-regulated. We observed that addition of anti-IL-10 antibody significantly increased cellular proliferation only in the old focus (percentage of positive responders: 41.9% and 20.7%, respectively with or without anti-IL-10 antibody, p = 0.007) (Fig 4A). A significant increase was also observed in the percentage of positive responders in asymptomatic and healed groups from OF (40.4 and 20.4%, p<0.0001 in asymptomatic group; 44.1 and 21.2%, p = 0.012, in healed group, respectively with or without anti-IL-10 antibody (Fig 4B). Anti-IL-10 antibody had no significant effect on the cellular response against SGE in individuals from the emerging focus. Median SI values were not affected by anti-IL-10 antibody addition, in all groups and foci.
PBMC were stimulated with P. papatasi salivary glands extracts (1gland/ml) with or without an anti-IL10 antibody (500ng/ml), for 5 days. Proliferation was analyzed in individuals from donors residing in an old (OF) or an emerging focus (EF) (a) and LST+/Scar+ (healed), LST+/Scar- (asymptomatics) and LST-/Scar- (naïve) groups (b), in each focus. Results were expressed as percentage of positive proliferative responders. The proliferative response was considered positive when SI was superior or equal to 2. Statistical significance was assigned to a value of p<0,05. Statistically significant differences between groups are showed.
IFN-γ production was significantly increased when anti-IL-10 antibody was added to cultures, in individuals from OF (712/1491 and 30/784, p = 0.0000, respectively with or without anti-IL-10 antibody) but not in those from EF (Fig 5A). Interestingly, addition of anti-IL-10 antibody allowed to detect significant levels of IFN-γ in asymptomatic (2004/2786, 1071/1995; in stimulated and non-stimulated cultures, p = 0.0000) and healed (1740/2483, 455/1943; p = 0.0000) groups from OF (Fig 5B). This cytokine was not detected in these groups, in the absence of anti-IL-10 antibody. Anti-IL-10 antibody did not affect IFN-γ production in groups from EF (Fig 5C).
IFN-γ was quantified by ELISA, in the culture supernatants of PBMC stimulated with P. papatasi salivary gland extracts (1 gland/ml), with or without an anti-IL10 antibody (500ng/ml), during 72h. Analysis of IFN-γ production was performed in individuals from OF and EF (a) and LST+/Scar+ (healed), LST+/Scar- (asymptomatics) and LST-/Scar- (naïve) groups in OF (b) and EF (c). Statistical significance was assigned to a value of p<0.05. Statistically significant differences between stimulated and non-stimulated cultures and between groups are showed. Horizontal lines represent median IFN-γ values.
IgG responses to P. papatasi salivary gland extract in individuals living in old versus emerging foci for ZCL
We measured the reactivity of SGE, using plasma samples from 184 and 606 individuals living in OF and EF foci, respectively. We showed a significantly higher proportion of seropositive individuals in the old focus (63.04%, 116/184) compared to the emerging one (50.8%, 308/606), (p = 0.004), (Fig 6A). Median ROD/IQR was also significantly higher in OF (2.46/2.3), in comparison with EF (2.02/2) (p = 0.0006). However, median ROD of seropositive individuals was similar in both foci (OF: 3.2/2.5; EF: 3.2/1.8). Furthermore, no significant differences were observed in terms of proportion of seropositive individuals or median ROD, between asymptomatic (n = 109) (64.2%; 3.31/3.4) and healed (n = 74) (60.8%; 3.16/1.9) groups from OF or asymptomatic (n = 159) (48.4%; 3.04/1.9, healed (n = 101) (48.3%; 2.8/1.9 and naïve (n = 331) (52.8%; 3.5/1.8) groups from EF (Fig 6B). No correlation was found between anti-saliva IgG response and LST in both foci.
Analysis of IgG response was performed in individuals from OF and EF (a) and LST+/Scar+ (healed), LST+/Scar- (asymptomatics) and LST-/Scar- (naïve) groups (b), in each focus. IgG antibodies were measured by ELISA. Results were expressed as relative OD (ROD) defined as the ratio of sample OD/mean OD of sera from negative controls. ROD superior or equal to 2 was considered positive. Negative sera were obtained from healthy controls living outside Tunisia in sand fly-free regions. Statistical significance was assigned to a value of p<0,05. Statistically significant differences between groups are showed for positive responders. Horizontal lines represent median ROD values and dotted lines represent cut-off level. The percentages of positive responders are presented for groups and foci.
In addition, we showed that the percentage of individuals with positive proliferation against SGE (SI+) was significantly higher among seropositive individuals (ROD+) compared to seronegative individuals (ROD-) in both foci (%SI+ in ROD+ and ROD-: 25% and 5.8% in OF; 39.5% and 16.5% in EF). Significant levels of IFN-γ were only observed in seropositive individuals in both foci (median IFN-γ/IQR in stimulated and non-stimulated cultures: 1038/2019, 689/1346; p = 0.001 in OF and 13/47, 9/28; p = 0.004 in EF). Significant correlations were found between SGE specific IgG responses and proliferative responses in both foci (OF: r = 0.38, p = 0.0004; EF: r = 0.3116, p <0.0001), and between IgG responses and IFN-γ responses only in EF (r = 0.16, p = 0.005). However, these correlations were not detected when asymptomatic, healed and naïve groups were considered.
Evaluation of the effects of specific-SGE cellular and humoral responses on LST conversion and on ZCL development
During the follow-up of one year throughout a L. major transmission season, a second LST (LST2) evaluation was carried out in individuals from both foci. To estimate the impact of the anti-saliva cellular response (proliferation and IFN-γ) on the conversion of the LST response, we compared the proportion of EF naïve individuals who converted their LST response without developing ZCL after a transmission season (LST-/LST2+/ZCL-) between naïve individuals with positive proliferation to saliva (n = 56) and those with no proliferation (n = 117). Despite a higher percentage of LST-/LST2+/ZCL- individuals in the first group (14.2%) compared to the second one (9.4%), no statistically significant differences were observed. IFN-γ was not detected in naïve individuals, whether or not they have converted their LST response, after a transmission season. Anti-IL-10 had no effect on IFN-γ response in these groups.
We also compared the percentage of EF naïve individuals who developed ZCL after a transmission season (LST-/ZCL+), between naïve individuals with positive proliferation to saliva and those with no proliferation, cited above. Out of 56 naïve individuals with positive saliva proliferation, no individual developed ZCL. All ZCL cases (n = 6) were observed among naïve individuals with no proliferation against SGE (5.12%, 6/117). However, no significant differences were detected between the two groups. Furthermore, IFN-γ was not detected in individuals who have developed ZCL. However, significant IFN-γ levels were observed in the group of individuals who remained naïves (n = 167) (14/42, 8/26, in stimulated compared to non-stimulated cultures, p = 0.01). The addition of anti-IL-10 had no effect on the production of IFN-γ in naïve individuals whether they develop ZCL or not.
No significant differences were observed for percentage of LST-/LST2+/ZCL- individuals between naïve SGE seropositive (10.8%: 19/175) or seronegative groups (8.9%: 14/156). Regarding the effects of anti-SGE humoral responses on the development of ZCL, we did not detect significant differences in the percentage of LST-/ZCL+ individuals between naïve SGE seropositive (2.3%: 4/175) or seronegative groups (1.3%: 2/156).
Multivariate analysis of risk factors for L. major infection in individuals exposed to P. papatasi
We undertook a multivariate analysis using logistic regression to identify risk factors associated with ZCL. During the one-year follow-up, 29 ZCL new cases were detected among individuals from both foci. The anti-SGE antibody response was identified as a risk factor for the development of ZCL. We showed that individuals with mROD≥4 had a risk factor of 2.65 (p = 0.023; 95% confidence interval, 1.14−6.14) to develop ZCL.
Here, we report an analysis of cellular and humoral responses developed against salivary gland extracts from P. papatasi, in a large cohort of 790 individuals living in areas characterized by hyperendemicity for ZCL infection caused by L. major. A close spatial association has been reported between the abundance of P. papatasi and the incidence of ZCL in central and southwestern Tunisia . Our study was conducted in old and emerging foci, in central Tunisia. The determinants of L. major infection were evaluated for both foci in a recent large LST epidemiologic study, showing a significantly higher prevalence of infection in the old focus as assessed by higher proportions of LST positive individuals compared to the emerging focus . In both foci, individuals were subdivided according to their LST response and the presence of typical ZCL scars into healed, asymptomatic and naïve individuals. Almost all participants had a positive LST, with either healed lesions or asymptomatic infection, in the old focus, whereas more than half of the individuals were naïve in the emerging focus. The LST reaction reflects a CD4+ Th1 cell-mediated immune response  and is a useful indicator of immune status in studies evaluating Leishmania vaccine candidates [45–47] and an important tool in epidemiological studies to assess exposure to Leishmania and the prevalence of infection [9,48–50].
We showed positive proliferative responses against P. papatasi saliva in less than 30% of individuals with relatively low proliferative indexes and low but significant IFN-γ levels, regardless of the immune status, in both foci. IFN-γ production, which was only observed in individuals with positive proliferation, was significantly higher in the old focus. According to the immune status, anti-saliva proliferation was also similar in healed, asymptomatic and naïve groups from both foci. However, IFN-γ was detected in LST positive individuals (asymptomatic and healed) in the old but not in the emerging focus. We did not observe a correlation between LST response and anti-saliva cellular responses both in terms of proliferation or IFN-γ responses. The observation of higher IFN-γ levels in the old focus suggest a more frequent exposure to sand fly bites in this focus. It has been observed that size of LST reaction, reflecting the time of exposure to the parasite and the intensity of the immune response, increased with lenght of residence in endemic areas [9,51]. In our cohort, the size of LST reaction was significantly higher in individuals from the old focus compared to those from the emerging one, suggesting longer exposure times to sand fly bites and probably higher infection rate of vectors leading to the development of an immunity against infection in almost all individuals living in the old focus. Within the emerging focus, where infecting bites are probably fewer than in the old focus, we showed a significant IFN-γ production only in naïve individuals, suggesting that the anti-saliva cellular immune responses vary depending on whether the host is exposed to saliva in the presence or absence of parasites. Host-parasite interactions within the insect vector before transmission could alter the vector salivary protein profile that may consequently influence the immune response of the human host . A role of the parasite in diminishing host antibody production against a salivary protein was suggested in a study showing a decrease in the immunogenicity of this protein in infected donors compared to donors exposed to sand fly bites but not infected .
Few studies have described human cellular responses against sand fly saliva in leishmaniasis endemic areas. A DTH-response (a manifestation of cell-mediated immunity) to P. papatasi, L. longipalpis or P. duboscqi bites, was previously reported in individuals wether after experimental or natural exposure [24,30,31]. An increase in the frequency of T CD4+ and T CD8+ cells, with IFN-γ and IL-10 production were described in PBMC from human volunteers experimentally exposed to L. longipalpis bites, stimulated with saliva . Similarly to our results, a specific proliferative response to P. papatasi saliva was showed in 30% of a small group of exposed donors in a CL endemic area and no correlation was observed between these responses and proliferation to soluble Leishmania antigens . However, absence of IFN-γ and IL-10 production by T CD8+ cells, in favor a Th2-type cellular response, was observed in this work. Authors showed the ability of activated anti-saliva CD4+ T cells to produce IFN-γ after being separated from CD8+ T cells . IFN-γ, IL-10, IL-5, IL-12 and IL-13 were produced in response to P. duboscqi saliva stimulation of PBMC from individuals living in a CL endemic area . More recently, a mixed T cell response with a predominance of IL-10 evidenced by elevated IL-10/IFN-γ and IL-10/IL-13 ratios was reported in individuals exposed to L. intermedia . The predominance of anti-saliva IL-10 response in individuals living in endemic areas led the authors to suggest that exposure to sand fly bites may facilitate Leishmania infection [41,54]. It was suggested that IL-10 production could be attributed to the presence of adenosine, a component of P. papatasi saliva, that have been reported as being able to inhibit NO synthesis and to induce IL-10 production in murine macrophages [55–57]. We did not detect IL-10 production in response to P. papatasi saliva in our study. Different culture conditions or differences in the sample size of the study groups or studied foci, may explain these different results. Furthermore, variability in the salivary repertoire of various sand fly species or populations that can induce different immune response pattern, may also explain these discrepancies [58–60]. Indeed, salivary molecules associated with immunomodulatory activities, such as adenosine, which is, present in P. papatasi saliva but absent from Lu. longipalpis and maxadilan which is only present in Lu. Longipalpis, has been identified . However, differences in immune responses elicited by salivary proteins from the same sand fly species could be attributed to genetic differences among hosts.
Despite the absence of IL-10 production in the peripheral blood in our study, it cannot be excluded that this cytokine is produced locally, at the site of infection or is present in peripheral blood at undetectable levels. Given the globally low anti-P. papatasi saliva cellular responses observed in our work and taking into account the published data about this sand fly saliva ability to induce IL-10 in humans, we further analyzed cellular responses using an anti-IL-10 antibody. We showed that addition of anti-IL-10 antibody induced a significant increase in both the percentage of individuals with positive anti-saliva T cell proliferation and IFN-γ production, in asymptomatic and healed groups, in the old focus. This result suggest that, in the old focus, saliva-specific T cell responses could be partially suppressed by IL-10 , even though not detected and emphasizes once again the difference between the anti-saliva immune response profiles, depending on the intensity of vector exposure. In endemic areas, individuals are mostly exposed to the bites of uninfected sand flies but are not protected against leishmaniasis. Exposure of mice to short- or long-term P. duboscqi bites followed by infection either immediately or with a delay, showed protection only in short-term exposed mice, suggesting that chronic exposure to salivary antigens may result in a change in the immune response of the host towards a Th2 profile . Addition of the anti-IL-10 antibody had no effect on cellular responses in individuals living in the emerging focus, suggesting that the absence of IFN-γ observed in the immune group would not be due to its inhibition by immunosuppressive cytokines such as IL-10.
We showed a significantly higher proportion of individuals with positive IgG responses to saliva in the old focus compared to the emerging one, which reveal once again differences in sand fly exposure between both foci. We did not show any change in the anti-saliva humoral response, depending on the LST response. No correlation was observed between anti-saliva cellular or humoral responses and LST, or between anti-saliva cellular and humoral responses, in both foci. The absence of correlation between proliferative and humoral response to P. papatasi saliva, was reported . Similar anti-L. whitmani IgG responses were also observed in LST positive and LST negative individuals in a recent study conducted in a CL endemic area with high transmission of L. braziliensis . However, unlike our findings, LST positivity was correlated to higher levels of P. duboscqi or L. longipalpis saliva specific IgG antibodies, in individuals living in CL or VL endemic areas, respectively [32,49]. Differences in salivary proteins from various sand fly species could explain these discrepancies [58–60,65].
Development of IgG antibodies against several sand fly species saliva, and the use of this response as an epidemiological tool for estimating exposure to various sandflies species, has already been reported in leishmaniasis endemic areas [33–36,40,53,54,64,66]. We evaluated the impact of the anti-saliva immune response on the conversion of the LST response and on the development of ZCL in naïve individuals of our cohort. No significant differences were observed for the proportion of individuals who converted their LST response without developing ZCL after a transmission season between individuals with positive proliferation or seropositive to saliva and those with no proliferation or seronegative. IFN-γ was not detected in naïve individuals, whether they converted or not their LST response. These results suggest that the development of a cellular or humoral response to P. papatasi saliva in naïve individuals, have no impact on the acquisition of a DTH to parasite during a subsequent transmission season, in our emerging focus. It has been reported in animal models that pre-exposition to uninfected sand fly bites is associated with the development of a protective cellular response against Leishmania infection [15,18,19,23,25,27,67,68]. However, a lack of protection against a challenge infection has also been reported in mice immunized with sand fly saliva . In VL endemic areas, increased anti-saliva IgG levels in individuals who developed a positive DTH to L. chagasi antigens, suggesting that the induction of a saliva humoral response could promote induction of a cellular response against the parasite [33,66,69].
We also showed that new ZCL cases were observed among individuals with no proliferation to saliva. Furthermore, no IFN-γ was produced in individuals who developed ZCL while a significant production was detected in those who remained naïve. It would be therefore tempting to suggest that the ability to develop a Th1 cell response against sandfly saliva prior to first contact with the parasite may lead to protection against infection. However, the statistical analysis did not reveal any significant difference, probably due to the low number of new ZCL cases detected during our study period. Our results are in line with data from a study showing that lymphocytes from donors experimentally exposed to sand fly bites seems to limit parasite burden in macrophages in in vitro autologous cell cutlture system . Finally, a multivariate analysis of ZCL risk factors showed that individuals with mROD≥4 had a risk factor of 2.65 (p = 0.023) to develop ZCL. Others have reported a correlation between the anti-saliva response and the risk of disease. Higher anti-saliva IgG levels were described in patients with active CL lesions in comparison with healthy or LST positive individuals [34,35,40]. More recently, it has been reported that individuals seropositive to saliva had a higher risk of developing CL than seronegative individuals .
Data presented in this study suggest that differences in sand fly exposure levels and in prevalence of infection are associated with different immune responses profiles against P. papatasi saliva. Higher anti-saliva IgG responses and IFN-γ levels and a skew towards a Th2-type cellular response were observed in immune individuals who are more exposed to sand fly bites and submitted to a higher prevalence of infection. We showed that anti-saliva IgG response did not interfere with the development of a DTH response to parasite but was a risk factor for the development of ZCL due to L. major. Together, these data can contribute to a better understanding of the mechanisms that govern the resistance or susceptibility to infection by L. major parasites upon transmission by the P. papatasi sandflies in endemic areas for ZCL.
We would like to thank all the patients of Sidi Bouzid and Kairouan who agreed to collaborate in this study. We also thank E. Zhioua and S. Cherni for providing the salivary glands of Phlebotomus papatasi and S. Marzouki for providing sera from healthy controls living outside Tunisia in sand fly-free regions. We are grateful to A. Gharbi, A. Zaatour, N. Belhaj Hamida for the logistical support in the field and W. Markikou-Ouni, M. Abdeladhim, N. Ben Hassouna and A. Millet for helpful assistance during the laboratory work. We thank Darragh Duffy (Laboratory of Dendritic Cell Immunobiology, Institut Pasteur, Paris, France) for the Luminex technology assistance. Experiments were performed in the laboratory of Dendritic Cell Immunobiology (Institut Pasteur, Paris, France).
- 1. Alvar J, Vélez ID, Bern C, Herrero M, Desjeux P, Cano J, et al. Leishmaniasis worldwide and global estimates of its incidence. PLoS One. 2012;7: e35671. pmid:22693548
- 2. Karimkhani C, Wanga V, Coffeng LE, Naghavi P, Dellavalle RP, Naghavi M. Global burden of cutaneous leishmaniasis: A cross-sectional analysis from the Global Burden of Disease Study 2013. Lancet Infect Dis. 2016;16: 584–591. pmid:26879176
- 3. Desjeux P. Leishmaniasis. Nat Rev Microbiol. 2004;2: 692. pmid:15378809
- 4. Bañuls a. L, Bastien P, Pomares C, Arevalo J, Fisa R, Hide M. Clinical pleiomorphism in human leishmaniases, with special mention of asymptomatic infection. Clin Microbiol Infect. 2011;17: 1451–1461. pmid:21933304
- 5. Sassi a., Louzir H, Salah a. Ben, Mokni M, Osman a. Ben, Dellagi K. Leishmanin skin test lymphoproliferative responses and cytokine production after symptomatic or asymptomatic Leishmania major infection in Tunisia. Clin Exp Immunol. 1999;116: 127–132. pmid:10209516
- 6. Andrade-Narvaez FJ, Loría-Cervera EN, Sosa-Bibiano EI, Van Wynsberghe NR. Asymptomatic infection with American cutaneous leishmaniasis: epidemiological and immunological studies. Mem Inst Oswaldo Cruz. 2016;111: 599–604. pmid:27759762
- 7. Oliveira LF, Schubach AO, Martins MM, Passos SL, Oliveira R V, Marzochi MC, et al. Systematic review of the adverse effects of cutaneous leishmaniasis treatment in the New World. Acta Trop. 2011;118: 87–96. pmid:21420925
- 8. Salah A Ben, Kamarianakis Y, Chlif S, Alaya N Ben, Prastacos P. Zoonotic cutaneous leishmaniasis in central Tunisia: Spatio-temporal dynamics. Int J Epidemiol. 2007;36: 991–1000. pmid:17591639
- 9. Bettaieb J, Toumi A, Chlif S, Chelghaf B, Boukthir A, Gharbi A, et al. Prevalence and determinants of Leishmania major infection in emerging and old foci in Tunisia. Parasit Vectors. 2014;7: 386. pmid:25142220
- 10. Chalghaf B, Chlif S, Mayala B, Ghawar W, Bettaieb J, Harrabi M, et al. Ecological Niche Modeling for the Prediction of the Geographic Distribution of Cutaneous Leishmaniasis in Tunisia. Am J Trop Med Hyg. 2016;94: 844–851. pmid:26856914
- 11. Ben Ismail R, Gramiccia M, Gradoni L, Helal H, Ben Rachid MS. Isolation of Leishmania major from Phlebotomus papatasi in Tunisia. Trans R Soc Trop Med Hyg. 1987;81: 749. Available: http://www.ncbi.nlm.nih.gov/pubmed/3449994 pmid:3449994
- 12. Chelbi I, Kaabi B, Béjaoui M, Derbali M, Zhioua E. Spatial correlation between Phlebotomus papatasi Scopoli (Diptera: Psychodidae) and incidence of zoonotic cutaneous leishmaniasis in Tunisia. J Med Entomol. 2009;46: 400–2. Available: http://www.ncbi.nlm.nih.gov/pubmed/19351095 pmid:19351095
- 13. Titus RG, Ribeiro JM. Salivary gland lysates from the sand fly Lutzomyia longipalpis enhance Leishmania infectivity. Science. 1988;239: 1306–8. Available: http://www.ncbi.nlm.nih.gov/pubmed/3344436 pmid:3344436
- 14. Theodos CM, Ribeiro JM, Titus RG. Analysis of enhancing effect of sand fly saliva on Leishmania infection in mice. Infect Immun. 1991;59: 1592–8. Available: http://www.ncbi.nlm.nih.gov/pubmed/2019430 pmid:2019430
- 15. Belkaid Y, Kamhawi S, Modi G, Valenzuela J, Noben-Trauth N, Rowton E, et al. Development of a natural model of cutaneous leishmaniasis: powerful effects of vector saliva and saliva preexposure on the long-term outcome of Leishmania major infection in the mouse ear dermis. J Exp Med. 1998;188: 1941–53. Available: http://www.ncbi.nlm.nih.gov/pubmed/9815271 pmid:9815271
- 16. Mbow ML, Bleyenberg JA, Hall LR, Titus RG. Phlebotomus papatasi sand fly salivary gland lysate down-regulates a Th1, but up-regulates a Th2, response in mice infected with Leishmania major. J Immunol. 1998;161: 5571–7. Available: http://www.ncbi.nlm.nih.gov/pubmed/9820534 pmid:9820534
- 17. Hall LR, Titus RG. Sand fly vector saliva selectively modulates macrophage functions that inhibit killing of Leishmania major and nitric oxide production. J Immunol. 1995;155: 3501–6. Available: http://www.ncbi.nlm.nih.gov/pubmed/7561045 pmid:7561045
- 18. Morris RV, Shoemaker CB, David JR, Lanzaro GC, Titus RG. Sandfly maxadilan exacerbates infection with Leishmania major and vaccinating against it protects against L. major infection. J Immunol. 2001;167: 5226–30. Available: http://www.ncbi.nlm.nih.gov/pubmed/11673536 pmid:11673536
- 19. Rohousová I, Volf P, Lipoldová M. Modulation of murine cellular immune response and cytokine production by salivary gland lysate of three sand fly species. Parasite Immunol. 2005;27: 469–73. pmid:16255746
- 20. Brodie TM, Smith MC, Morris R V, Titus RG. Immunomodulatory effects of the Lutzomyia longipalpis salivary gland protein maxadilan on mouse macrophages. Infect Immun. 2007;75: 2359–65. pmid:17339357
- 21. Wheat WH, Pauken KE, Morris R V, Titus RG. Lutzomyia longipalpis salivary peptide maxadilan alters murine dendritic cell expression of CD80/86, CCR7, and cytokine secretion and reprograms dendritic cell-mediated cytokine release from cultures containing allogeneic T cells. J Immunol. 2008;180: 8286–98. Available: http://www.ncbi.nlm.nih.gov/pubmed/18523295 pmid:18523295
- 22. Gomes R, Oliveira F. The Immune Response to Sand Fly Salivary Proteins and Its Influence on Leishmania Immunity. Front Immunol. 2012;3. pmid:22593758
- 23. Kamhawi S, Belkaid Y, Modi G, Rowton E, Sacks D. Protection against cutaneous leishmaniasis resulting from bites of uninfected sand flies. Science. 2000;290: 1351–4. Available: http://www.ncbi.nlm.nih.gov/pubmed/11082061 pmid:11082061
- 24. Belkaid Y, Valenzuela JG, Kamhawi S, Rowton E, Sacks DL, Ribeiro JM. Delayed-type hypersensitivity to Phlebotomus papatasi sand fly bite: An adaptive response induced by the fly? Proc Natl Acad Sci U S A. 2000;97: 6704–9. Available: http://www.ncbi.nlm.nih.gov/pubmed/10841567 pmid:10841567
- 25. Valenzuela JG, Belkaid Y, Garfield MK, Mendez S, Kamhawi S, Rowton ED, et al. Toward a defined anti-Leishmania vaccine targeting vector antigens: characterization of a protective salivary protein. J Exp Med. 2001;194: 331–42. Available: http://www.ncbi.nlm.nih.gov/pubmed/11489952 pmid:11489952
- 26. Oliveira F, Lawyer PG, Kamhawi S, Valenzuela JG. Immunity to Distinct Sand Fly Salivary Proteins Primes the Anti-Leishmania Immune Response towards Protection or Exacerbation of Disease. Lehane MJ, editor. PLoS Negl Trop Dis. 2008;2: e226. pmid:18414648
- 27. Gomes R, Teixeira C, Teixeira MJ, Oliveira F, Menezes MJ, Silva C, et al. Immunity to a salivary protein of a sand fly vector protects against the fatal outcome of visceral leishmaniasis in a hamster model. Proc Natl Acad Sci U S A. 2008;105: 7845–50. pmid:18509051
- 28. Oliveira F, Rowton E, Aslan H, Gomes R, Castrovinci PA, Alvarenga PH, et al. A sand fly salivary protein vaccine shows efficacy against vector-transmitted cutaneous leishmaniasis in nonhuman primates. Sci Transl Med. 2015;7: 290ra90. pmid:26041707
- 29. Reed SG, Coler RN, Mondal D, Kamhawi S, Valenzuela JG. Leishmania vaccine development: exploiting the host-vector-parasite interface. Expert Rev Vaccines. 2016;15: 81–90. pmid:26595093
- 30. Vinhas V, Andrade BB, Paes F, Bomura A, Clarencio J, Miranda JC, et al. Human anti-saliva immune response following experimental exposure to the visceral leishmaniasis vector,Lutzomyia longipalpis. Eur J Immunol. 2007;37: 3111–3121. pmid:17935072
- 31. Oliveira F, Traoré B, Gomes R, Faye O, Gilmore DC, Keita S, et al. Delayed-type hypersensitivity to sand fly saliva in humans from a leishmaniasis-endemic area of Mali is Th1-mediated and persists to midlife. J Invest Dermatol. 2013;133: 452–9. pmid:22992802
- 32. Barral A, Honda E, Caldas A, Costa J, Vinhas V, Rowton ED, et al. Human immune response to sand fly salivary gland antigens: a useful epidemiological marker? Am J Trop Med Hyg. 2000;62: 740–5. Available: http://www.ncbi.nlm.nih.gov/pubmed/11304066 pmid:11304066
- 33. Gomes RB, Brodskyn C, de Oliveira CI, Costa J, Miranda JC, Caldas A, et al. Seroconversion against Lutzomyia longipalpis saliva concurrent with the development of anti-Leishmania chagasi delayed-type hypersensitivity. J Infect Dis. 2002;186: 1530–4. pmid:12404176
- 34. Rohousova I, Ozensoy S, Ozbel Y, Volf P. Detection of species-specific antibody response of humans and mice bitten by sand flies. Parasitology. 2005;130: 493–9. Available: http://www.ncbi.nlm.nih.gov/pubmed/15991492 pmid:15991492
- 35. Marzouki S, Ben Ahmed M, Boussoffara T, Abdeladhim M, Ben Aleya-Bouafif N, Namane A, et al. Characterization of the antibody response to the saliva of Phlebotomus papatasi in people living in endemic areas of cutaneous leishmaniasis. Am J Trop Med Hyg. 2011;84: 653–61. pmid:21540371
- 36. Clements MF, Gidwani K, Kumar R, Hostomska J, Dinesh DS, Kumar V, et al. Measurement of recent exposure to Phlebotomus argentipes, the vector of Indian visceral Leishmaniasis, by using human antibody responses to sand fly saliva. Am J Trop Med Hyg. 2010;82: 801–7. pmid:20439958
- 37. Teixeira C, Gomes R, Collin N, Reynoso D, Jochim R, Oliveira F, et al. Discovery of markers of exposure specific to bites of Lutzomyia longipalpis, the vector of Leishmania infantum chagasi in Latin America. Milon G, editor. PLoS Negl Trop Dis. 2010;4: e638. pmid:20351786
- 38. Souza AP, Andrade BB, Aquino D, Entringer P, Miranda JC, Alcantara R, et al. Using recombinant proteins from Lutzomyia longipalpis saliva to estimate human vector exposure in visceral Leishmaniasis endemic areas. Milon G, editor. PLoS Negl Trop Dis. 2010;4: e649. pmid:20351785
- 39. Andrade BB, Teixeira CR. Biomarkers for exposure to sand flies bites as tools to aid control of leishmaniasis. Front Immunol. 2012;3: 121. pmid:22661974
- 40. de Moura TR, Oliveira F, Novais FO, Miranda JC, Clarêncio J, Follador I, et al. Enhanced Leishmania braziliensis infection following pre-exposure to sandfly saliva. Jaffe C, editor. PLoS Negl Trop Dis. 2007;1: e84. pmid:18060088
- 41. Abdeladhim M, Ben Ahmed M, Marzouki S, Belhadj Hmida N, Boussoffara T, Belhaj Hamida N, et al. Human cellular immune response to the saliva of Phlebotomus papatasi is mediated by IL-10-producing CD8+ T cells and Th1-polarized CD4+ lymphocytes. Bates PA, editor. PLoS Negl Trop Dis. 2011;5: e1345. pmid:21991402
- 42. Sokal JE. Editorial: Measurement of delayed skin-test responses. N Engl J Med. 1975;293: 501–2. pmid:1152865
- 43. Weigle KA, Valderrama L, Arias AL, Santrich C, Saravia NG. Leishmanin skin test standardization and evaluation of safety, dose, storage, longevity of reaction and sensitization. Am J Trop Med Hyg. 1991;44: 260–271. pmid:2035747
- 44. Alvarado R, Enk C, Jaber K, Schnur L, Frankenburg S. Delayed-type hypersensitivity and lymphocyte proliferation in response to Leishmania major infection in a group of children in Jericho. Trans R Soc Trop Med Hyg. 83: 189–92. Available: http://www.ncbi.nlm.nih.gov/pubmed/2609368 pmid:2609368
- 45. Castés M, Blackwell J, Trujillo D, Formica S, Cabrera M, Zorrilla G, et al. Immune response in healthy volunteers vaccinated with killed leishmanial promastigotes plus BCG. I: Skin-test reactivity, T-cell proliferation and interferon-gamma production. Vaccine. 1994;12: 1041–51. Available: http://www.ncbi.nlm.nih.gov/pubmed/7975845 pmid:7975845
- 46. Khalil EA, El Hassan AM, Zijlstra EE, Mukhtar MM, Ghalib HW, Musa B, et al. Autoclaved Leishmania major vaccine for prevention of visceral leishmaniasis: a randomised, double-blind, BCG-controlled trial in Sudan. Lancet (London, England). 2000;356: 1565–9. Available: http://www.ncbi.nlm.nih.gov/pubmed/11075771
- 47. Armijos RX, Weigel MM, Calvopina M, Hidalgo A, Cevallos W, Correa J. Safety, immunogenecity, and efficacy of an autoclaved Leishmania amazonensis vaccine plus BCG adjuvant against New World cutaneous leishmaniasis. Vaccine. 2004;22: 1320–6. pmid:15003662
- 48. Weigle KA, Santrich C, Martinez F, Valderrama L, Saravia NG. Epidemiology of cutaneous leishmaniasis in Colombia: environmental and behavioral risk factors for infection, clinical manifestations, and pathogenicity. J Infect Dis. 1993;168: 709–14. Available: http://www.ncbi.nlm.nih.gov/pubmed/8354913 pmid:8354913
- 49. Traoré B, Oliveira F, Faye O, Dicko A, Coulibaly CA, Sissoko IM, et al. Prevalence of Cutaneous Leishmaniasis in Districts of High and Low Endemicity in Mali. Boelaert M, editor. PLoS Negl Trop Dis. 2016;10: e0005141. pmid:27898671
- 50. Oliveira F, Doumbia S, Anderson JM, Faye O, Diarra SS, Traoré P, et al. Discrepant prevalence and incidence of Leishmania infection between two neighboring villages in Central Mali based on Leishmanin skin test surveys. Louzir H, editor. PLoS Negl Trop Dis. 2009;3: e565. pmid:20016847
- 51. Moral L, Rubio EM, Moya M. A leishmanin skin test survey in the human population of l’Alacantí region (Spain): implications for the epidemiology of Leishmania infantum infection in southern Europe. Trans R Soc Trop Med Hyg. 96: 129–32. Available: http://www.ncbi.nlm.nih.gov/pubmed/12055798 pmid:12055798
- 52. Gazos-Lopes F, Mesquita RD, Silva-Cardoso L, Senna R, Silveira AB, Jablonka W, et al. Glycoinositolphospholipids from Trypanosomatids Subvert Nitric Oxide Production in Rhodnius prolixus Salivary Glands. Rodrigues MM, editor. PLoS One. 2012;7: e47285. pmid:23077586
- 53. Geraci NS, Mukbel RM, Kemp MT, Wadsworth MN, Lesho E, Stayback GM, et al. Profiling of human acquired immunity against the salivary proteins of Phlebotomus papatasi reveals clusters of differential immunoreactivity. Am J Trop Med Hyg. 2014;90: 923–38. pmid:24615125
- 54. Carvalho a. M, Cristal JR, Muniz a. C, Carvalho LP, Gomes R, Miranda JC, et al. Interleukin 10-Dominant Immune Response and Increased Risk of Cutaneous Leishmaniasis After Natural Exposure to Lutzomyia intermedia Sand Flies. J Infect Dis. 2015; 1–9. pmid:25596303
- 55. Ribeiro JM, Katz O, Pannell LK, Waitumbi J, Warburg A. Salivary glands of the sand fly Phlebotomus papatasi contain pharmacologically active amounts of adenosine and 5’-AMP. J Exp Biol. 1999;202: 1551–9. Available: http://www.ncbi.nlm.nih.gov/pubmed/10229701 pmid:10229701
- 56. Katz O, Waitumbi JN, Zer R, Warburg A. Adenosine, AMP, and protein phosphatase activity in sandfly saliva. Am J Trop Med Hyg. 2000;62: 145–50. Available: http://www.ncbi.nlm.nih.gov/pubmed/10761741 pmid:10761741
- 57. Haskó G, Kuhel DG, Chen JF, Schwarzschild MA, Deitch EA, Mabley JG, et al. Adenosine inhibits IL-12 and TNF-[alpha] production via adenosine A2a receptor-dependent and independent mechanisms. FASEB J. 2000;14: 2065–74. pmid:11023991
- 58. Volf P, Tesarová P, Nohýnkova EN. Salivary proteins and glycoproteins in phlebotomine sandflies of various species, sex and age. Med Vet Entomol. 2000;14: 251–6. Available: http://www.ncbi.nlm.nih.gov/pubmed/11016431 pmid:11016431
- 59. Abdeladhim M, Kamhawi S, Valenzuela JG. What’s behind a sand fly bite? The profound effect of sand fly saliva on host hemostasis, inflammation and immunity. Infect Genet Evol. 2014;28: 691–703. pmid:25117872
- 60. Lestinova T, Rohousova I, Sima M, De Oliveira CI, Volf P. Insights into the sand fly saliva: Blood-feeding and immune interactions between sand flies, hosts, and Leishmania. pmid:28704370
- 61. Lerner EA, Shoemaker CB. Maxadilan. Cloning and functional expression of the gene encoding this potent vasodilator peptide. J Biol Chem. 1992;267: 1062–6. Available: http://www.ncbi.nlm.nih.gov/pubmed/1730635 pmid:1730635
- 62. Wilke CM, Wei S, Wang L, Kryczek I, Kao J, Zou W. Dual biological effects of the cytokines interleukin-10 and interferon-γ. Cancer Immunol Immunother. 2011;60: 1529–41. pmid:21918895
- 63. Rohoušová I, Hostomská J, Vlková M, Kobets T, Lipoldová M, Volf P. The protective effect against Leishmania infection conferred by sand fly bites is limited to short-term exposure. Int J Parasitol. 2011;41: 481–5. pmid:21310158
- 64. Gomes R, Cavalcanti K, Teixeira C, Carvalho AM, Mattos PS, Cristal JR, et al. Immunity to Lutzomyia whitmani Saliva Protects against Experimental Leishmania braziliensis Infection. Engwerda CR, editor. PLoS Negl Trop Dis. 2016;10: e0005078. pmid:27812113
- 65. Sima M, Novotny M, Pravda L, Sumova P, Rohousova I, Volf P. The Diversity of Yellow-Related Proteins in Sand Flies (Diptera: Psychodidae). Traub-Csekö YM, editor. PLoS One. 2016;11: e0166191. pmid:27812196
- 66. Barral A, Honda E, Caldas A, Costa J, Vinhas V, Rowton ED, et al. Human immune response to sand fly salivary gland antigens: a useful epidemiological marker? Am J Trop Med Hyg. 2000;62: 740–5. Available: http://www.ncbi.nlm.nih.gov/pubmed/11304066 pmid:11304066
- 67. Ben Hadj Ahmed S, Kaabi B, Chelbi I, Cherni S, Derbali M, Laouini D, et al. Colonization of Phlebotomus papatasi changes the effect of pre-immunization with saliva from lack of protection towards protection against experimental challenge with Leishmania major and saliva. Parasit Vectors. 2011;4: 126. pmid:21726438
- 68. Xu X, Oliveira F, Chang BW, Collin N, Gomes R, Teixeira C, et al. Structure and function of a "yellow" protein from saliva of the sand fly Lutzomyia longipalpis that confers protective immunity against Leishmania major infection. J Biol Chem. 2011;286: 32383–93. pmid:21795673
- 69. Aquino DMC, Caldas AJM, Miranda JC, Silva AAM, Barral-Netto M, Barral A. Epidemiological study of the association between anti-Lutzomyia longipalpis saliva antibodies and development of delayed-type hypersensitivity to Leishmania antigen. Am J Trop Med Hyg. 2010;83: 825–7. pmid:20889873