Cutaneous leishmaniasis causes a spectrum of diseases from self-healing to severe nonhealing lesions. Defining the factors contributing to lesion resolution may help in developing new therapies for those patients with chronic disease. We found that infection with Leishmania major increases the expression of vascular endothelial growth factor-A and vascular endothelial growth factor receptor (VEGFR)-2 and is associated with significant changes in the blood and lymphatic vasculature at the site of infection. Ab blockade of VEGFR-2 during infection led to a reduction in lymphatic endothelial cell proliferation and simultaneously increased lesion size without altering the parasite burden. These data show that L. major infection initiates enhanced vascular endothelial growth factor-A/VEGFR-2 signaling and suggest that VEGFR-2-dependent lymphangiogenesis is a mechanism that restricts tissue inflammation in leishmaniasis.

Cutaneous leishmaniasis has a wide spectrum of clinical manifestations, ranging from self-healing to chronic debilitating disease. There is no vaccine for human leishmaniasis, the drugs against the parasite are extremely toxic, and, furthermore, patients are often refractory to treatment (13). Leishmania parasites are killed by macrophages in response to IFN-γ produced by CD4+ Th1 cells; thus, severe disease occurs in the absence of a strong Th1 response (4). However, even when an appropriate adaptive immune response develops and parasites are controlled, cutaneous lesions often persist, suggesting that the inflammatory response can drive pathology (58). Thus, defining the factors that control lesion development and resolution is important in developing novel therapies to address the disease in patients.

Although many factors were shown to contribute to the development and severity of leishmanial lesions (5, 79), the role of changes in the vasculature in cutaneous leishmaniasis has not been explored. Vascular remodeling is a hallmark of inflammation and leads to morphological and functional changes in the vascular network that can influence the recruitment of cells, as well as their exit from the tissue (1013). The formation of new blood vessels during inflammation and the increased vascular permeability enhance cell recruitment but, concomitantly, may also promote pathology (1014). Moreover, the expansion of the lymphatic vasculature, which supports the initial immune response, also provides a route for drainage of fluid and inflammatory cells out of the tissue (10, 15, 16).

Members of the vascular endothelial growth factor (VEGF) family, including VEGF-A, VEGF-C, and VEGF-D, are produced by a variety of cell types and induce changes in the vascular network during cancer, wound healing, and inflammation. VEGF-A binds VEGF receptor (VEGFR)-2, which is primarily expressed on blood endothelial cells (BECs), and promotes angiogenesis and vascular permeability (17). Alternatively, VEGF-C and VEGF-D bind to VEGFR-3 on lymphatic endothelial cells (LECs), promoting lymphangiogenesis (18). VEGF family members are elevated in inflammatory settings, and neutralization of VEGFR-2 signaling reduces inflammation, whereas VEGFR-3 blockade increases inflammation in the skin (15, 19). Although VEGF-A binding VEGFR-2 on BECs primarily leads to angiogenesis, VEGF-A can also mediate inflammation-induced lymphangiogenesis, and VEGF-A produced in the skin can have profound effects at other sites, such as the draining lymph node (dLN) (2024). Thus, VEGF signaling might be important in the development and resolution of leishamanial lesions, and the role of this pathway has not been defined during infection.

To address this question, we examined the role of VEGF signaling in cutaneous leishmaniasis. In this article, we show that Leishmania major infection induces vascular remodeling in the skin and that the expression of VEGF-A and its receptor VEGFR-2 are elevated. The kinetics of the expression of these mediators mirror the presence of leishmanial lesions and coincide with increased BEC and LEC proliferation at the infection site. Moreover, inhibition of VEGF-A/VEGFR-2 signaling during L. major infection specifically affected LECs and led to increased disease pathology. Taken together, these data suggest that L. major infection activates the VEGF-A/VEGFR-2 signaling pathway, leading to vascular remodeling, and suggest that VEGFR-2–mediated lymphangiogenesis is a mechanism that limits inflammation and promotes lesion resolution.

Female C57BL/6 mice were purchased from the National Cancer Institute, and Tie2-GFP transgenic mice, in which GFP is predominantly expressed by endothelial cells (ECs), were purchased from The Jackson Laboratory and bred at the University of Pennsylvania. Mice were housed in the School of Veterinary Medicine at the University of Pennsylvania under pathogen-free conditions and used for experiments between 6 and 8 wk of age. All procedures were performed in accordance with the guidelines of the University of Pennsylvania Institutional Animal Care and Use Committee.

The L. major (WHO/MHOM/IL/80/Friedlin) strain was used for experiments. Parasites were grown in vitro in Schneider’s Drosophila medium (Life Technologies) supplemented with 20% heat-inactivated FBS (Invitrogen), 2 mM l-glutamine and 2 mM l-glutamine (Sigma), 100 U/ml penicillin, and 100 μg/ml streptomycin (Sigma). Metacyclic stationary-phase promastigotes were isolated from 4–5-d cultures by Ficoll density gradient separation (Sigma) (25). For dermal ear infections, 2 × 106 parasites in 10 μl PBS (Lonza) were injected intradermally into the ear. Lesion development was monitored weekly by measuring ear thickness and lesion area with a caliper, and the lesion volume was calculated. Pathology was also scored using a previously published scoring system: no lesion (0), swelling/redness (1), deformation of the ear pinna (2), ulceration (3), partial tissue loss (4), or total tissue loss (5) (26). To determine parasite loads and cellular content, ears were enzymatically digested using 0.25 mg/ml Liberase (Roche) and 10 μg/ml DNase I (Sigma) in incomplete RPMI 1640 (Life Technologies) for 90 min at 37°C. To determine parasite burdens in the tissue, limiting dilution assays were performed (27).

For histology, L. major–infected and naive ears were fixed in 10% buffered formalin and embedded in paraffin. Longitudinal 6-μm sections were subjected to Masson’s trichrome stain. For immunofluorescence microscopy, ears and dLNs were frozen in Tissue-Tek OCT (Sakura). Longitudinal 6-μm sections were fixed with 3% formaldehyde, blocked with 10% goat serum (Sigma) for 1 h, and stained with rat anti-CD31 (BD Pharmingen) and rabbit anti–LYVE-1 (AngioBio) overnight, followed by secondary Abs goat anti-rabbit Alexa Fluor 488 (Life Technologies) or goat anti-rat DyLight 549 (Jackson ImmunoResearch) for 1 h. Sections were stained with DAPI (Molecular Probes) and mounted with Prolong Gold (Life Technologies). Images were captured using a Nikon Digital Sight DS-Fi1 color camera on a Nikon E600 microscope.

Mice were anesthetized and maintained at a core temperature of 37°C. Ears were attached to an imaging platform using tissue glue. The mice were injected retro-orbitally with 50 μl of 1% Evans blue (Alfa Aesar) and imaged immediately. Imaging was performed with a Leica SP5 two photon microscope system (Leica Microsystems) equipped with a picosecond or femtosecond laser (Coherent). The standard wavelength used for the two-photon imaging was 900 nm, which allowed optimal excitation of the used fluorophores. Images were obtained using a 20× water-dipping lens. The total volume of the fluorescence of Evans blue was calculated using Volocity (Perkin Elmer) at time point zero (∼2 min after injection), and the calculated fluorescence at time point zero was subtracted from subsequent time points.

Near infrared (NIR) nanopolymersomes were formed by dissolving an amphiphilic diblock copolymer composed of polyethylene glycol and polybutadiene in chloroform with the lipophilic NIR fluorophore DiR (Thermo Fisher Scientific). This solution was evaporated to form a thin polymer film, and polymersomes were formed by film rehydration at 60°C overnight. Nanopolymersomes were synthesized by sonication and extrusion to an average diameter of 200 nm, as previously described (28). Mice were injected i.v. with NIR nanopolymersomes, and the accumulation of NIR fluorescence was imaged from whole-mount tissues of infected and contralateral ears after 3 h using an Odyssey infrared imaging system (LI-COR Biosciences). For the Miles assay, 100 μl of 4% Evans blue was injected i.v., and ears were weighed and imaged after 2 h. The dye was extracted from the tissue by placing ears in formamide for 48 h at 60°C. The samples were centrifuged, and the OD at 650 nm was recorded. The amount of Evans blue/ear was calculated using a standard curve and the weight of the tissue.

Cells were incubated with fixable Aqua dye (Invitrogen) to assess viability. Fc receptors were blocked with CD16/32 (eBioscience, San Diego, CA), and cells were surface stained using anti-CD45, anti-Ly6C, and anti-VEGFR-2 (from eBioscience), as well as anti-CD64, anti-CD11b, anti-CD31, and anti-podoplanin–biotin (from BioLegend), followed by streptavidin-PECy7 (eBioscience). For intracellular cytokine staining, dLN cells were stimulated with Brefeldin A (3 μg/ml; eBioscience) with PMA (100 ng/ml) and ionomycin (1 μg/ml; both from Sigma) for 4 h before surface staining for anti-CD4. dLN cells were fixed with 2% paraformaldehyde (Electron Microscopy Sciences), permeabilized with 0.2% saponin buffer, and stained with anti–IFN-γ (BioLegend). Cell events were acquired on an LSR II Fortessa flow cytometer (BD Biosciences, San Jose, CA) and analyzed using FlowJo software (TreeStar).

BrdU (BD Pharmingen) was placed in the drinking water (0.8 mg/ml) for 3 d, and mice were injected i.p. with 1 mg of BrdU 24 and 1 h prior to euthanasia. After tissue processing and cell surface staining, cells were fixed and permeabilized with a BrdU kit (BD Pharmingen) and intracellularly stained with anti-BrdU–allophycocyanin after DNase I treatment, according to the manufacturer’s instructions.

mRNA was extracted by the RNeasy Plus Mini kit (QIAGEN). RNA was reverse transcribed with a High-Capacity cDNA Reverse Transcription Kit (Applied Biosystems). Quantitative real-time PCR was performed using SYBR Green PCR Master Mix on a ViiA 7 Real-Time PCR system (Life Technologies). Primer sequences are listed in Supplemental Table I. The results were normalized to the housekeeping gene ribosomal protein S14 gene (RPSII) using the comparative threshold cycle method (2−ΔΔCT) for relative quantification (8, 29).

Mice were given anti–VEGFR-2 DC101 Ab (0.8 mg/mouse i.p. every 3 d; kindly provided by Bronislaw Pytowski, Eli Lilly and Company, New York, NY) or rat IgG control (clone 2A3; Bio X Cell) starting 1 d prior to infection and continuing for the first 4 wk of infection.

Statistical significance was determined using a two-tailed Student unpaired t test with a p value <0.05 in GraphPad Prism 5.

Given that human leishmanial lesions exhibit VEGF-A and VEGFR-2 expression, and VEGF-A/VEGFR-2 signaling can contribute to the inflammatory response through remodeling of the vasculature, the role of vascular remodeling was evaluated in cutaneous leishmaniasis. Cutaneous leishmaniasis was studied using the well-characterized experimental murine model in which C57BL/6 mice were dermally infected with L. major parasites. In this model, dermal lesions develop slowly, peaking at 4–6 wk postinfection (p.i.), with 106–107 parasites in the lesions. Once parasites are controlled by the immune response, dermal lesions in C57BL/6 mice spontaneously resolve by 8–10 wk p.i. (Fig. 1A). To visualize the ECs and vasculature following infection, Tie2-GFP reporter mice were used. Mice were infected with L. major parasites in the ear, and the vasculature was analyzed using multiphoton intravital microscopy (30). Compared with naive mice, at 5 wk p.i. when lesions are at their peak, the structure and morphology of the vessels in infected skin are highly tortuous and irregularly shaped with sprouts, reminiscent of tumor vessels (Fig. 1B, 1F). Histology of infected ears also revealed an increase in the number of vessels in the skin over time that mirror lesion development (Fig. 1C, 1D). In addition, immunofluorescence microscopy showed an increase in CD31 staining, a marker of ECs, in the ears of infected mice compared with controls (Fig. 1E). Taken together, these data show that L. major infection induces vascular remodeling in the skin, and increases in CD31+ cells suggest an expansion of the EC network.

FIGURE 1.

Dermal vascular remodeling occurs during L. major infection. (A) C57BL/6 mice were infected with 2 × 106L. major metacyclic parasites in the ear dermis, and lesion progression was monitored over time. (B) Multiphoton microscopy of the dermis of Tie2-GFP–expressing mice infected for 5 wk compared with uninfected controls. (C) Masson’s trichrome stained infected and contralateral control ears from C57BL/6 mice showing vessels containing RBCs. (D) Total number of vessels/ear section quantified by microscopy at 5 wk p.i. (n = 3–5 mice per group). Data are mean + SEM. (E) Confocal microscopy of frozen sections stained with Abs against CD31 (red) showing alterations in the EC network with infection compared with naive control C57BL/6 mice. Dashed lines represent the tissue edge. (F) EC sprouting is detected in infected mice (arrowheads) after i.v. Evans blue to visualize the vasculature. Scale bars, control 47 μm (B, left); infection 43 μm (B, right); 100 μm (C and E); and 16 μm (F). *p < 0.05, t test.

FIGURE 1.

Dermal vascular remodeling occurs during L. major infection. (A) C57BL/6 mice were infected with 2 × 106L. major metacyclic parasites in the ear dermis, and lesion progression was monitored over time. (B) Multiphoton microscopy of the dermis of Tie2-GFP–expressing mice infected for 5 wk compared with uninfected controls. (C) Masson’s trichrome stained infected and contralateral control ears from C57BL/6 mice showing vessels containing RBCs. (D) Total number of vessels/ear section quantified by microscopy at 5 wk p.i. (n = 3–5 mice per group). Data are mean + SEM. (E) Confocal microscopy of frozen sections stained with Abs against CD31 (red) showing alterations in the EC network with infection compared with naive control C57BL/6 mice. Dashed lines represent the tissue edge. (F) EC sprouting is detected in infected mice (arrowheads) after i.v. Evans blue to visualize the vasculature. Scale bars, control 47 μm (B, left); infection 43 μm (B, right); 100 μm (C and E); and 16 μm (F). *p < 0.05, t test.

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Vascular remodeling is often associated with the proliferation of ECs. Therefore, the frequency of proliferating ECs during L. major infection was examined by BrdU incorporation in C57BL/6 mice. At 20 d p.i., when lesions are starting to develop in infected mice, flow cytometric analysis of CD45 CD31+ ECs revealed an increase in the percentage and number of BrdU+ proliferating ECs compared with naive control skin (Fig. 2A–C). The percentage of BrdU+ proliferating ECs was elevated as early as 12 d p.i., when lesions are not yet apparent in infected mice, and increased percentages of BrdU+ ECs persisted in infected ears at day 35 p.i., when lesions are at their peak (Fig. 2B). BECs and LECs can be distinguished based on their podoplanin expression: BECs are CD31+ podoplanin, and LECs are CD31+ podoplanin+ (Fig. 2D). When BEC and LEC populations were examined individually, flow cytometric analysis showed increased percentages and numbers of BrdU+ BECs in infected mice compared with naive controls (Fig. 2E–G). Similarly, L. major–infected ears displayed increased percentages and numbers of BrdU+ LECs compared with controls (Fig. 2E, 2H, 2I). Altogether, these results clearly indicate that EC proliferation is ongoing at the site of infection and suggest that L. major infection triggers angiogenesis and lymphangiogenesis in the dermis; this rapid expansion of the blood and lymphatic vasculature likely has a strong effect on vessel permeability within the infected tissue.

FIGURE 2.

Endothelial proliferation is detected at the site of infection during leishmaniasis. (A) Representative flow cytometry plots showing gating of CD31+ CD45 ECs and the percentage of ECs incorporating BrdU from naive control and 20-d infected ears. Cells were gated on total, live, singlets previously. Percentage (B) and number (C) of proliferating BrdU+ ECs in naive ears and at 12, 20, and 35 d p.i. (D) Dermal BECs and LECs were separated by podoplanin expression during FACS analysis. (E) Representative flow plots showing the percentages of BECs and LECs incorporating BrdU from naive and infected ears at 20 d p.i. Frequency (F) and number (G) of BrdU+ BECs from naive and infected ears at 12, 20, and 35 d p.i. Frequency (H) and number (I) of BrdU+ LECs from naive and infected ears at 12, 20, and 35 d p.i. Data are representative of at least one experiment depending on the time point, with four or five mice per group per time point. Data are mean + SEM. *p < 0.05, **p < 0.005, ***p < 0.0005, t test.

FIGURE 2.

Endothelial proliferation is detected at the site of infection during leishmaniasis. (A) Representative flow cytometry plots showing gating of CD31+ CD45 ECs and the percentage of ECs incorporating BrdU from naive control and 20-d infected ears. Cells were gated on total, live, singlets previously. Percentage (B) and number (C) of proliferating BrdU+ ECs in naive ears and at 12, 20, and 35 d p.i. (D) Dermal BECs and LECs were separated by podoplanin expression during FACS analysis. (E) Representative flow plots showing the percentages of BECs and LECs incorporating BrdU from naive and infected ears at 20 d p.i. Frequency (F) and number (G) of BrdU+ BECs from naive and infected ears at 12, 20, and 35 d p.i. Frequency (H) and number (I) of BrdU+ LECs from naive and infected ears at 12, 20, and 35 d p.i. Data are representative of at least one experiment depending on the time point, with four or five mice per group per time point. Data are mean + SEM. *p < 0.05, **p < 0.005, ***p < 0.0005, t test.

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Vascular remodeling often coincides with enhanced vascular permeability, which allows cells and inflammatory mediators to migrate into inflamed tissues. To determine whether L. major infection leads to increased vascular permeability in the skin, C57BL/6 mice were infected in the ear; at 5 wk p.i., they were injected i.v. with nanopolymersomes containing an NIR fluorophore 3 h prior to ex vivo imaging of whole-mount ear tissue. The size of the nanopolymersomes (average diameter of 200 nm) prevents these particles from escaping through normal vasculature, but they were shown to accumulate at sites of enhanced permeability (31). Imaging showed infected ears had an increase in fluorescence compared with controls, suggesting that more polymersomes accumulated in the infected ear compared with the contralateral ear of the same mouse (Fig. 3A, 3B). Moreover, the polymersomes specifically collected at lesions within the infected ears, suggesting increased vascular leakage directly at the infection site. In complementary experiments, C57BL/6 mice were subjected to a Miles assay at 5 wk p.i. in which Evans blue was injected i.v. (32). Infected ears had a greater amount of Evans blue collect in the tissue compared with contralateral ears (Fig. 3C, 3D). To examine the extent and proximity of vascular leakage with parasitized cells, intravital multiphoton imaging was used to visualize cutaneous vascular leakage in real time p.i. with DsRed-labeled parasites. For these experiments, naive and infected Tie2-GFP mice were given Evans blue i.v. and imaged immediately. Images of the ear were acquired every 30 s for 30 min. Time-lapse images revealed that the Evans blue stayed confined within the vasculature in naive control ears (Fig. 3E–G, Supplemental Videos 1, 3). In contrast, the Evans blue escaped out of the vessels and into the extravascular space in infected ears (Fig. 3E–G, Supplemental Videos 2, 4). Collectively, these data demonstrate that L. major infection induces dramatic changes in the cutaneous vasculature, including an increase in vascular permeability.

FIGURE 3.

Increased vascular permeability in the dermis is associated with L. major infection. C57BL/6 mice were infected with L. major parasites in the ear dermis and analyzed 5–6 wk p.i. (A) Mice were injected i.v. with polymersomes containing NIR fluorophores, and the accumulation of fluorescence was imaged from whole-mount tissue sections of infected and control contralateral ears after 3 h. (B) The total infrared fluorescence of each ear was quantified. Data are representative of one experiment with four mice per group. (C) Images of infected ears and naive controls 2 h after i.v. Evans blue, with corresponding quantification from one representative experiment of two with three to five mice per group (D). (EG) Tie2-GFP–expressing mice were infected with DsRed-labeled L. major parasites intradermally, and the dermis of infected and naive mice was analyzed by multiphoton microscopy for 30 min. (E) Time-lapse two-dimensional images were taken every 10 min starting immediately after i.v. Evans blue. Scale bars, 45 μm. (F) Three-dimensional images were acquired immediately after i.v. Evans blue and 30 min later (Supplemental Videos 1–4). Parasites are red, Tie2 is green, and Evans blue is white. Scale bars, 60 μm. (G) Quantification of the volume of fluorescence intensity of Evans blue in the extravascular space was calculated every 30 s from the multiphoton imaging. Representative data from one infected mouse and one control mouse is shown; imaging data were collected from at least five mice per group. Data are mean + SEM. ***p < 0.0005, t test.

FIGURE 3.

Increased vascular permeability in the dermis is associated with L. major infection. C57BL/6 mice were infected with L. major parasites in the ear dermis and analyzed 5–6 wk p.i. (A) Mice were injected i.v. with polymersomes containing NIR fluorophores, and the accumulation of fluorescence was imaged from whole-mount tissue sections of infected and control contralateral ears after 3 h. (B) The total infrared fluorescence of each ear was quantified. Data are representative of one experiment with four mice per group. (C) Images of infected ears and naive controls 2 h after i.v. Evans blue, with corresponding quantification from one representative experiment of two with three to five mice per group (D). (EG) Tie2-GFP–expressing mice were infected with DsRed-labeled L. major parasites intradermally, and the dermis of infected and naive mice was analyzed by multiphoton microscopy for 30 min. (E) Time-lapse two-dimensional images were taken every 10 min starting immediately after i.v. Evans blue. Scale bars, 45 μm. (F) Three-dimensional images were acquired immediately after i.v. Evans blue and 30 min later (Supplemental Videos 1–4). Parasites are red, Tie2 is green, and Evans blue is white. Scale bars, 60 μm. (G) Quantification of the volume of fluorescence intensity of Evans blue in the extravascular space was calculated every 30 s from the multiphoton imaging. Representative data from one infected mouse and one control mouse is shown; imaging data were collected from at least five mice per group. Data are mean + SEM. ***p < 0.0005, t test.

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Given that VEGFs are the primary vascular mediators responsible for EC proliferation and increased vascular permeability, the kinetics of the expression of multiple VEGF family members during L. major infection in vivo were examined. At 5 wk p.i., the expression of VEGF-A (Fig. 4A) and its receptor, VEGFR-2 (Fig. 4B), were elevated at the site of infection in C57BL/6 mice compared with naive skin. VEGF-C and VEGF-D, as well as their receptor, VEGFR-3, were not significantly elevated at the time points analyzed (Fig. 4C–E). The kinetics of VEGF-A expression mirror the development and resolution of leishmanial lesions and coincide with increased EC proliferation. Because VEGF-A transcript levels were elevated in the skin at the site of infection, the expression of VEGF-A was also examined in the dLN. Similar to the site of infection, the expression of VEGF-A was significantly increased in the lymph nodes draining infected ears compared with naive controls (Fig. 4F). Transcript levels of HIF1α, which drives VEGF-A expression, were increased in the ear and dLN at 5 wk p.i., compared with naive controls (Fig. 4G, 4H). Taken together, these data show that the angiogenic HIF1α/VEGF-A/VEGFR-2 pathway is elevated during leishmaniasis and may play a role in vascular remodeling at the site of infection.

FIGURE 4.

VEGF-A and the corresponding receptor, VEGFR-2, are expressed at increased levels at the site of L. major infection. C57BL/6 mice were infected with L. major parasites in the ear dermis, and ears were analyzed by quantitative real-time PCR at 5, 9, and 15 wk p.i. for the expression of VEGF-A (A), VEGFR-2 (B), VEGF-C (C), VEGF-D (D), and VEGFR-3 (E). (F) Quantitative real-time PCR for VEGF-A expression in lymph nodes draining infected and naive control ears at 5 wk p.i. Quantitative real-time PCR for HIF1α expression at 5 wk p.i. in naive and infected ears (G) and dLNs (H). Relative mRNA expression normalized to the housekeeping gene RPS11 is presented as the mean + SEM with five mice per group. Data are representative of at least two independent experiments. *p < 0.05, **p < 0.005, ***p < 0.0005, t test.

FIGURE 4.

VEGF-A and the corresponding receptor, VEGFR-2, are expressed at increased levels at the site of L. major infection. C57BL/6 mice were infected with L. major parasites in the ear dermis, and ears were analyzed by quantitative real-time PCR at 5, 9, and 15 wk p.i. for the expression of VEGF-A (A), VEGFR-2 (B), VEGF-C (C), VEGF-D (D), and VEGFR-3 (E). (F) Quantitative real-time PCR for VEGF-A expression in lymph nodes draining infected and naive control ears at 5 wk p.i. Quantitative real-time PCR for HIF1α expression at 5 wk p.i. in naive and infected ears (G) and dLNs (H). Relative mRNA expression normalized to the housekeeping gene RPS11 is presented as the mean + SEM with five mice per group. Data are representative of at least two independent experiments. *p < 0.05, **p < 0.005, ***p < 0.0005, t test.

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Because vascular remodeling, including increased EC proliferation and vascular leakiness, was associated with elevated VEGF-A and VEGFR-2 levels during L. major infection, the role of the VEGF-A/VEGFR-2 pathway was analyzed in vivo using an Ab-neutralization strategy. Inhibition of VEGFR-2 signaling from the onset of infection reduced CD31+ EC proliferation, as assessed by BrdU incorporation at 2 wk p.i. (Fig. 5A, 5B). Additionally, BECs and LECs were analyzed individually based on their podoplanin expression; VEGFR-2 blockade led to a decrease in the percentage of BrdU+ LECs during infection (Fig. 5A, 5D). However, surprisingly, the frequency of BrdU+ BECs was not altered with VEGFR-2 blockade, suggesting that VEGFR-2 inhibition specifically limited the proliferation of LECs without affecting BEC proliferation at the site of infection (Fig. 5A, 5C, 5D). Given that inhibition of VEGFR-2 signaling targeted LECs and not BECs, we next compared the expression of VEGFR-2 on dermal BECs and LECs. Flow cytometric results showed that LECs expressed higher levels of VEGFR-2 than BECs in the skin of naive and infected mice, supporting the preferential effects seen in LECs with Ab blockade (Fig. 5E). To determine whether VEGFR-2 blockade altered parasite control or disease pathology, C57BL/6 mice were infected and given anti–VEGFR-2 Ab for the first 4 wk of infection. Importantly, mice receiving anti–VEGFR-2 Ab exhibited increased lesion volumes compared with isotype control–treated mice (Fig. 5F). Additionally, inhibition of VEGFR-2 signaling exacerbated pathology, and some mice developed ulcerations, which was rarely seen in the control group (Fig. 5G, 5H). These increased lesion sizes were not associated with impaired parasite control because parasite burdens in mice treated with anti–VEGFR-2 Ab were not higher at 4 wk p.i. compared with controls (Fig. 5I). Additionally, there were no significant differences in the percentages of immune cell populations, including macrophages, Ly6C+ inflammatory monocytes, or CD4+ T cells, in the dermis of mice treated with anti–VEGFR-2 Ab compared with controls at 4 wk p.i. (Fig. 5J). Furthermore, the frequency of CD4+ T cells producing IFN-γ in the dLN was similar between groups (Fig. 5J). Similarly, the transcript levels of IFN-γ, IL-4, or IL-10 at the site of infection did not differ between mice treated with anti–VEGFR-2 Ab and controls (data not shown). Given that regulatory T cells and plasmacytoid dendritic cells are known to express VEGFR-2, the frequency of these immune cell populations was also examined following infection; however, the percentage of regulatory T cells and plasmacytoid dendritic cells was similar with and without VEGFR-2 blockade following infection (data not shown). Histologically, ears from mice given anti–VEGFR-2 Ab showed a cellular infiltrate that exceeded that in control animals (Fig. 5K). Additionally, ears were stained with LYVE-1 to examine the lymphatic vasculature by immunofluorescence. VEGFR-2 blockade led to a decrease in lymphatic vessel diameter compared with isotype-treated controls but did not alter the density of lymphatic vessels at the site of infection (Fig. 5L–N). Taken together, these data suggest that VEGF-A/VEGFR-2 signaling mediates the proliferation of LECs and contributes to lymphangiogenesis during infection and that inhibition of this pathway exacerbates pathology, demonstrating a protective role for the VEGF-A/VEGFR-2 signaling pathway during infection-induced inflammation.

FIGURE 5.

VEGFR-2 blockade targets LECs and increases pathology during L. major infection. Mice were treated with anti–VEGFR-2 neutralization Ab or rat isotype control for the first 4 wk of infection. Naive mice are shown as controls. (A) Representative flow cytometry plots showing the percentage of BrdU+ cells after gating on total, live, singlet, CD31+ CD45 ECs or on BECs (CD31+ podoplanin) or LECs (CD31+ podoplanin+) separately. Quantification of the percentage of proliferating BrdU+ cells of total ECs (B), BECs (C), and LECs (D) from the skin of naive controls or infected mice at 2 wk p.i. (E) Representative flow cytometry graphs of BECs and LECs showing VEGFR-2 expression. (F) Lesion volume was monitored over time. (G) Lesion pathology was scored over time. (H) Representative images of anti–VEGFR-2–treated and control ears at 4 wk p.i. The dashed circles show the lesion area. (I) After 4 wk of anti–VEGFR-2 Ab treatment, the numbers of parasites in the ear were quantified by LDA. (J) Cells were previously gated on total, live, singlets from anti–VEGFR-2–treated or control animals at 4 wk p.i. Quantification of the percentage of CD64+ macrophages, Ly6C+ inflammatory monocytes of CD11b+ cells, and CD4+ T cells of CD45+ cells in the skin was assessed by flow cytometry. Cells were previously gated on total, live, singlets from anti–VEGFR-2–treated or control animals at 4 wk p.i. The quantification of the percentage of CD4+ T cells from the dLN producing IFN-γ was assessed by flow cytometry after ex vivo treatment with PMA and ionomycin in the presence of Brefeldin A for 4 h. (K) Representative H&E-stained ear sections of anti–VEGFR-2–treated and control ears at 4 wk p.i. Scale bars, 100 μm. (L) Confocal microscopy of frozen sections from anti–VEGFR-2–treated and control animals stained for LYVE-1 (green) at 4 wk p.i. Scale bars, 100 μm. Quantification of the percent area of LYVE-1 staining (M) and luminal area of LYVE-1+ vessels (N). Data are representative of at least two independent experiments, with four or five mice per group. Data are mean ± SEM. *p < 0.05, **p < 0.005, t test.

FIGURE 5.

VEGFR-2 blockade targets LECs and increases pathology during L. major infection. Mice were treated with anti–VEGFR-2 neutralization Ab or rat isotype control for the first 4 wk of infection. Naive mice are shown as controls. (A) Representative flow cytometry plots showing the percentage of BrdU+ cells after gating on total, live, singlet, CD31+ CD45 ECs or on BECs (CD31+ podoplanin) or LECs (CD31+ podoplanin+) separately. Quantification of the percentage of proliferating BrdU+ cells of total ECs (B), BECs (C), and LECs (D) from the skin of naive controls or infected mice at 2 wk p.i. (E) Representative flow cytometry graphs of BECs and LECs showing VEGFR-2 expression. (F) Lesion volume was monitored over time. (G) Lesion pathology was scored over time. (H) Representative images of anti–VEGFR-2–treated and control ears at 4 wk p.i. The dashed circles show the lesion area. (I) After 4 wk of anti–VEGFR-2 Ab treatment, the numbers of parasites in the ear were quantified by LDA. (J) Cells were previously gated on total, live, singlets from anti–VEGFR-2–treated or control animals at 4 wk p.i. Quantification of the percentage of CD64+ macrophages, Ly6C+ inflammatory monocytes of CD11b+ cells, and CD4+ T cells of CD45+ cells in the skin was assessed by flow cytometry. Cells were previously gated on total, live, singlets from anti–VEGFR-2–treated or control animals at 4 wk p.i. The quantification of the percentage of CD4+ T cells from the dLN producing IFN-γ was assessed by flow cytometry after ex vivo treatment with PMA and ionomycin in the presence of Brefeldin A for 4 h. (K) Representative H&E-stained ear sections of anti–VEGFR-2–treated and control ears at 4 wk p.i. Scale bars, 100 μm. (L) Confocal microscopy of frozen sections from anti–VEGFR-2–treated and control animals stained for LYVE-1 (green) at 4 wk p.i. Scale bars, 100 μm. Quantification of the percent area of LYVE-1 staining (M) and luminal area of LYVE-1+ vessels (N). Data are representative of at least two independent experiments, with four or five mice per group. Data are mean ± SEM. *p < 0.05, **p < 0.005, t test.

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Although the immune response to L. major has been well characterized, changes in the vasculature that might influence disease progression have not been investigated. Thus, many studies have focused on the frequency and activation status of immune cells at the site of infection, but little work has gone into understanding how the vasculature contributes to the immune response during infection or whether manipulation of the pathways involved in vascular remodeling alters pathogen control, pathology, and/or lesion resolution. In this article, we showed that L. major infection leads to dramatic morphological changes in the dermal vascular network. The vessels from infected mice were found to be highly tortuous and dilated, reminiscent of the vasculature associated with tumors, and they exhibited increased permeability, confirming recent findings in this model (33). Infection also increased EC proliferation and the levels of VEGF-A and VEGFR-2. These findings are relevant to human leishmaniasis, because VEGF-A and VEGFR-2 are expressed in leishmanial lesions from humans along with dilated vessels, corroborating our findings in the mouse model (34). To determine whether VEGF-A/VEGFR-2 signaling contributes to pathogen control or the severity of disease, infected mice were given VEGFR-2–blocking Abs, which specifically decreased the percentage of proliferating LECs and exacerbated pathology. These data suggest that VEGF-A/VEGFR-2 signaling mediates vascular remodeling and contributes to infection-induced lymphangiogenesis that serves as a protective mechanism to restrict the inflammatory response.

During L. major infection, we found elevated levels of HIF1α, which is the major transcription factor that responds to oxygen deprivation and cellular stress. HIF activation can occur in response to hypoxia, cytokines, TLR ligation, and reactive oxygen species (ROS), all of which are present during L. major infection. Thus, we hypothesize that HIF1α activation is driving VEGF-A expression and production during infection. Parasites may directly activate HIF1α themselves, or the hypoxic environment rich in proinflammatory cytokines, such as TNF-α and IL-1, may also contribute to HIF activation and, thus, VEGF-A production (3537). Furthermore, Leishmania spp. can activate TLRs, so VEGF-A production may result from HIF1α activation in response to TLR ligation (29, 3840). Additionally, Leishmania spp. can induce ROS from monocytes and macrophages, so ROS may activate HIF1α and, thus, VEGF-A production during infection (41). As a result, ongoing work in the laboratory is focused on dissecting out the individual roles of host and parasite factors that may act alone or in a combinatorial fashion to enhance HIF activation and, thus, VEGF-A expression and production following L. major infection.

The cellular source of VEGF-A has not been identified, and it remains to be determined whether VEGF-A expression is mediated directly by HIF1α during leishmaniasis. Given their role in other inflammatory settings and cancer, we hypothesize that myeloid cells, which are present in high numbers in leishmanial lesions, are responding to factors in the tissue driving HIF1α activation and, thus, VEGF-A expression and production (20, 22, 42, 43). Therefore, we speculate that myeloid cells are the major producers of VEGF-A in vivo following L. major infection. However, other cells, such as keratinocytes, dendritic cells, and B cells, can contribute to VEGF-A production in wound healing and inflammation and may play a role in this setting (21, 44). Therefore, future studies aim to identify the cellular source of VEGF-A and to determine whether VEGF-A production is mediated by HIF1α activation following L. major infection. During infection, we hypothesize that myeloid cells located at the lesion site produce VEGF-A that predominantly targets LECs in the skin to initiate lymphangiogenesis. We speculate that ECs are the primary targets of VEGF-A, given their high expression levels of VEGFR-2; however, other cells types, such as macrophages, also express VEGFR-2, suggesting that VEGF-A may also be acting on hematopoietic cells during infection. As a result, future studies will characterize the effects of VEGF-A on immune cell populations during L. major infection.

During L. major infection, VEGFR-2 signaling limits pathology by increasing lymphatic vessel number and function. Surprisingly, we found that VEGFR-2 signaling induces LEC proliferation and lymphangiogenesis following L. major infection. This result was unexpected, given that VEGFR-2 signaling is typically thought to mediate angiogenesis and promote dermal pathology (19). Although less appreciated, VEGF-A can drive lymphangiogenesis, but lymphangiogenesis is primarily mediated by VEGF-C/VEGF-D/VEGFR-3 signaling (19, 22, 24, 4547). In addition to decreased lymphangiogenesis, lymphatic vessel diameter was decreased with VEGFR-2 blockade, similar to the results obtained using skin UV-B irradiation (48). These findings suggest that collapsed lymphatic vessels may be functionally impaired in their ability to transport fluid and cells away from the site of inflammation. Although VEGFR-2 blockade resulted in larger leishmanial lesions, these mice controlled parasites similar to controls, which is consistent with a lack of differences in the immune response between groups (data not shown). These data suggest that the pre-existing lymphatic vasculature is sufficient to drive the adaptive immune response, but VEGFR-2–mediated lymphangiogenesis is required to drain fluid and newly infiltrated cells away from the site of infection. Moreover, lymphangiogenesis is also required for normal wound repair, and the increased pathology during L. major infection may reflect a defect in lesion resolution (42, 46, 49). Altogether, our studies show that L. major infection induces VEGF-A/VEGFR-2 signaling that promotes lymphangiogenesis and protects against exacerbating pathology.

Our data suggest that VEGF-A/VEGFR-2 signaling is not essential for angiogenesis during L. major infection, so other factors, such as FGF-2, PDGF, and angiopoietins, may contribute to angiogenesis. Importantly, these pathways can induce angiogenesis in the absence of VEGFR-2 signaling (5056). Therefore, we hypothesize that other angiogenic mediators, such as FGF-2, which is elevated during infection, may drive the expansion of the blood vasculature during leishmaniasis (57). Alternatively, these angiogenic factors may serve a redundant or compensatory role only upon the neutralization of VEGFR-2.

In this study, L. major infection led to elevated levels of VEGF-A and increased EC proliferation in the dermis and the dLN. However, VEGFR-2 blockade led to a reduction in EC proliferation in the ear, whereas no differences were detected in the dLNs. Consistent with this observation, the size and cellularity of the dLNs were similar between mice treated with anti–VEGFR-2 Ab and controls. In the dLN, the VEGFR-2 and VEGFR-3 pathways may compensate for one another for EC proliferation. Alternatively, other signaling pathways, such as lymphotoxin β (LTβ)/LTβR or LIGHT/LTβR, which were shown to be important in dLN hypertrophy and L. major infection, may also contribute to EC proliferation (5861). Furthermore, the lack of inhibition of EC proliferation in the total dLN may not reflect differences in distinct regions of the dLN (62). In addition, we found higher expression levels of VEGFR-2 in the dermis compared with the dLN, so blockade strategies may preferentially affect cells in the skin. As a result, studies are ongoing in the laboratory to identify the pathways responsible for angiogenesis and lymphangiogenesis in the dLN during infection.

In summary, we showed that L. major infection activates VEGF-A/VEGFR-2 signaling that promotes vascular remodeling and drives the expansion of the lymphatic network, providing a conduit for fluid and cells to migrate to the dLN. Upon blockade of VEGFR-2–mediated lymphangiogenesis, we speculate that fluid, debris, and cells are sequestered at the site of infection, resulting in exacerbated pathology. Because the lesions of many patients with leishmaniasis exhibit an immense inflammatory infiltrate, despite low numbers of parasites, stimulation of lymphangiogenesis may provide a novel target to alleviate the inflammation. Thus, understanding the role of the vasculature during infection and inflammation has important implications for the development of novel strategies targeting the vasculature to limit the pathology associated with parasites, as well as other infectious and inflammatory diseases.

We thank Drs. Gudrun Debes, Sydney Evans, and Celeste Simon for discussions and Ba Nguyen for technical assistance (University of Pennsylvania). We acknowledge the Abramson Cancer Center Histology Core and the Flow Cytometry Core Facility in the School of Medicine, as well as the PennVet Imaging Core Facility and, specifically, Dr. Gordon Ruthel (University of Pennsylvania).

This work was supported by National Institutes of Health Grants R01 AI 106842 (to P.S.) and R01 AI 41158 (to C.A.H.). Imaging experiments were carried out on instrumentation supported by National Institutes of Health Grant S10RR027128, the School of Veterinary Medicine, the University of Pennsylvania, and the Commonwealth of Pennsylvania. T.W. was supported by National Institutes of Health Postdoctoral Research Grant F32 AI 114080.

The online version of this article contains supplemental material.

Abbreviations used in this article:

     
  • BEC

    blood endothelial cell

  •  
  • dLN

    draining lymph node

  •  
  • EC

    endothelial cell

  •  
  • LEC

    lymphatic endothelial cell

  •  
  • LTβ

    lymphotoxin β

  •  
  • NIR

    near infrared

  •  
  • p.i.

    postinfection

  •  
  • ROS

    reactive oxygen species

  •  
  • VEGF

    vascular endothelial growth factor

  •  
  • VEGFR

    VEGF receptor.

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The authors have no financial conflicts of interest.