Nonhealing Infection despite Th1 Polarization Produced by a Strain of Leishmania major in C57BL/6 Mice
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免疫学杂志 2005年第5期
Abstract
Experimental Leishmania major infection in mice has been of immense interest because it was among the first models to demonstrate the importance of the Th1/Th2 balance to infection outcome in vivo. However, the Th2 polarization that promotes the development of nonhealing cutaneous lesions in BALB/c mice has failed to adequately explain the mechanisms underlying nonhealing forms of leishmaniasis in humans. We have studied a L. major strain from a patient with nonhealing lesions that also produces nonhealing lesions with ulcerations and high parasite burden in conventionally resistant C57BL/6 mice. Surprisingly, these mice develop a strong, polarized, and sustained Th1 response, as evidenced by high levels of IFN- produced by Leishmania-specific cells in the draining lymph node and in the ear lesion, and an absence of IL-4 or IL-13. The parasites fail to be effectively cleared despite high level induction of inducible NO synthase in the lesion, and despite their sensitivity to killing by IFN--activated macrophages in vitro. Infection of IL-10–/– mice, blockade of the IL-10R, or depletion of CD25+ cells during the chronic phase promotes parasite killing, indicating that IL-10 and regulatory T cells play a role in rendering the Th1 responses ineffective at controlling infection in the skin. Mice with nonhealing primary lesions are nonetheless resistant to reinfection in the other ear. We suggest that nonhealing infections in animal models that are explained not by aberrant Th2 development, but by overactivation of homeostatic pathways designed to control inflammation, provide better models to understand nonhealing or reactivation forms of leishmaniasis in humans.
Introduction
Acquired resistance to the intracellular protozoan Leishmania major is dependent on the development of a Th1-type immune response, marked by the induction of CD4+ and CD8+ T cells mediating IFN--dependent macrophage microbicidal activity. In the murine model for cutaneous leishmaniasis, the mechanisms of acquired resistance to L. major have been well documented using genetically resistant C57BL/6 mice, which reproduce the self-cure disease outcome typically seen in humans (1, 2). Intradermal inoculation of these mice with a biologically appropriate low dose of infectious stage parasites produces an initial prepatent phase of intracellular growth, absent of visible lesions or pathology, followed by lesion development and the accumulation of IFN--producing CD4+ and CD8+ T cells in the lesion (3, 4, 5). Following resolution of the lesion, a low number of parasites persists indefinitely in the site, and the host is protected against a rechallenge infection at a distal site (3, 5).
In contrast to the self-limiting infections with L. major observed in C57BL/6 mice, BALB/c mice develop progressive, nonhealing lesions that are associated with an early, sustained, and dominant Th2 response, especially IL-4 (1, 2). Furthermore, ablation of the cells or cytokines involved in normal Th1 development in resistant mice results in Th2 deviation and progressive infection (6, 7, 8, 9). The Th1/Th2 balance that controls immunity to L. major in these murine models has fostered the main conceptual framework for understanding healing and nonhealing forms of clinical disease. However, it has not been possible to clearly associate a Th2 polarity with nonhealing, systemic, or reactivation forms of leishmaniasis in humans. IFN--producing cells or mRNA remain readily detectable in patients with kala-azar, postkala-azar dermal leishmaniasis, or chronic cutaneous leishmaniasis, and the opposing cytokine most commonly found in these clinical settings is not IL-4, but IL-10 (10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21). Interestingly, IL-10 has also been found to contribute to BALB/c susceptibility to L. major (22), although its influence is generally obscured by the Th2 bias that is intrinsic to this mouse strain (23). In contrast, a role for IL-10 in regulating immunity in resistant mice has been conclusively shown, in this case mediating not susceptibility per se, but the persistence of low numbers of parasites in the skin following clinical cure (4). Thus, IL-10–/– mice achieve sterile immunity, as do healed C57BL/6 mice treated during the chronic phase with anti-IL-10 receptor Abs. More recently, IL-10-producing naturally occurring CD4+CD25+ regulatory T cells have been shown to home to L. major lesions, and to be necessary for parasite persistence in healed mice (24). These cells may fulfill a primarily homeostatic function, preventing excessive pathology mediated by Leishmania-specific Th1 cells in the inflammatory site. Although regulatory and effector T cell subsets appear to function in equilibrium to maintain latency in healed mice, these findings raise the possibility that an imbalance in regulatory cells or cytokines might control nonhealing or systemic disease outcomes.
In the present study, we have analyzed the role of these opposing host factors in the immune response of C57BL/6 mice to a strain of L. major that was isolated from a patient with nonhealing lesions (25). The lesions persisted for months following multiple courses of treatment, but the patient was found to have a positive skin test and proliferative responses to Leishmania Ags. In the current study, despite the induction of a strong and polarized Th1 response, the parasites are not effectively controlled, and the dermal lesions fail to resolve. We examine the degree of suppression mediated by IL-10 and by CD4+CD25+ regulatory T cells and argue that these are major contributing factors to the evolution of these nonhealing infections.
Materials and Methods
Mice
C57BL/6 mice were purchased from the Division of Cancer Treatment, National Cancer Institute (Frederick, MD). C57BL/10SgSnAi, C57BL/10SgSnAi-[KO]IL-10, and C57BL/6SgSnAi-[KO]IL-4 mice were purchased from Taconic Farms. All mice were maintained in the National Institute of Allergy and Infectious Diseases animal care facility under specific pathogen-free conditions.
Antibodies
For in vivo Ab treatments, mice were given 1.0 mg i.p. injections at the indicated times with anti-CD25 (PC61), anti-IL-10R (1B1.3a), or an isotype control (GL113). CD25 depletion was determined by staining cells with Ab clone 7D4, which showed depletion to be >80% compared with control-treated mice. The following fluorochrome-conjugated Abs for cell surface and intracytoplasmic staining were purchased from BD Pharmingen: PE-anti-CD25 (PC61), FITC-anti-CD25 (clone 7D4), FITC-anti-TCR- (H57-597), PE-Cy5 anti-CD4 (L3T4), PE-Cy5 anti-CD8 (53–6.7), and PE anti-IFN- (XMG1.2). The isotype controls used (all from BD Pharmingen) were rat IgG2b (A95-1), rat IgG2a (R35-95), rat IgG1 (R3-34), and hamster IgG group 2 (Ha4/8). For ELISA measurements of cytokines, the following Ab pairs were purchased from eBioscience. IL-4: 11B11, BVD6-24G2; IL-10: JES5-16E3, JES5-2A5; IFN-: XMG1.2, R4-6A2.
Parasite preparation and intradermal inoculation
L. major clones V1 (MHOM/IL/80/Friedlin) and Sd (MHOM/SN/74/SD) were cultured in medium 199 with 20% heat-inactivated FCS (Gemini Bio-Products), 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mM L-glutamine, 40 mM HEPES, 0.1 mM adenine (in 50 mM HEPES), 5 mg/ml hemin (in 50% triethanolamine), and 1 mg/ml 6-biotin (M199/S). The Sd strain was isolated from a patient who was admitted to the National Institutes of Health for treatment of lesions that persisted following multiple courses of therapy (25). Infective-stage metacyclic promastigotes of L. major were isolated from stationary cultures (4–5 days old) by density gradient centrifugation as described previously (26). Metacyclic promastigotes (1000) were inoculated into the ear dermis using a 30-gauge needle in a volume of 5 μl. The development of the lesion was monitored by measuring the diameter of the ear lesion with a direct-reading vernier caliper (Thomas Scientific).
Processing of ear tissue and estimation of parasite load
Parasite titrations were performed as previously described (3). The two sheets of infected ears were separated, deposited in DMEM containing 100 U/ml penicillin, 100 μg/ml streptomycin, and 0.2 mg/ml Liberase CI purified enzyme blend (Roche Diagnostic Systems), and incubated for 2 h at 37°C. Sheets were then placed in a grinder and processed in a Medimachine tissue homogenizer (BD Biosciences) for 4 min. Tissue homogenates were then filtered through a 70-μm cell strainer (Falcon Products), and aliquots of these were serially diluted in 96-well flat-bottom microtiter plates containing biphasic medium prepared using 50 μl of NNN medium containing 30% of defibrinated rabbit blood overlaid with 50 μl of M199/S. The number of viable parasites in each sample was determined from the highest dilution at which promastigotes could be grown out after 7 days of incubation at 25°C.
Analysis of intradermal lymphocytes, APC preparation, and cytokine analysis
To characterize leukocytes in the inoculation sites, the ears were collected, and the ventral and dorsal dermal sheets were prepared as described above. For cell surface phenotype, cells were fixed in PBS/2% paraformaldehyde for 15 min at 25°C, washed, and resuspended in staining buffer (PBS with 0.1% BSA and 0.01% NaN3) for phenotypic analysis by flow cytometry. For in vitro restimulation, unfractionated lymph node (LN)3 cells or dermal cells were incubated with uninfected or amastigote-infected bone marrow-derived dendritic cells (BMDCs) as a source of APCs. BMDCs were generated from bone marrow in the presence of GM-CSF as previously described (27). LN cells (3 x 106) or dermal cells pooled from six ears were resuspended in RPMI 1640 containing 10% FCS, 10 mM HEPES, 100 U/ml penicillin, and 100 μg/ml streptomycin, and 1 ml of the cell suspension was incubated with uninfected or amastigote-infected BMDCs at a ratio of 5:1 in 24-well plates at 37°C in 5% CO2 for 18 h, with brefeldin A added during the last 5 h. The cells were then fixed and analyzed for surface markers and intracellular IFN-. In some experiments, unfractionated LN or dermal cells were incubated for 48 h with BMDCs or amastigote-infected BMDCs, and IFN-, IL-10, and IL-4 from culture supernatants were quantitated by ELISA (eBioscience) according to the manufacturer’s protocol. The limits of detection for the cytokines were 4 pg/ml for IL-4, 15 pg/ml for IL-10, and 15 pg/ml for IFN-.
Immunolabeling and flow cytometry
Before staining, LN or dermal cells were incubated with an anti-FcIIIR/IIR (clone 2.4G2; BD Pharmingen) mAb in PBS containing 0.1% BSA and 0.01% NaN3. The staining of surface markers and intracellular cytokines was performed sequentially. The cells were stained first for their surface markers (TCR chain receptor, CD8, or CD4), followed by a permeabilization step and staining with anti-IFN-. The lymphocytes were identified by characteristic size (forward light scatter) and granularity (side light scatter), in combination with anti-TCR- chain and anti-CD4 or anti-CD8, or anti-CD25 for surface staining. For each sample, 150,000 events were acquired for analysis. The data were collected and analyzed using CellQuest Pro software and a FACSCalibur flow cytometer (BD Biosciences).
Real-time PCR
For analysis of gene expression, ears were removed and immediately placed in RNAlater stabilization buffer (Qiagen) to prevent degradation. Tissue was then disrupted by placing the ears in mortars containing liquid nitrogen and grinding with a pestle. The disrupted tissue was then homogenized using Qiashredder homogenizers and processed for RNA isolation using the RNeasy minikit (Qiagen), following the manufacturer’s protocol, with all samples being treated with a DNase step to remove genomic DNA. Reverse transcription was performed on 500 ng of total RNA from each ear sample using Superscript III First-Strand synthesis system for RT-PCR (Invitrogen Life Technologies). Real-time PCR was performed on an ABI Prism 7900HT sequence detection system (Applied Biosystems) using 10% of the cDNA reaction in a total volume of 20 μl of PCR mixture. FAM-MGB-labeled primer/probe sets for IFN-, IL-10, IL-13, inducible NO synthase-2 (iNOS-2), and Foxp3 were designed from Applied Biosystems, and VIC-MGB-labeled 18 S rRNA was used as the endogenous control. The relative quantitation of products was determined by the comparative threshold cycle method using the equation 2–cT to determine the fold increase in product. Each gene of interest was normalized to the 18 S rRNA endogenous control, and the fold change in expression was displayed as relative to naive controls. Four ears were used for each time point, and each sample was run in triplicate during the PCR.
In vitro killing assay
Peritoneal macrophages from C57BL/6 mice were plated in eight-chamber Lab-Tek Permanox tissue culture slides (Miles Laboratories) at 5 x 104 cells/well in a volume of 0.4 ml of RPMI 1640 complete. V1 or Sd tissue-derived amastigotes were added to the wells at a ratio of 5:1, and infection was allowed to establish for 72 h at 35°C. At this time, 0.1 μg/ml LPS and titrating amounts of recombinant mouse IFN- were added. After an additional 72 h, the slides were fixed for 1 min in anhydrous methanol and stained with Diff-Quick solutions. The numbers of infected and non-infected macrophages and the number of amastigotes were counted. The percentage of killing relative to untreated macrophages was calculated on the basis of the comparison of total amastigotes per 100 macrophages.
Statistical analysis
To determine whether differences were statistically significant, Student’s t test was performed, using a two-tailed distribution with unpaired samples.
Results
L. major Sd produces a nonhealing phenotype in C57BL/6 mice
To characterize the host response to the L. major Sd strain that produced nonhealing lesions in a patient, C57BL/6 mice were inoculated with 1000 metacyclic promastigotes, and monitored for lesion development in reference to the L. major V1 strain. As previously described (3), mice infected intradermally with V1 parasites produced lesions that healed spontaneously in 8–12 wk following inoculation (Fig. 1). In contrast, when mice were infected with the L. major Sd strain, an initial prepatent phase was followed by the development of lesions that failed to heal. Lesions grew slowly, but progressively larger in diameter, resulting in ulcerations and eventual loss of dermal tissue (Fig. 1). The lesions did not disseminate to other dermal sites nor was there evidence of visceralization beyond the local draining LN (DLN) (data not shown). Lesions harboring the V1 strain showed a peak in parasite number at 6 wk, which by 10 wk had been 90% cleared from the site, and by 22 wk fewer than 100 parasites remained in the healed lesion. In contrast, lesions harboring the Sd strain had a slow progressive increase in the number of parasites throughout the 22 wk that lesions were monitored.
FIGURE 1. Sd strain of L. major produces a nonhealing phenotype in C57BL/6 mice. A total of 103 V1 or Sd metacyclic promastigotes were injected in the ear dermis of mice. A, Development of ear lesions over the course of 22 wk. The mean and SD of six ears per group are shown. B, Parasite loads in ear lesions, showing the mean and SD from six ears per group. C, Photographs of the ears were taken 22 wk postinfection The results are representative of at least three independent experiments.
Priming and recruitment of Th1 cells is not deficient in L. major Sd-infected mice
Because the L. major Sd strain produced a nonhealing phenotype, we investigated the possibility that this L. major substrain induces an aberrant Th2 response in this normally resistant host. As early as 3 wk postinfection, Ag-specific IFN- production was detectable in high levels in the DLN of the Sd-infected mice, at a time when Th1 priming was barely in evidence in the mice infected with V1 (Table I). IFN- production in the Sd-infected mice was maintained at high levels at 6 and 11 wk, whereas IFN- production peaked at around 6 wk in the V1-infected mice and declined significantly by 11 wk, at a time when the majority of parasites had been eliminated. Importantly, at no time point did the DLN cells from the Sd or V1 mice produce significant amounts of IL-4. In contrast, IL-10 was detected by 3 wk in the Sd mice, with high levels being produced by 6 wk postinfection. Apart from the early time point, IL-10 was secreted in even higher levels by the DLN cells from the V1-infected mice.
Table I. ELISA cytokine measurements of draining lymph node cellsa
The number of CD4+ and CD8+ T cells in the ear dermis was analyzed to determine whether the Sd infection was inhibiting the recruitment of effector cells to the site of infection. Fig. 2 shows the frequency of TCR-+CD4+ and TCR-+CD8+ cells present in the lesions at 3, 6, and 10 wk postinfection, along with the total numbers of these subsets. The recruitment of CD4+ T cells was apparent in both groups by 3 wk, though it was slightly higher in the Sd lesions at 3 and 6 wk, and 2-fold higher at 10 wk, when the V1 dermal lesions were beginning to resolve. Recruitment of CD8+ T cells was generally comparable in both groups at the earlier time points, with a slight decrease of CD8+ cells seen in the Sd infections compared with V1 at 10 wk.
FIGURE 2. Sd infection is not defective in the recruitment of lymphocytes to the lesion. At various times postinfection, dermal cells were immunolabeled for TCR- and CD4 or CD8 and analyzed by flow cytometry. Lymphocytes were gated according to forward scatter and side scatter properties. The numbers shown in the upper right quadrant are the absolute numbers of cells per ear (x104) and are the results of four to six ears pooled per time point. The numbers represent the mean and SD of three independent experiments.
The ability of the lesional CD4+ T cells to secrete cytokines in response to infected DC was measured by intracellular staining and by ELISA. Ag-specific IFN--producing cells were detected in both groups as early as 3 wk, increasing in absolute numbers at wk 6, and maintained at high frequency at 10 wk. Strikingly, the Sd infection contained more than twice the number of Th1 effectors at 10 wk, suggesting the higher parasite load at this time was driving the continued expansion and recruitment of effector cells (Fig. 3). Cytokines were also measured by ELISA (Table II). An earlier, more elevated and sustained Ag-specific IFN- response by Sd lesional cells was also noted by ELISA. IL-4-producing cells were not detectable in either group, either by intracellular staining (data not shown), or by ELISA, except for the slightly elevated levels produced by the V1 lesional cells at wk 6 (Table II). In contrast, substantial amounts of IL-10 were produced by ear lesional cells from both groups by 3 wk, and high leve
ls continued to be produced by the V1 and Sd dermal cells throughout the healing and chronic phase, respectively. Interestingly, the majority of the IL-10 produced by the dermal cells did not require Ag re-stimulation for release. Taken together, these results indicate that there is not a defect in Th1 priming, nor a defect in recruitment of Th1 cells to the site of the infection. The Sd parasites persist and produce nonhealing lesions despite the development of a polarized Th1 response.
FIGURE 3. Sd-infected ears contain high numbers of IFN- secreting effector cells. Dermal cells from six V1- or Sd-infected ears were pooled and restimulated for 18 h in the presence of BMDCs or amastigote-infected BMDCs (BMDCs/Lm). Brefeldin A was added during the last 5 h, and cells were then fixed and permeabilized for intracellular cytokine staining. The numbers in the upper right quadrant are the percentages of CD4+ cells secreting IFN-. The data are representative of at least three independent experiments.
Table II. ELISA cytokine measurements of cells from ear lesionsa
To determine whether the Sd strain might persist in the lesion due to an intrinsic resistance to NO-dependent killing by activated macrophages, we performed an in vitro killing assay, in which infected macrophages were activated with LPS and titrated amounts of rIFN-, providing both the priming and triggering stimuli needed to generate optimal levels of NO (Fig. 4). Both strains were efficiently killed when the infected macrophages were activated using 10 ng/ml rIFN-, and although a broad dose range defined the LD50 for each of the strains, the Sd strain did not display a greater resistance to killing at any of the concentrations of rIFN- used. Thus the resistance of the Sd strain to killing in vivo is apt to reflect differences in the activation state of the infected cells in the inflammatory site.
FIGURE 4. V1 and Sd strains are equally susceptible to killing by activated macrophages in vitro. Peritoneal macrophages were plated in 8-well chamber slides and infected with V1 or Sd amastigotes. Macrophages were activated by the addition of LPS and titrated amounts of rIFN-, 72 h after infection. After an additional 72 h, macrophages were fixed and stained for parasite quantitation. The percentage of killing was determined by the number of amastigotes per 100 macrophages and was based on the comparison with unactivated macrophages.
The role of IL-10 and CD4+CD25+ Treg cells in the development of nonhealing lesions due to L. major Sd
Because the presence of IL-10 has been shown to permit the persistence of L. major in the face of a robust Th1 response in healed mice (4), we considered the possibility that the nonhealing phenotype might be due to a modest shift in the balance between IFN- and IL-10 cytokines, not revealed in the ELISA assays of Ag re-stimulated dermal cells. As IL-10 has been shown to be produced by keratinocytes in dermal lesions (13), and by Leishmania-infected macrophages in vitro (22, 28), real-time RT-PCR on RNA extracted directly from the ears was used to better quantitate total IL-10 produced by the different possible sources of IL-10 in the inflammatory site. Compared with V1, the Sd strain activated elevated levels of IFN- mRNA, in agreement with the flow cytometry and ELISA data, but also contained significantly increased message for IL-10 at 11 and 14 wk postinfection (Fig. 5). The difference seen at these time points, especially at wk 14, is apt to be secondary to the inflammatory conditions associated with active vs healed lesions. No difference was observed at the 6-wk time point, around the time when the clinical outcomes were beginning to diverge. Additionally, IL-13 was measured by real-time PCR to exclude the possible role of this Th2-associated cytokine. At both 6 and 12 wk postinfection, neither group expressed significant IL-13 messages above that detectable in naive ears. The data reinforce the absence of a Th2 immune deviation in the L. major Sd-infected ears.
FIGURE 5. Sd lesions contain elevated levels of mRNA for IFN-, IL-10, and Foxp3, and low amounts of IL-13. Total RNA was isolated directly from the ears at the indicated times during the infection, reverse transcribed, and analyzed by real-time PCR. The target genes were normalized to the endogenous control, and the values shown are the fold increase relative to expression in naive ears. Each bar is the geometric mean and SD of four individual ears. (*, p < 0.05; **, p < 0.01).
A potential source of IL-10 in the nonhealing lesions is CD4+CD25+ regulatory T cells. The presence of these cells has been shown to control the persistence of L. major in healed mice (24), and adoptive transfer of these cells, or their Ag-induced expansion, has been recently shown to reactivate parasite growth and dermal pathology in healed lesions (29). Thus it was important to determine whether there was an increased frequency of CD24+CD25+ Treg in the Sd mice. The forkhead/winged helix transcription factor Foxp3 has been shown to be specifically expressed by CD25+ Treg, as well as by CD25– T cells with regulatory activity (30, 31). Compared with V1, the Sd lesions contained a significantly increased level of Foxp3 mRNA at 11 and 14 wk postinfection and a small, though nonsignificant, increase at 6 wk (Fig. 5). And whereas the number of Foxp3 transcripts remained steady throughout the acute, healing, and latent stages of V1 infection, there was a steady increase during progressive lesion development in the Sd non-cure mice. Flow cytometry analysis to measure the frequency of CD4+CD25+ and CD4+CD25– T cells in the lesions revealed no major differences between the two infections during the first 11 wk (Fig. 6). There was a modest increase in the total number of CD25+ Treg in the Sd lesions at 6 wk, and especially at 11 wk when there was an 3-fold increase compared with V1, consistent with the Foxp3 real-time PCR data.
FIGURE 6. V1- and Sd-infected ears contain high numbers of CD4+CD25+ cells. Dermal cells were removed from ear lesions at the indicated times and stained for CD4 and CD25. The numbers in the upper right quadrant represent the percentage of total CD4+ cells that express CD25 and the absolute numbers of CD4+CD25+ cells (x104) per ear. The data are from one representative experiment of at least three independent experiments.
To determine what role IL-10 may play in prevention of healing, we first took a genetic approach by infecting IL-10–/– mice with either V1 or Sd (Fig. 7). At 10 wk postinfection, there was a 2-log reduction in L. major Sd parasite burden in the IL-10–/– mice compared with the wild-type B/10 mice (Fig. 7B). However, sterile cure of the Sd strain was not achieved as was apparent with V1, consistent with previous observations (4). Despite the reduced parasite load, the Sd lesions in the IL-10–/– mice were larger than in the wild-type mice throughout the course of infection, consistent with a critical role of IL-10 in modulating immune-mediated pathology (Fig. 7A). In contrast to the IL-10–/– mice, the IL-4–/– mice were found to be equally susceptible to L. major Sd as was wild-type B/6 mice, both in terms of the severity of their nonhealing lesions (not shown) and the failure to effectively reduce the parasite burden during the chronic stage (Fig. 7B). To determine the effect of a transient block in IL-10 signaling, mice were treated with anti-IL-10R Ab during the chronic phase of the infection. Mice were sacrificed 1 wk after the course of treatment, and the parasite burden was measured (Fig. 7C). The treatment had a dramatic curative effect in the Sd-infected mice, with 99% of parasites being cleared from the site. The treatment also reduced the parasite burden in the healing V1 lesions, as previously reported. In neither infection did the treatment result in sterile cure, though it is possible that more prolonged treatment, or treatment during more latent stage of infection with V1, might have produced the complete clearance of V1 achieved in the prior studies. The more effective clearance of parasites following anti-IL-10R treatment was associated in both groups with increased IFN- and iNOS expression in the lesion, as measured by real-time PCR. However, it should be noted that the increased iNOS expression in the L. major Sd lesions was slight, and perhaps more importantly, the levels in the untreated Sd-infected mice were already high (Fig. 8). Sd-infected mice were also depleted of CD25+ cells during the chronic stage of the infection by treating with anti-CD25 Ab over a course of 2 wk, and analyzing the parasite load and immune response 1 wk following the last treatment. The anti-CD25-treated mice had a significant, 10-fold reduction in parasite number compared with the isotype controls (Fig. 9), and slight increases in both IFN- and iNOS-2 expression in the lesion, though again the levels of both transcripts were already high in the untreated mice.
FIGURE 7. IL-10 deficiency or anti-CD25 treatment promotes parasite killing in the Sd-infected mice. Parasite loads in ear lesions in C57BL/10 or IL-10–/– mice or C57BL/6 or IL-4–/– mice infected with 103 metacyclics and sacrificed at wk 10 for parasite enumeration (A), C57BL/6 mice treated with anti-IL-10R Ab every 4 days during the last 2 wk of infection and sacrificed at wk 10 for parasite enumeration (B), and C57BL/6 mice treated with anti-CD25 Ab every 4 days during the last 2 wk of infection and sacrificed at wk 12 for parasite enumeration (C). The data are representative of two independent experiments (*, p < 0.05 compared with control mice).
FIGURE 8. Anti-IL-10R- and anti-CD25-treated mice have increased expression of IFN- and iNOS-2 mRNA. At 12 wk postinfection, mice were given two i.p. injections of Ab 3 days apart. Three days after the second treatment, total RNA was isolated from ears, reverse transcribed, and analyzed by real-time PCR. The target genes were normalized to the endogenous control, and the values shown are the fold increase relative to expression in naive mice. Each bar is the geometric mean and SD of four individual ears (*, p < 0.05 compared with control mice).
FIGURE 9. Sd-infected mice develop protective immunity to a rechallenge. Fourteen weeks following primary infection, mice were rechallenged in the opposite ear with 103 metacyclics, along with naive animals as a comparison. A, Ear lesion development in rechallenged and naive mice. The mean and SD of five mice per group are shown. B, Parasite burdens in individual ears in the primary site and secondary rechallenge site 8 wk after rechallenge.
Mice with nonhealing L. major Sd infection develop concomitant immunity to reinfection
Mice that developed nonhealing L. major Sd lesions at 14 wk were rechallenged in the contralateral ear with the Sd strain to determine whether protective immunity against reinfection had been generated during the primary infection. Despite the inability to heal the primary lesion, the rechallenge ears did not develop measurable pathology, and 8 wk after rechallenge they had a 10-fold reduction in parasite burden compared with naive-infected controls (Fig. 9). Coinfection with the Sd and V1 strains using 1000 metacyclics in each ear did not compromise the ability of the mice to control and resolve the V1 lesions, again despite development of nonhealing Sd lesions in the contralateral ear (data not shown).
Discussion
Using low dose intradermal inoculation in conventionally resistant C57BL/6 mice, we have characterized the host immune response to a strain of L. major that produces a nonhealing phenotype. This occurs despite the development of a strong and polarized Th1 response, as measured by IFN- production in the DLN as well as in the site of the infection, and the correspondingly low levels of IL-4 and IL-13 produced in these sites. In addition, the nonhealing phenotype developed despite normal recruitment of CD4+ and CD8+ lymphocytes, and despite high level induction of iNOS in the lesion, conditions that are collectively associated with effective control of L. major infection in the skin (2). The possibility that the Sd strain is intrinsically more resistant to IFN--mediated killing was addressed by determining the concentrations of IFN- required to activate peritoneal macrophages for leishmanicidal capacity in vitro, and no significant difference compared with the V1 strain was observed. The non-cure phenotype seen in this model is clearly divergent from the BALB/c model of susceptibility, in which the infection is marked by a strong Th2 bias. The disease-promoting cytokine that could be identified in this model is not IL-4, but IL-10, as indicated by the enhanced host resistance observed in Sd-infected IL-10–/– B10 mice, or in B6 mice treated with anti-IL-10R Ab. CD4+CD25+ regulatory T cells were also shown to play a role, as the in vivo depletion of CD25+ cells promoted greater clearance of parasites from the skin. It is important to consider that the L. major BALB/c model has in general failed to adequately explain severe forms of clinical disease, which seem not to be associated with a Th1 response defect per se, but with concomitant expression of IL-10 (10, 11, 13, 14, 15, 16, 17, 18, 19, 20, 21).
Although IL-10 and CD4+CD25+ T cells are induced by L. major Sd and are required for the evolution of the nonhealing lesions, these responses also accompany the dermal infections with V1 that produce a healing phenotype in B6 mice. In this setting, IL-10 produced by naturally occurring regulatory T cells has been shown to prevent sterile cure, and the regulatory cells seem to function in equilibrium with Leishmania-specific effectors to establish and maintain latent infection in the skin (24). Might the inability to heal the primary Sd lesion be the result of an imbalance in suppressor cells or cytokines in the inflammatory site? While IL-10 production and both IL-10 and Foxp3 mRNA were maintained at elevated levels in the nonhealing compared with the healing L. major lesions, at the critical earlier stage of infection, around wk 6, when the V1 parasites began to be effectively controlled, the inability of an equally potent Th1 response to clear the Sd parasites was not associated with increased IL-10 production or an increased frequency of CD4+CD25+ T cells in the site. In fact, the most striking difference between the immune response induced by these two strains is the high level IFN- production by cells from the DLN or ear lesions in the Sd-infected mice as early as 3 wk postchallenge, at a time when the Th1 response had yet to develop in the mice infected with V1. How this paradoxical finding relates to the evolution of the non-cure phenotype is not known. It is of course possible that additional suppressor factors other than IL-10 might accompany the early, elevated IFN- response in the Sd mice to shift the balance in favor of parasite survival. One indication that the L. major Sd strain induces factors in addition to IL-10 that promote infection is the observation that a low level infection was maintained in the IL-10–/– mice, whereas complete cure was achieved in the IL-10–/– mice infected with V1, confirming our prior studies (4).
Although IL-10 production may not be a sufficient condition for the evolution the non-cure phenotype, it nonetheless contributes in an essential way to this outcome, as indicated by the more effective clearance of the Sd strain in the IL-10-deficient and anti-IL-10R-treated mice. The source of the IL-10 induced by Sd infection is not clear from these studies. Depletion of the CD25+ cells did not appreciably reduce the amount of IL-10 produced in the site (data not shown). Furthermore, the IL-10R blockade consistently promoted greater parasite clearance than did the CD25+ cell depletion, suggesting alternative sources of IL-10. It has been shown that amastigote-infected macrophages produce IL-10 (28, 32), as do macrophages following FcR ligation (22), which in this context might involve anti-Leishmania Abs and released Ags. Keratinocytes are also known to produce IL-10 (13, 33), and their ability to do so in visceral leishmaniasis patients is associated with reactivation disease (postkala azar dermal leishmaniasis) (13). It is interesting that in contrast to the IFN- response, the IL-10 produced by the lesional cells was consistently at high levels in the absence of Ag restimulation, suggesting that non-Ag-specific sources of IL-10 are present in the inflammatory site. It is possible that differences in the frequency or activation state of these IL-10-producing cells, obscured by the quantitation of total IL-10, represent the critical difference between the V1 and Sd inflammatory sites. Autocrine IL-10 produced by infected macrophages is apt, for example, to be far more effective at deactivating infected cells than IL-10 produced by Treg. And although the quantitation of total iNOS in the lesion indicates a highly activated tissue environment, the deactivated tissue might be confined to the infected macrophages or other parasitized cells.
The nonhealing infections produced by L. major Sd in B6 mice mimics in some respects the pattern of susceptibility displayed by "resistant" mice infected with new world cutaneous Leishmania amazonensis strains and their related subspecies Leishmania mexicana (34, 35). In fact, the L. amazonensis studies in resistant mice (C3H and C57BL/6) provided an early challenge to the role of IL-4 and Th2 polarization as a necessary condition for the evolution of nonhealing disseminating forms of leishmaniasis, as L. amazonensis-infected IL-4 knockout mice and anti-IL-4-treated mice still developed nonhealing lesions (35). In contrast, results from two studies involving IL-10 knockout mice indicated a role for IL-10 in promoting parasite growth, since 1–2 log reductions were observed in the deficient mice (36, 37). A critical difference between these non-cure phenotypes and the non-cure Sd infections reported here is that whereas the new world strains induced only low level IFN- responses throughout infection, Sd elicits a strong sustained Th1 response.
We observed in our studies of L. major Sd infection in B6 mice that despite their inability to heal their primary lesion, when the animals were rechallenged in the contralateral ear, they were substantially protected. Thus the IL-10, Treg, and possibly other suppressive mechanisms that function within the primary inflammatory site to prevent effective clearance by Th1 cells do not inhibit the expression of Th1 effector activity in a naive re-challenge site. We have argued that the activation of Treg during infection provides a benefit to the host by not only controlling the severity of inflammation, but by promoting parasite persistence the Treg will indirectly maintain a memory pool necessary for resistance to reinfection (24). Since Treg may be preferentially recruited to or expanded by signals that accumulate within inflamed tissue, they may not be able to home to naive rechallenge sites in sufficient time or numbers to suppress the expression of concomitant immunity.
The availability of a non-cure phenotype of L. major infection associated not with aberrant Th2 development but with ongoing Th1 responses rendered ineffective by an imbalance in homeostatic suppressive cells or cytokines may better reflect the mechanisms controlling non-curing forms of cutaneous visceral leishmaniasis in humans.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 Current address: Department of Microbiology and Tropical Medicine, George Washington University, Washington, DC 20037.
2 Address correspondence and reprint requests to Dr. David L. Sacks, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Laboratory of Parasitic Diseases, Building 4, Room 126, 4 Center Drive MSC 0425, Bethesda, MD 20892-0425. E-mail address: dsacks{at}nih.gov
3 Abbreviations used in this paper: LN, lymph node; BMDC, bone marrow-derived dendritic cell; iNOS, inducible NO synthase; DLN, draining LN.
Received for publication September 8, 2004. Accepted for publication December 16, 2004.
References
Reiner, S. L., R. M. Locksley. 1995. The regulation of immunity to Leishmania major. Annu. Rev. Immunol. 13:151.
Sacks, D., N. Noben-Trauth. 2002. The immunology of susceptibility and resistance to Leishmania major in mice. Nat. Rev. Immunol. 2:845.
Belkaid, Y., S. Mendez, R. Lira, N. Kadambi, G. Milon, D. Sacks. 2000. A natural model of Leishmania major infection reveals a prolonged "silent" phase of parasite amplification in the skin before the onset of lesion formation and immunity. J. Immunol. 165:969.
Belkaid, Y., K. F. Hoffmann, S. Mendez, S. Kamhawi, M. C. Udey, T. A. Wynn, D. L. Sacks. 2001. The role of interleukin (IL)-10 in the persistence of Leishmania major in the skin after healing and the therapeutic potential of anti-IL-10 receptor antibody for sterile cure. J. Exp. Med. 194:1497.
Belkaid, Y., E. Von Stebut, S. Mendez, R. Lira, E. Caler, S. Bertholet, M. C. Udey, D. Sacks. 2002. CD8+ T cells are required for primary immunity in C57BL/6 mice following low-dose, intradermal challenge with Leishmania major. J. Immunol. 168:3992.
Belosevic, M., D. S. Finbloom, P. H. Van Der Meide, M. V. Slayter, C. A. Nacy. 1989. Administration of monoclonal anti-IFN- antibodies in vivo abrogates natural resistance of C3H/HeN mice to infection with Leishmania major. J. Immunol. 143:266.
Ghalib, H. W., M. R. Piuvezam, Y. A. Skeiky, M. Siddig, F. A. Hashim, A. M. el-Hassan, D. M. Russo, S. G. Reed. 1993. Interleukin 10 production correlates with pathology in human Leishmania donovani infections. J. Clin. Invest. 92:324.
Karp, C. L., S. H. el-Safi, T. A. Wynn, M. M. Satti, A. M. Kordofani, F. A. Hashim, M. Hag-Ali, F. A. Neva, T. B. Nutman, D. L. Sacks. 1993. In vivo cytokine profiles in patients with kala-azar: marked elevation of both interleukin-10 and interferon-. J. Clin. Invest. 91:1644.
Kenney, R. T., D. L. Sacks, A. A. Gam, H. W. Murray, S. Sundar. 1998. Splenic cytokine responses in Indian kala-azar before and after treatment. J. Infect. Dis. 177:815.
Gasim, S., A. M. Elhassan, E. A. Khalil, A. Ismail, A. M. Kadaru, A. Kharazmi, T. G. Theander. 1998. High levels of plasma IL-10 and expression of IL-10 by keratinocytes during visceral leishmaniasis predict subsequent development of post-kala-azar dermal leishmaniasis. [Published erratum appears in 1998 Clin. Exp. Immunol. 112:547.
Ribeiro-de-Jesus, A., R. P. Almeida, H. Lessa, O. Bacellar, E. M. Carvalho. 1998. Cytokine profile and pathology in human leishmaniasis. Braz. J. Med. Biol. Res. 31:143.
Holaday, B. J., M. M. Pompeu, S. Jeronimo, M. J. Texeira, A. Sousa Ade, A. W. Vasconcelos, R. D. Pearson, J. S. Abrams, R. M. Locksley. 1993. Potential role for interleukin-10 in the immunosuppression associated with kala azar. J. Clin. Invest. 92:2626
Ismail, A., A. M. El Hassan, K. Kemp, S. Gasim, A. E. Kadaru, T. Moller, A. Kharazmi, T. G. Theander. 1999. Immunopathology of post kala-azar dermal leishmaniasis (PKDL): T-cell phenotypes and cytokine profile. J. Pathol. 189:615.
Gasim, S., A. M. Elhassan, A. Kharazmi, E. A. Khalil, A. Ismail, T. G. Theander. 2000. The development of post-kala-azar dermal leishmaniasis (PKDL) is associated with acquisition of Leishmania reactivity by peripheral blood mononuclear cells (PBMC). Clin. Exp. Immunol. 119:523.
Ajdary, S., M. H. Alimohammadian, M. B. Eslami, K. Kemp, A. Kharazmi. 2000. Comparison of the immune profile of nonhealing cutaneous Leishmaniasis patients with those with active lesions and those who have recovered from infection. Infect. Immun. 68:1760.
Louzir, H., P. C. Melby, A. Ben Salah, H. Marrakchi, K. Aoun, R. Ben Ismail, K. Dellagi. 1998. Immunologic determinants of disease evolution in localized cutaneous leishmaniasis due to Leishmania major. J. Infect. Dis. 177:1687.
Akuffo, H., K. Maasho, M. Blostedt, B. Hojeberg, S. Britton, M. Bakhiet. 1997. Leishmania aethiopica derived from diffuse leishmaniasis patients preferentially induce mRNA for interleukin-10 while those from localized leishmaniasis patients induce interferon-. J. Infect. Dis. 175:737.
Melby, P. C., F. J. Andrade-Narvaez, B. J. Darnell, G. Valencia-Pacheco, V. V. Tryon, A. Palomo-Cetina. 1994. Increased expression of proinflammatory cytokines in chronic lesions of human cutaneous leishmaniasis. Infect. Immun. 62:837.
Kane, M. M., D. M. Mosser. 2001. The role of IL-10 in promoting disease progression in leishmaniasis. J. Immunol. 166:1141.
Noben-Trauth, N., R. Lira, H. Nagase, W. E. Paul, D. L. Sacks. 2003. The relative contribution of IL-4 receptor signaling and IL-10 to susceptibility to Leishmania major. J. Immunol. 170:5152.
Lutz, M. B., N. Kukutsch, A. L. Ogilvie, S. Rossner, F. Koch, N. Romani, G. Schuler. 1999. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J. Immunol. Methods 223:77.
Carrera, L., R. T. Gazzinelli, R. Badolato, S. Hieny, W. Muller, R. Kuhn, D. L. Sacks. 1996. Leishmania promastigotes selectively inhibit interleukin 12 induction in bone marrow-derived macrophages from susceptible and resistant mice. J. Exp. Med. 183:515.
Mendez, S., S. K. Reckling, C. A. Piccirillo, D. Sacks, Y. Belkaid. 2004. Role for CD4+CD25+ regulatory T cells in reactivation of persistent leishmaniasis and control of concomitant immunity. J. Exp. Med. 200:201.
Fontenot, J. D., M. A. Gavin, A. Y. Rudensky. 2003. Foxp3 programs the development and function of CD4+CD25+ regulatory T cells. Nat. Immunol. 4:330.
Hori, S., T. Nomura, S. Sakaguchi. 2003. Control of regulatory T cell development by the transcription factor Foxp3. Science 299:1057.
Guizani-Tabbane, L., K. Ben-Aissa, M. Belghith, A. Sassi, K. Dellagi. 2004. Leishmania major amastigotes induce p50/c-Rel NF-B transcription factor in human macrophages: involvement in cytokine synthesis. Infect. Immun. 72:2582.
Moore, K. W., R. de Waal Malefyt, R. L. Coffman, A. O’Garra. 2001. Interleukin-10 and the interleukin-10 receptor. Annu. Rev. Immunol. 19:683.
Roberts, M., J. Alexander, J. Blackwell. 1990. Genetic analysis of Leishmania mexicana infection in mice single gene Scl-2 controlled predisposition to cutaneous lesion development. J. Immunogenet. 17:89
Afonso, L., P. Scott. 1993. Immune responses associated with susceptibility of C57BL/10 mice to Leishmania amazonensis. Infect. Immun. 61:2952.
Padigel, U. M., J. Alexander, J. P. Farrell. 2003. The role of interleukin-10 in susceptibility of BALB/c mice to infection with Leishmania mexicana and Leishmania amazonensis. J. Immunol. 171:3705.
Jones, D. E., M. R. Ackermann, U. Wille, C. A. Hunter, P. Scott. 2002. Early enhanced Th1 response after Leishmania amazonensis infection of C57BL/6 interleukin-10-deficient mice does not lead to resolution of infection. Infect. Immun. 70:2151.(Charles F. Anderson, Susa)
Experimental Leishmania major infection in mice has been of immense interest because it was among the first models to demonstrate the importance of the Th1/Th2 balance to infection outcome in vivo. However, the Th2 polarization that promotes the development of nonhealing cutaneous lesions in BALB/c mice has failed to adequately explain the mechanisms underlying nonhealing forms of leishmaniasis in humans. We have studied a L. major strain from a patient with nonhealing lesions that also produces nonhealing lesions with ulcerations and high parasite burden in conventionally resistant C57BL/6 mice. Surprisingly, these mice develop a strong, polarized, and sustained Th1 response, as evidenced by high levels of IFN- produced by Leishmania-specific cells in the draining lymph node and in the ear lesion, and an absence of IL-4 or IL-13. The parasites fail to be effectively cleared despite high level induction of inducible NO synthase in the lesion, and despite their sensitivity to killing by IFN--activated macrophages in vitro. Infection of IL-10–/– mice, blockade of the IL-10R, or depletion of CD25+ cells during the chronic phase promotes parasite killing, indicating that IL-10 and regulatory T cells play a role in rendering the Th1 responses ineffective at controlling infection in the skin. Mice with nonhealing primary lesions are nonetheless resistant to reinfection in the other ear. We suggest that nonhealing infections in animal models that are explained not by aberrant Th2 development, but by overactivation of homeostatic pathways designed to control inflammation, provide better models to understand nonhealing or reactivation forms of leishmaniasis in humans.
Introduction
Acquired resistance to the intracellular protozoan Leishmania major is dependent on the development of a Th1-type immune response, marked by the induction of CD4+ and CD8+ T cells mediating IFN--dependent macrophage microbicidal activity. In the murine model for cutaneous leishmaniasis, the mechanisms of acquired resistance to L. major have been well documented using genetically resistant C57BL/6 mice, which reproduce the self-cure disease outcome typically seen in humans (1, 2). Intradermal inoculation of these mice with a biologically appropriate low dose of infectious stage parasites produces an initial prepatent phase of intracellular growth, absent of visible lesions or pathology, followed by lesion development and the accumulation of IFN--producing CD4+ and CD8+ T cells in the lesion (3, 4, 5). Following resolution of the lesion, a low number of parasites persists indefinitely in the site, and the host is protected against a rechallenge infection at a distal site (3, 5).
In contrast to the self-limiting infections with L. major observed in C57BL/6 mice, BALB/c mice develop progressive, nonhealing lesions that are associated with an early, sustained, and dominant Th2 response, especially IL-4 (1, 2). Furthermore, ablation of the cells or cytokines involved in normal Th1 development in resistant mice results in Th2 deviation and progressive infection (6, 7, 8, 9). The Th1/Th2 balance that controls immunity to L. major in these murine models has fostered the main conceptual framework for understanding healing and nonhealing forms of clinical disease. However, it has not been possible to clearly associate a Th2 polarity with nonhealing, systemic, or reactivation forms of leishmaniasis in humans. IFN--producing cells or mRNA remain readily detectable in patients with kala-azar, postkala-azar dermal leishmaniasis, or chronic cutaneous leishmaniasis, and the opposing cytokine most commonly found in these clinical settings is not IL-4, but IL-10 (10, 11, 12, 13, 14, 15, 16, 17, 18, 19, 20, 21). Interestingly, IL-10 has also been found to contribute to BALB/c susceptibility to L. major (22), although its influence is generally obscured by the Th2 bias that is intrinsic to this mouse strain (23). In contrast, a role for IL-10 in regulating immunity in resistant mice has been conclusively shown, in this case mediating not susceptibility per se, but the persistence of low numbers of parasites in the skin following clinical cure (4). Thus, IL-10–/– mice achieve sterile immunity, as do healed C57BL/6 mice treated during the chronic phase with anti-IL-10 receptor Abs. More recently, IL-10-producing naturally occurring CD4+CD25+ regulatory T cells have been shown to home to L. major lesions, and to be necessary for parasite persistence in healed mice (24). These cells may fulfill a primarily homeostatic function, preventing excessive pathology mediated by Leishmania-specific Th1 cells in the inflammatory site. Although regulatory and effector T cell subsets appear to function in equilibrium to maintain latency in healed mice, these findings raise the possibility that an imbalance in regulatory cells or cytokines might control nonhealing or systemic disease outcomes.
In the present study, we have analyzed the role of these opposing host factors in the immune response of C57BL/6 mice to a strain of L. major that was isolated from a patient with nonhealing lesions (25). The lesions persisted for months following multiple courses of treatment, but the patient was found to have a positive skin test and proliferative responses to Leishmania Ags. In the current study, despite the induction of a strong and polarized Th1 response, the parasites are not effectively controlled, and the dermal lesions fail to resolve. We examine the degree of suppression mediated by IL-10 and by CD4+CD25+ regulatory T cells and argue that these are major contributing factors to the evolution of these nonhealing infections.
Materials and Methods
Mice
C57BL/6 mice were purchased from the Division of Cancer Treatment, National Cancer Institute (Frederick, MD). C57BL/10SgSnAi, C57BL/10SgSnAi-[KO]IL-10, and C57BL/6SgSnAi-[KO]IL-4 mice were purchased from Taconic Farms. All mice were maintained in the National Institute of Allergy and Infectious Diseases animal care facility under specific pathogen-free conditions.
Antibodies
For in vivo Ab treatments, mice were given 1.0 mg i.p. injections at the indicated times with anti-CD25 (PC61), anti-IL-10R (1B1.3a), or an isotype control (GL113). CD25 depletion was determined by staining cells with Ab clone 7D4, which showed depletion to be >80% compared with control-treated mice. The following fluorochrome-conjugated Abs for cell surface and intracytoplasmic staining were purchased from BD Pharmingen: PE-anti-CD25 (PC61), FITC-anti-CD25 (clone 7D4), FITC-anti-TCR- (H57-597), PE-Cy5 anti-CD4 (L3T4), PE-Cy5 anti-CD8 (53–6.7), and PE anti-IFN- (XMG1.2). The isotype controls used (all from BD Pharmingen) were rat IgG2b (A95-1), rat IgG2a (R35-95), rat IgG1 (R3-34), and hamster IgG group 2 (Ha4/8). For ELISA measurements of cytokines, the following Ab pairs were purchased from eBioscience. IL-4: 11B11, BVD6-24G2; IL-10: JES5-16E3, JES5-2A5; IFN-: XMG1.2, R4-6A2.
Parasite preparation and intradermal inoculation
L. major clones V1 (MHOM/IL/80/Friedlin) and Sd (MHOM/SN/74/SD) were cultured in medium 199 with 20% heat-inactivated FCS (Gemini Bio-Products), 100 U/ml penicillin, 100 μg/ml streptomycin, 2 mM L-glutamine, 40 mM HEPES, 0.1 mM adenine (in 50 mM HEPES), 5 mg/ml hemin (in 50% triethanolamine), and 1 mg/ml 6-biotin (M199/S). The Sd strain was isolated from a patient who was admitted to the National Institutes of Health for treatment of lesions that persisted following multiple courses of therapy (25). Infective-stage metacyclic promastigotes of L. major were isolated from stationary cultures (4–5 days old) by density gradient centrifugation as described previously (26). Metacyclic promastigotes (1000) were inoculated into the ear dermis using a 30-gauge needle in a volume of 5 μl. The development of the lesion was monitored by measuring the diameter of the ear lesion with a direct-reading vernier caliper (Thomas Scientific).
Processing of ear tissue and estimation of parasite load
Parasite titrations were performed as previously described (3). The two sheets of infected ears were separated, deposited in DMEM containing 100 U/ml penicillin, 100 μg/ml streptomycin, and 0.2 mg/ml Liberase CI purified enzyme blend (Roche Diagnostic Systems), and incubated for 2 h at 37°C. Sheets were then placed in a grinder and processed in a Medimachine tissue homogenizer (BD Biosciences) for 4 min. Tissue homogenates were then filtered through a 70-μm cell strainer (Falcon Products), and aliquots of these were serially diluted in 96-well flat-bottom microtiter plates containing biphasic medium prepared using 50 μl of NNN medium containing 30% of defibrinated rabbit blood overlaid with 50 μl of M199/S. The number of viable parasites in each sample was determined from the highest dilution at which promastigotes could be grown out after 7 days of incubation at 25°C.
Analysis of intradermal lymphocytes, APC preparation, and cytokine analysis
To characterize leukocytes in the inoculation sites, the ears were collected, and the ventral and dorsal dermal sheets were prepared as described above. For cell surface phenotype, cells were fixed in PBS/2% paraformaldehyde for 15 min at 25°C, washed, and resuspended in staining buffer (PBS with 0.1% BSA and 0.01% NaN3) for phenotypic analysis by flow cytometry. For in vitro restimulation, unfractionated lymph node (LN)3 cells or dermal cells were incubated with uninfected or amastigote-infected bone marrow-derived dendritic cells (BMDCs) as a source of APCs. BMDCs were generated from bone marrow in the presence of GM-CSF as previously described (27). LN cells (3 x 106) or dermal cells pooled from six ears were resuspended in RPMI 1640 containing 10% FCS, 10 mM HEPES, 100 U/ml penicillin, and 100 μg/ml streptomycin, and 1 ml of the cell suspension was incubated with uninfected or amastigote-infected BMDCs at a ratio of 5:1 in 24-well plates at 37°C in 5% CO2 for 18 h, with brefeldin A added during the last 5 h. The cells were then fixed and analyzed for surface markers and intracellular IFN-. In some experiments, unfractionated LN or dermal cells were incubated for 48 h with BMDCs or amastigote-infected BMDCs, and IFN-, IL-10, and IL-4 from culture supernatants were quantitated by ELISA (eBioscience) according to the manufacturer’s protocol. The limits of detection for the cytokines were 4 pg/ml for IL-4, 15 pg/ml for IL-10, and 15 pg/ml for IFN-.
Immunolabeling and flow cytometry
Before staining, LN or dermal cells were incubated with an anti-FcIIIR/IIR (clone 2.4G2; BD Pharmingen) mAb in PBS containing 0.1% BSA and 0.01% NaN3. The staining of surface markers and intracellular cytokines was performed sequentially. The cells were stained first for their surface markers (TCR chain receptor, CD8, or CD4), followed by a permeabilization step and staining with anti-IFN-. The lymphocytes were identified by characteristic size (forward light scatter) and granularity (side light scatter), in combination with anti-TCR- chain and anti-CD4 or anti-CD8, or anti-CD25 for surface staining. For each sample, 150,000 events were acquired for analysis. The data were collected and analyzed using CellQuest Pro software and a FACSCalibur flow cytometer (BD Biosciences).
Real-time PCR
For analysis of gene expression, ears were removed and immediately placed in RNAlater stabilization buffer (Qiagen) to prevent degradation. Tissue was then disrupted by placing the ears in mortars containing liquid nitrogen and grinding with a pestle. The disrupted tissue was then homogenized using Qiashredder homogenizers and processed for RNA isolation using the RNeasy minikit (Qiagen), following the manufacturer’s protocol, with all samples being treated with a DNase step to remove genomic DNA. Reverse transcription was performed on 500 ng of total RNA from each ear sample using Superscript III First-Strand synthesis system for RT-PCR (Invitrogen Life Technologies). Real-time PCR was performed on an ABI Prism 7900HT sequence detection system (Applied Biosystems) using 10% of the cDNA reaction in a total volume of 20 μl of PCR mixture. FAM-MGB-labeled primer/probe sets for IFN-, IL-10, IL-13, inducible NO synthase-2 (iNOS-2), and Foxp3 were designed from Applied Biosystems, and VIC-MGB-labeled 18 S rRNA was used as the endogenous control. The relative quantitation of products was determined by the comparative threshold cycle method using the equation 2–cT to determine the fold increase in product. Each gene of interest was normalized to the 18 S rRNA endogenous control, and the fold change in expression was displayed as relative to naive controls. Four ears were used for each time point, and each sample was run in triplicate during the PCR.
In vitro killing assay
Peritoneal macrophages from C57BL/6 mice were plated in eight-chamber Lab-Tek Permanox tissue culture slides (Miles Laboratories) at 5 x 104 cells/well in a volume of 0.4 ml of RPMI 1640 complete. V1 or Sd tissue-derived amastigotes were added to the wells at a ratio of 5:1, and infection was allowed to establish for 72 h at 35°C. At this time, 0.1 μg/ml LPS and titrating amounts of recombinant mouse IFN- were added. After an additional 72 h, the slides were fixed for 1 min in anhydrous methanol and stained with Diff-Quick solutions. The numbers of infected and non-infected macrophages and the number of amastigotes were counted. The percentage of killing relative to untreated macrophages was calculated on the basis of the comparison of total amastigotes per 100 macrophages.
Statistical analysis
To determine whether differences were statistically significant, Student’s t test was performed, using a two-tailed distribution with unpaired samples.
Results
L. major Sd produces a nonhealing phenotype in C57BL/6 mice
To characterize the host response to the L. major Sd strain that produced nonhealing lesions in a patient, C57BL/6 mice were inoculated with 1000 metacyclic promastigotes, and monitored for lesion development in reference to the L. major V1 strain. As previously described (3), mice infected intradermally with V1 parasites produced lesions that healed spontaneously in 8–12 wk following inoculation (Fig. 1). In contrast, when mice were infected with the L. major Sd strain, an initial prepatent phase was followed by the development of lesions that failed to heal. Lesions grew slowly, but progressively larger in diameter, resulting in ulcerations and eventual loss of dermal tissue (Fig. 1). The lesions did not disseminate to other dermal sites nor was there evidence of visceralization beyond the local draining LN (DLN) (data not shown). Lesions harboring the V1 strain showed a peak in parasite number at 6 wk, which by 10 wk had been 90% cleared from the site, and by 22 wk fewer than 100 parasites remained in the healed lesion. In contrast, lesions harboring the Sd strain had a slow progressive increase in the number of parasites throughout the 22 wk that lesions were monitored.
FIGURE 1. Sd strain of L. major produces a nonhealing phenotype in C57BL/6 mice. A total of 103 V1 or Sd metacyclic promastigotes were injected in the ear dermis of mice. A, Development of ear lesions over the course of 22 wk. The mean and SD of six ears per group are shown. B, Parasite loads in ear lesions, showing the mean and SD from six ears per group. C, Photographs of the ears were taken 22 wk postinfection The results are representative of at least three independent experiments.
Priming and recruitment of Th1 cells is not deficient in L. major Sd-infected mice
Because the L. major Sd strain produced a nonhealing phenotype, we investigated the possibility that this L. major substrain induces an aberrant Th2 response in this normally resistant host. As early as 3 wk postinfection, Ag-specific IFN- production was detectable in high levels in the DLN of the Sd-infected mice, at a time when Th1 priming was barely in evidence in the mice infected with V1 (Table I). IFN- production in the Sd-infected mice was maintained at high levels at 6 and 11 wk, whereas IFN- production peaked at around 6 wk in the V1-infected mice and declined significantly by 11 wk, at a time when the majority of parasites had been eliminated. Importantly, at no time point did the DLN cells from the Sd or V1 mice produce significant amounts of IL-4. In contrast, IL-10 was detected by 3 wk in the Sd mice, with high levels being produced by 6 wk postinfection. Apart from the early time point, IL-10 was secreted in even higher levels by the DLN cells from the V1-infected mice.
Table I. ELISA cytokine measurements of draining lymph node cellsa
The number of CD4+ and CD8+ T cells in the ear dermis was analyzed to determine whether the Sd infection was inhibiting the recruitment of effector cells to the site of infection. Fig. 2 shows the frequency of TCR-+CD4+ and TCR-+CD8+ cells present in the lesions at 3, 6, and 10 wk postinfection, along with the total numbers of these subsets. The recruitment of CD4+ T cells was apparent in both groups by 3 wk, though it was slightly higher in the Sd lesions at 3 and 6 wk, and 2-fold higher at 10 wk, when the V1 dermal lesions were beginning to resolve. Recruitment of CD8+ T cells was generally comparable in both groups at the earlier time points, with a slight decrease of CD8+ cells seen in the Sd infections compared with V1 at 10 wk.
FIGURE 2. Sd infection is not defective in the recruitment of lymphocytes to the lesion. At various times postinfection, dermal cells were immunolabeled for TCR- and CD4 or CD8 and analyzed by flow cytometry. Lymphocytes were gated according to forward scatter and side scatter properties. The numbers shown in the upper right quadrant are the absolute numbers of cells per ear (x104) and are the results of four to six ears pooled per time point. The numbers represent the mean and SD of three independent experiments.
The ability of the lesional CD4+ T cells to secrete cytokines in response to infected DC was measured by intracellular staining and by ELISA. Ag-specific IFN--producing cells were detected in both groups as early as 3 wk, increasing in absolute numbers at wk 6, and maintained at high frequency at 10 wk. Strikingly, the Sd infection contained more than twice the number of Th1 effectors at 10 wk, suggesting the higher parasite load at this time was driving the continued expansion and recruitment of effector cells (Fig. 3). Cytokines were also measured by ELISA (Table II). An earlier, more elevated and sustained Ag-specific IFN- response by Sd lesional cells was also noted by ELISA. IL-4-producing cells were not detectable in either group, either by intracellular staining (data not shown), or by ELISA, except for the slightly elevated levels produced by the V1 lesional cells at wk 6 (Table II). In contrast, substantial amounts of IL-10 were produced by ear lesional cells from both groups by 3 wk, and high leve
ls continued to be produced by the V1 and Sd dermal cells throughout the healing and chronic phase, respectively. Interestingly, the majority of the IL-10 produced by the dermal cells did not require Ag re-stimulation for release. Taken together, these results indicate that there is not a defect in Th1 priming, nor a defect in recruitment of Th1 cells to the site of the infection. The Sd parasites persist and produce nonhealing lesions despite the development of a polarized Th1 response.
FIGURE 3. Sd-infected ears contain high numbers of IFN- secreting effector cells. Dermal cells from six V1- or Sd-infected ears were pooled and restimulated for 18 h in the presence of BMDCs or amastigote-infected BMDCs (BMDCs/Lm). Brefeldin A was added during the last 5 h, and cells were then fixed and permeabilized for intracellular cytokine staining. The numbers in the upper right quadrant are the percentages of CD4+ cells secreting IFN-. The data are representative of at least three independent experiments.
Table II. ELISA cytokine measurements of cells from ear lesionsa
To determine whether the Sd strain might persist in the lesion due to an intrinsic resistance to NO-dependent killing by activated macrophages, we performed an in vitro killing assay, in which infected macrophages were activated with LPS and titrated amounts of rIFN-, providing both the priming and triggering stimuli needed to generate optimal levels of NO (Fig. 4). Both strains were efficiently killed when the infected macrophages were activated using 10 ng/ml rIFN-, and although a broad dose range defined the LD50 for each of the strains, the Sd strain did not display a greater resistance to killing at any of the concentrations of rIFN- used. Thus the resistance of the Sd strain to killing in vivo is apt to reflect differences in the activation state of the infected cells in the inflammatory site.
FIGURE 4. V1 and Sd strains are equally susceptible to killing by activated macrophages in vitro. Peritoneal macrophages were plated in 8-well chamber slides and infected with V1 or Sd amastigotes. Macrophages were activated by the addition of LPS and titrated amounts of rIFN-, 72 h after infection. After an additional 72 h, macrophages were fixed and stained for parasite quantitation. The percentage of killing was determined by the number of amastigotes per 100 macrophages and was based on the comparison with unactivated macrophages.
The role of IL-10 and CD4+CD25+ Treg cells in the development of nonhealing lesions due to L. major Sd
Because the presence of IL-10 has been shown to permit the persistence of L. major in the face of a robust Th1 response in healed mice (4), we considered the possibility that the nonhealing phenotype might be due to a modest shift in the balance between IFN- and IL-10 cytokines, not revealed in the ELISA assays of Ag re-stimulated dermal cells. As IL-10 has been shown to be produced by keratinocytes in dermal lesions (13), and by Leishmania-infected macrophages in vitro (22, 28), real-time RT-PCR on RNA extracted directly from the ears was used to better quantitate total IL-10 produced by the different possible sources of IL-10 in the inflammatory site. Compared with V1, the Sd strain activated elevated levels of IFN- mRNA, in agreement with the flow cytometry and ELISA data, but also contained significantly increased message for IL-10 at 11 and 14 wk postinfection (Fig. 5). The difference seen at these time points, especially at wk 14, is apt to be secondary to the inflammatory conditions associated with active vs healed lesions. No difference was observed at the 6-wk time point, around the time when the clinical outcomes were beginning to diverge. Additionally, IL-13 was measured by real-time PCR to exclude the possible role of this Th2-associated cytokine. At both 6 and 12 wk postinfection, neither group expressed significant IL-13 messages above that detectable in naive ears. The data reinforce the absence of a Th2 immune deviation in the L. major Sd-infected ears.
FIGURE 5. Sd lesions contain elevated levels of mRNA for IFN-, IL-10, and Foxp3, and low amounts of IL-13. Total RNA was isolated directly from the ears at the indicated times during the infection, reverse transcribed, and analyzed by real-time PCR. The target genes were normalized to the endogenous control, and the values shown are the fold increase relative to expression in naive ears. Each bar is the geometric mean and SD of four individual ears. (*, p < 0.05; **, p < 0.01).
A potential source of IL-10 in the nonhealing lesions is CD4+CD25+ regulatory T cells. The presence of these cells has been shown to control the persistence of L. major in healed mice (24), and adoptive transfer of these cells, or their Ag-induced expansion, has been recently shown to reactivate parasite growth and dermal pathology in healed lesions (29). Thus it was important to determine whether there was an increased frequency of CD24+CD25+ Treg in the Sd mice. The forkhead/winged helix transcription factor Foxp3 has been shown to be specifically expressed by CD25+ Treg, as well as by CD25– T cells with regulatory activity (30, 31). Compared with V1, the Sd lesions contained a significantly increased level of Foxp3 mRNA at 11 and 14 wk postinfection and a small, though nonsignificant, increase at 6 wk (Fig. 5). And whereas the number of Foxp3 transcripts remained steady throughout the acute, healing, and latent stages of V1 infection, there was a steady increase during progressive lesion development in the Sd non-cure mice. Flow cytometry analysis to measure the frequency of CD4+CD25+ and CD4+CD25– T cells in the lesions revealed no major differences between the two infections during the first 11 wk (Fig. 6). There was a modest increase in the total number of CD25+ Treg in the Sd lesions at 6 wk, and especially at 11 wk when there was an 3-fold increase compared with V1, consistent with the Foxp3 real-time PCR data.
FIGURE 6. V1- and Sd-infected ears contain high numbers of CD4+CD25+ cells. Dermal cells were removed from ear lesions at the indicated times and stained for CD4 and CD25. The numbers in the upper right quadrant represent the percentage of total CD4+ cells that express CD25 and the absolute numbers of CD4+CD25+ cells (x104) per ear. The data are from one representative experiment of at least three independent experiments.
To determine what role IL-10 may play in prevention of healing, we first took a genetic approach by infecting IL-10–/– mice with either V1 or Sd (Fig. 7). At 10 wk postinfection, there was a 2-log reduction in L. major Sd parasite burden in the IL-10–/– mice compared with the wild-type B/10 mice (Fig. 7B). However, sterile cure of the Sd strain was not achieved as was apparent with V1, consistent with previous observations (4). Despite the reduced parasite load, the Sd lesions in the IL-10–/– mice were larger than in the wild-type mice throughout the course of infection, consistent with a critical role of IL-10 in modulating immune-mediated pathology (Fig. 7A). In contrast to the IL-10–/– mice, the IL-4–/– mice were found to be equally susceptible to L. major Sd as was wild-type B/6 mice, both in terms of the severity of their nonhealing lesions (not shown) and the failure to effectively reduce the parasite burden during the chronic stage (Fig. 7B). To determine the effect of a transient block in IL-10 signaling, mice were treated with anti-IL-10R Ab during the chronic phase of the infection. Mice were sacrificed 1 wk after the course of treatment, and the parasite burden was measured (Fig. 7C). The treatment had a dramatic curative effect in the Sd-infected mice, with 99% of parasites being cleared from the site. The treatment also reduced the parasite burden in the healing V1 lesions, as previously reported. In neither infection did the treatment result in sterile cure, though it is possible that more prolonged treatment, or treatment during more latent stage of infection with V1, might have produced the complete clearance of V1 achieved in the prior studies. The more effective clearance of parasites following anti-IL-10R treatment was associated in both groups with increased IFN- and iNOS expression in the lesion, as measured by real-time PCR. However, it should be noted that the increased iNOS expression in the L. major Sd lesions was slight, and perhaps more importantly, the levels in the untreated Sd-infected mice were already high (Fig. 8). Sd-infected mice were also depleted of CD25+ cells during the chronic stage of the infection by treating with anti-CD25 Ab over a course of 2 wk, and analyzing the parasite load and immune response 1 wk following the last treatment. The anti-CD25-treated mice had a significant, 10-fold reduction in parasite number compared with the isotype controls (Fig. 9), and slight increases in both IFN- and iNOS-2 expression in the lesion, though again the levels of both transcripts were already high in the untreated mice.
FIGURE 7. IL-10 deficiency or anti-CD25 treatment promotes parasite killing in the Sd-infected mice. Parasite loads in ear lesions in C57BL/10 or IL-10–/– mice or C57BL/6 or IL-4–/– mice infected with 103 metacyclics and sacrificed at wk 10 for parasite enumeration (A), C57BL/6 mice treated with anti-IL-10R Ab every 4 days during the last 2 wk of infection and sacrificed at wk 10 for parasite enumeration (B), and C57BL/6 mice treated with anti-CD25 Ab every 4 days during the last 2 wk of infection and sacrificed at wk 12 for parasite enumeration (C). The data are representative of two independent experiments (*, p < 0.05 compared with control mice).
FIGURE 8. Anti-IL-10R- and anti-CD25-treated mice have increased expression of IFN- and iNOS-2 mRNA. At 12 wk postinfection, mice were given two i.p. injections of Ab 3 days apart. Three days after the second treatment, total RNA was isolated from ears, reverse transcribed, and analyzed by real-time PCR. The target genes were normalized to the endogenous control, and the values shown are the fold increase relative to expression in naive mice. Each bar is the geometric mean and SD of four individual ears (*, p < 0.05 compared with control mice).
FIGURE 9. Sd-infected mice develop protective immunity to a rechallenge. Fourteen weeks following primary infection, mice were rechallenged in the opposite ear with 103 metacyclics, along with naive animals as a comparison. A, Ear lesion development in rechallenged and naive mice. The mean and SD of five mice per group are shown. B, Parasite burdens in individual ears in the primary site and secondary rechallenge site 8 wk after rechallenge.
Mice with nonhealing L. major Sd infection develop concomitant immunity to reinfection
Mice that developed nonhealing L. major Sd lesions at 14 wk were rechallenged in the contralateral ear with the Sd strain to determine whether protective immunity against reinfection had been generated during the primary infection. Despite the inability to heal the primary lesion, the rechallenge ears did not develop measurable pathology, and 8 wk after rechallenge they had a 10-fold reduction in parasite burden compared with naive-infected controls (Fig. 9). Coinfection with the Sd and V1 strains using 1000 metacyclics in each ear did not compromise the ability of the mice to control and resolve the V1 lesions, again despite development of nonhealing Sd lesions in the contralateral ear (data not shown).
Discussion
Using low dose intradermal inoculation in conventionally resistant C57BL/6 mice, we have characterized the host immune response to a strain of L. major that produces a nonhealing phenotype. This occurs despite the development of a strong and polarized Th1 response, as measured by IFN- production in the DLN as well as in the site of the infection, and the correspondingly low levels of IL-4 and IL-13 produced in these sites. In addition, the nonhealing phenotype developed despite normal recruitment of CD4+ and CD8+ lymphocytes, and despite high level induction of iNOS in the lesion, conditions that are collectively associated with effective control of L. major infection in the skin (2). The possibility that the Sd strain is intrinsically more resistant to IFN--mediated killing was addressed by determining the concentrations of IFN- required to activate peritoneal macrophages for leishmanicidal capacity in vitro, and no significant difference compared with the V1 strain was observed. The non-cure phenotype seen in this model is clearly divergent from the BALB/c model of susceptibility, in which the infection is marked by a strong Th2 bias. The disease-promoting cytokine that could be identified in this model is not IL-4, but IL-10, as indicated by the enhanced host resistance observed in Sd-infected IL-10–/– B10 mice, or in B6 mice treated with anti-IL-10R Ab. CD4+CD25+ regulatory T cells were also shown to play a role, as the in vivo depletion of CD25+ cells promoted greater clearance of parasites from the skin. It is important to consider that the L. major BALB/c model has in general failed to adequately explain severe forms of clinical disease, which seem not to be associated with a Th1 response defect per se, but with concomitant expression of IL-10 (10, 11, 13, 14, 15, 16, 17, 18, 19, 20, 21).
Although IL-10 and CD4+CD25+ T cells are induced by L. major Sd and are required for the evolution of the nonhealing lesions, these responses also accompany the dermal infections with V1 that produce a healing phenotype in B6 mice. In this setting, IL-10 produced by naturally occurring regulatory T cells has been shown to prevent sterile cure, and the regulatory cells seem to function in equilibrium with Leishmania-specific effectors to establish and maintain latent infection in the skin (24). Might the inability to heal the primary Sd lesion be the result of an imbalance in suppressor cells or cytokines in the inflammatory site? While IL-10 production and both IL-10 and Foxp3 mRNA were maintained at elevated levels in the nonhealing compared with the healing L. major lesions, at the critical earlier stage of infection, around wk 6, when the V1 parasites began to be effectively controlled, the inability of an equally potent Th1 response to clear the Sd parasites was not associated with increased IL-10 production or an increased frequency of CD4+CD25+ T cells in the site. In fact, the most striking difference between the immune response induced by these two strains is the high level IFN- production by cells from the DLN or ear lesions in the Sd-infected mice as early as 3 wk postchallenge, at a time when the Th1 response had yet to develop in the mice infected with V1. How this paradoxical finding relates to the evolution of the non-cure phenotype is not known. It is of course possible that additional suppressor factors other than IL-10 might accompany the early, elevated IFN- response in the Sd mice to shift the balance in favor of parasite survival. One indication that the L. major Sd strain induces factors in addition to IL-10 that promote infection is the observation that a low level infection was maintained in the IL-10–/– mice, whereas complete cure was achieved in the IL-10–/– mice infected with V1, confirming our prior studies (4).
Although IL-10 production may not be a sufficient condition for the evolution the non-cure phenotype, it nonetheless contributes in an essential way to this outcome, as indicated by the more effective clearance of the Sd strain in the IL-10-deficient and anti-IL-10R-treated mice. The source of the IL-10 induced by Sd infection is not clear from these studies. Depletion of the CD25+ cells did not appreciably reduce the amount of IL-10 produced in the site (data not shown). Furthermore, the IL-10R blockade consistently promoted greater parasite clearance than did the CD25+ cell depletion, suggesting alternative sources of IL-10. It has been shown that amastigote-infected macrophages produce IL-10 (28, 32), as do macrophages following FcR ligation (22), which in this context might involve anti-Leishmania Abs and released Ags. Keratinocytes are also known to produce IL-10 (13, 33), and their ability to do so in visceral leishmaniasis patients is associated with reactivation disease (postkala azar dermal leishmaniasis) (13). It is interesting that in contrast to the IFN- response, the IL-10 produced by the lesional cells was consistently at high levels in the absence of Ag restimulation, suggesting that non-Ag-specific sources of IL-10 are present in the inflammatory site. It is possible that differences in the frequency or activation state of these IL-10-producing cells, obscured by the quantitation of total IL-10, represent the critical difference between the V1 and Sd inflammatory sites. Autocrine IL-10 produced by infected macrophages is apt, for example, to be far more effective at deactivating infected cells than IL-10 produced by Treg. And although the quantitation of total iNOS in the lesion indicates a highly activated tissue environment, the deactivated tissue might be confined to the infected macrophages or other parasitized cells.
The nonhealing infections produced by L. major Sd in B6 mice mimics in some respects the pattern of susceptibility displayed by "resistant" mice infected with new world cutaneous Leishmania amazonensis strains and their related subspecies Leishmania mexicana (34, 35). In fact, the L. amazonensis studies in resistant mice (C3H and C57BL/6) provided an early challenge to the role of IL-4 and Th2 polarization as a necessary condition for the evolution of nonhealing disseminating forms of leishmaniasis, as L. amazonensis-infected IL-4 knockout mice and anti-IL-4-treated mice still developed nonhealing lesions (35). In contrast, results from two studies involving IL-10 knockout mice indicated a role for IL-10 in promoting parasite growth, since 1–2 log reductions were observed in the deficient mice (36, 37). A critical difference between these non-cure phenotypes and the non-cure Sd infections reported here is that whereas the new world strains induced only low level IFN- responses throughout infection, Sd elicits a strong sustained Th1 response.
We observed in our studies of L. major Sd infection in B6 mice that despite their inability to heal their primary lesion, when the animals were rechallenged in the contralateral ear, they were substantially protected. Thus the IL-10, Treg, and possibly other suppressive mechanisms that function within the primary inflammatory site to prevent effective clearance by Th1 cells do not inhibit the expression of Th1 effector activity in a naive re-challenge site. We have argued that the activation of Treg during infection provides a benefit to the host by not only controlling the severity of inflammation, but by promoting parasite persistence the Treg will indirectly maintain a memory pool necessary for resistance to reinfection (24). Since Treg may be preferentially recruited to or expanded by signals that accumulate within inflamed tissue, they may not be able to home to naive rechallenge sites in sufficient time or numbers to suppress the expression of concomitant immunity.
The availability of a non-cure phenotype of L. major infection associated not with aberrant Th2 development but with ongoing Th1 responses rendered ineffective by an imbalance in homeostatic suppressive cells or cytokines may better reflect the mechanisms controlling non-curing forms of cutaneous visceral leishmaniasis in humans.
Disclosures
The authors have no financial conflict of interest.
Footnotes
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
1 Current address: Department of Microbiology and Tropical Medicine, George Washington University, Washington, DC 20037.
2 Address correspondence and reprint requests to Dr. David L. Sacks, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Laboratory of Parasitic Diseases, Building 4, Room 126, 4 Center Drive MSC 0425, Bethesda, MD 20892-0425. E-mail address: dsacks{at}nih.gov
3 Abbreviations used in this paper: LN, lymph node; BMDC, bone marrow-derived dendritic cell; iNOS, inducible NO synthase; DLN, draining LN.
Received for publication September 8, 2004. Accepted for publication December 16, 2004.
References
Reiner, S. L., R. M. Locksley. 1995. The regulation of immunity to Leishmania major. Annu. Rev. Immunol. 13:151.
Sacks, D., N. Noben-Trauth. 2002. The immunology of susceptibility and resistance to Leishmania major in mice. Nat. Rev. Immunol. 2:845.
Belkaid, Y., S. Mendez, R. Lira, N. Kadambi, G. Milon, D. Sacks. 2000. A natural model of Leishmania major infection reveals a prolonged "silent" phase of parasite amplification in the skin before the onset of lesion formation and immunity. J. Immunol. 165:969.
Belkaid, Y., K. F. Hoffmann, S. Mendez, S. Kamhawi, M. C. Udey, T. A. Wynn, D. L. Sacks. 2001. The role of interleukin (IL)-10 in the persistence of Leishmania major in the skin after healing and the therapeutic potential of anti-IL-10 receptor antibody for sterile cure. J. Exp. Med. 194:1497.
Belkaid, Y., E. Von Stebut, S. Mendez, R. Lira, E. Caler, S. Bertholet, M. C. Udey, D. Sacks. 2002. CD8+ T cells are required for primary immunity in C57BL/6 mice following low-dose, intradermal challenge with Leishmania major. J. Immunol. 168:3992.
Belosevic, M., D. S. Finbloom, P. H. Van Der Meide, M. V. Slayter, C. A. Nacy. 1989. Administration of monoclonal anti-IFN- antibodies in vivo abrogates natural resistance of C3H/HeN mice to infection with Leishmania major. J. Immunol. 143:266.
Ghalib, H. W., M. R. Piuvezam, Y. A. Skeiky, M. Siddig, F. A. Hashim, A. M. el-Hassan, D. M. Russo, S. G. Reed. 1993. Interleukin 10 production correlates with pathology in human Leishmania donovani infections. J. Clin. Invest. 92:324.
Karp, C. L., S. H. el-Safi, T. A. Wynn, M. M. Satti, A. M. Kordofani, F. A. Hashim, M. Hag-Ali, F. A. Neva, T. B. Nutman, D. L. Sacks. 1993. In vivo cytokine profiles in patients with kala-azar: marked elevation of both interleukin-10 and interferon-. J. Clin. Invest. 91:1644.
Kenney, R. T., D. L. Sacks, A. A. Gam, H. W. Murray, S. Sundar. 1998. Splenic cytokine responses in Indian kala-azar before and after treatment. J. Infect. Dis. 177:815.
Gasim, S., A. M. Elhassan, E. A. Khalil, A. Ismail, A. M. Kadaru, A. Kharazmi, T. G. Theander. 1998. High levels of plasma IL-10 and expression of IL-10 by keratinocytes during visceral leishmaniasis predict subsequent development of post-kala-azar dermal leishmaniasis. [Published erratum appears in 1998 Clin. Exp. Immunol. 112:547.
Ribeiro-de-Jesus, A., R. P. Almeida, H. Lessa, O. Bacellar, E. M. Carvalho. 1998. Cytokine profile and pathology in human leishmaniasis. Braz. J. Med. Biol. Res. 31:143.
Holaday, B. J., M. M. Pompeu, S. Jeronimo, M. J. Texeira, A. Sousa Ade, A. W. Vasconcelos, R. D. Pearson, J. S. Abrams, R. M. Locksley. 1993. Potential role for interleukin-10 in the immunosuppression associated with kala azar. J. Clin. Invest. 92:2626
Ismail, A., A. M. El Hassan, K. Kemp, S. Gasim, A. E. Kadaru, T. Moller, A. Kharazmi, T. G. Theander. 1999. Immunopathology of post kala-azar dermal leishmaniasis (PKDL): T-cell phenotypes and cytokine profile. J. Pathol. 189:615.
Gasim, S., A. M. Elhassan, A. Kharazmi, E. A. Khalil, A. Ismail, T. G. Theander. 2000. The development of post-kala-azar dermal leishmaniasis (PKDL) is associated with acquisition of Leishmania reactivity by peripheral blood mononuclear cells (PBMC). Clin. Exp. Immunol. 119:523.
Ajdary, S., M. H. Alimohammadian, M. B. Eslami, K. Kemp, A. Kharazmi. 2000. Comparison of the immune profile of nonhealing cutaneous Leishmaniasis patients with those with active lesions and those who have recovered from infection. Infect. Immun. 68:1760.
Louzir, H., P. C. Melby, A. Ben Salah, H. Marrakchi, K. Aoun, R. Ben Ismail, K. Dellagi. 1998. Immunologic determinants of disease evolution in localized cutaneous leishmaniasis due to Leishmania major. J. Infect. Dis. 177:1687.
Akuffo, H., K. Maasho, M. Blostedt, B. Hojeberg, S. Britton, M. Bakhiet. 1997. Leishmania aethiopica derived from diffuse leishmaniasis patients preferentially induce mRNA for interleukin-10 while those from localized leishmaniasis patients induce interferon-. J. Infect. Dis. 175:737.
Melby, P. C., F. J. Andrade-Narvaez, B. J. Darnell, G. Valencia-Pacheco, V. V. Tryon, A. Palomo-Cetina. 1994. Increased expression of proinflammatory cytokines in chronic lesions of human cutaneous leishmaniasis. Infect. Immun. 62:837.
Kane, M. M., D. M. Mosser. 2001. The role of IL-10 in promoting disease progression in leishmaniasis. J. Immunol. 166:1141.
Noben-Trauth, N., R. Lira, H. Nagase, W. E. Paul, D. L. Sacks. 2003. The relative contribution of IL-4 receptor signaling and IL-10 to susceptibility to Leishmania major. J. Immunol. 170:5152.
Lutz, M. B., N. Kukutsch, A. L. Ogilvie, S. Rossner, F. Koch, N. Romani, G. Schuler. 1999. An advanced culture method for generating large quantities of highly pure dendritic cells from mouse bone marrow. J. Immunol. Methods 223:77.
Carrera, L., R. T. Gazzinelli, R. Badolato, S. Hieny, W. Muller, R. Kuhn, D. L. Sacks. 1996. Leishmania promastigotes selectively inhibit interleukin 12 induction in bone marrow-derived macrophages from susceptible and resistant mice. J. Exp. Med. 183:515.
Mendez, S., S. K. Reckling, C. A. Piccirillo, D. Sacks, Y. Belkaid. 2004. Role for CD4+CD25+ regulatory T cells in reactivation of persistent leishmaniasis and control of concomitant immunity. J. Exp. Med. 200:201.
Fontenot, J. D., M. A. Gavin, A. Y. Rudensky. 2003. Foxp3 programs the development and function of CD4+CD25+ regulatory T cells. Nat. Immunol. 4:330.
Hori, S., T. Nomura, S. Sakaguchi. 2003. Control of regulatory T cell development by the transcription factor Foxp3. Science 299:1057.
Guizani-Tabbane, L., K. Ben-Aissa, M. Belghith, A. Sassi, K. Dellagi. 2004. Leishmania major amastigotes induce p50/c-Rel NF-B transcription factor in human macrophages: involvement in cytokine synthesis. Infect. Immun. 72:2582.
Moore, K. W., R. de Waal Malefyt, R. L. Coffman, A. O’Garra. 2001. Interleukin-10 and the interleukin-10 receptor. Annu. Rev. Immunol. 19:683.
Roberts, M., J. Alexander, J. Blackwell. 1990. Genetic analysis of Leishmania mexicana infection in mice single gene Scl-2 controlled predisposition to cutaneous lesion development. J. Immunogenet. 17:89
Afonso, L., P. Scott. 1993. Immune responses associated with susceptibility of C57BL/10 mice to Leishmania amazonensis. Infect. Immun. 61:2952.
Padigel, U. M., J. Alexander, J. P. Farrell. 2003. The role of interleukin-10 in susceptibility of BALB/c mice to infection with Leishmania mexicana and Leishmania amazonensis. J. Immunol. 171:3705.
Jones, D. E., M. R. Ackermann, U. Wille, C. A. Hunter, P. Scott. 2002. Early enhanced Th1 response after Leishmania amazonensis infection of C57BL/6 interleukin-10-deficient mice does not lead to resolution of infection. Infect. Immun. 70:2151.(Charles F. Anderson, Susa)