In this work, parasites of the species L. amazonensis (MHOM/BR/75/JOSEFA) were used. The promastigotes were maintained in 25cm² culture flasks with M199 medium (pH 7.2), supplemented with 10% fetal bovine serum, 5µg/mL bovine hemin, 50U/mL penicillin, and 50µg/mL streptomycin.

Mice from BALB/c lineage, females aged 8–12 weeks, were used. The mice came from the Biotério Central de Camundongos of the Centro de Ciências da Saúde (CCS) from the Universidade Federal do Rio de Janeiro, mice were maintained under the Protocol of the Ethics Committee on the Use of Experimental Animals No. 133/23 of the CCS.

The parasites were cultivated until the beginning of the stationary phase. The culture containing the promastigote forms were collected, washed with phosphate buffer saline (PBS); and centrifuged for 10 minutes at 4°C at 1000 x G force. After centrifugation, the supernatant was discarded, and the cells were washed with PBS. The procedure was repeated twice. Then the cells were counted and adjusted to a concentration of 2x10⁸ parasites/mL. The animals were sedated with ketamine and xylazine and infected with 2x10⁶ parasites using Hamilton^® syringes coupled with 34G needles. Different infection methods were implemented: intradermal in the ear performed at an angle of 20°; Cutaneous Nose, with the inoculum carried out at the apex of the mouse’s nose; Septum, with the inoculum occurring in the mouse septum and “Nasobasal”, with the inoculum being carried out at an angle of 60° towards the lower “floor” part of the nose. The lesion was monitored with photos. Weekly measurements of the mice’s ears were taken using Mitutoyo™ thickness gauges.

Animals were sedated and placed in the LabPET8 Flex Triumph Gamma Medica™ tomography system at CENABIO, UIPA. The equipment settings included 60kV, 480µA, 1024 projections, 8-minute acquisition, and 30x magnification with a 39.46mm field of view, focusing on the snout and mucous membrane. After acquisition, data was reconstructed and analyzed using 3D Slicer™ software, with the Scissors tool applied to generate the snout area.

Tissues from animal snouts were fixed in 4% formaldehyde in PBS for 15 days. Decalcification was performed for 28 days using 10% EDTA. After fixation and decalcification, tissues were dehydrated through a series of ethanol and xylene solutions, then embedded in paraffin. Sections (5µm thick) were cut, rehydrated, and stained with Hematoxylin & Eosin (H&E). Tissue processing was done by the Histotechnology Platform at IOC - FIOCRUZ. Histology was performed with n = 5 per group.

At the end of each experiment, parasite load was assessed using the limiting dilution method (LDA). Animals were euthanized with an overdose of Ketamine and Xylazine, and tissues (snout, nose tip, premaxilla, ear, cervical lymph node, ear lymph node, NALT, and spleen) were collected, weighed, and macerated in 1mL of M199. Serial 1:4 dilutions were made, and the results were plotted. Three experiments (n=3-5) were conducted, with two examining the entire snout and one focusing on segmented parts, 12 weeks post-infection.

Flow cytometric analysis of cells from cervical and ear lymph nodes of infected or naïve animals was performed. Tissues were macerated, and cells counted with 0.1% trypan blue dye. 1x10⁶ cells were plated in a 96-well plate, centrifuged, and stimulated with PMA (20ng/mL), ionomycin (1µg/mL), and monensin (2.5µg/mL) in Complete RPMI for 4 hours at 37°C. Cells were blocked with anti-FcR, surface stained, fixed using the eBioscience™ FoxP3 kit, and intracellularly labeled. The samples were analyzed on a BD LSR Fortessa X-20 cytometer. Supplementary Table S1 denotes the cytometer configuration. Supplementary Table S2 shows the antibodies (BioLegend™) used and their concentrations. Supplementary Figure S1 denotes the performed gating strategy.

Results are represented by Standard Error of the Mean (SEM). Statistical significance was performed by unpaired two tailed Student t test with 95% confidence interval (p<0,05) for the respective data: Lesion profile by tomography, parasite load quantification and cytometry. While One Way ANOVA with 95% interval of confidence (p<0,05) was performed using Tukey post-test, for the ELISA OD Sum data. Statistically significant differences were defined as * for p<0,05; ** for p<0,005; *** for p<0,0005.

To assess Leishmania infection in the nasal mucosa, 2x10⁶ promastigotes of L. amazonensis were directly inoculated into different nasal sites of mice. After 6 weeks, the Nose Cutaneous (NC) and Nasobasal (NB) groups developed visible lesions with edema and redness ( Figure 1A ). These lesions progressed and became necrotic by 12 weeks in some cases. The muzzle volume was significantly larger in the NC and NB groups (387mm³ and 616mm³) compared to the Sham group ( Figure 1B ), resembling the progressive ear infection, which reached 1.8mm in thickness ( Figures 1C, D ). In contrast, the Septum group showed no visible lesions throughout the study ( Figure 1A ). These results indicate that nasal inoculation can cause tissue damage, depending on the infection site.

Parasite load in infected mice ranged from 10⁶ to 10⁸ parasites per gram of tissue in the Nose Cutaneous (NC) and Nasobasal (NB) groups, while the Septum (ST) group showed only 10⁴ parasites, with Only 20% of animals testing positive ( Figure 2A ). Ear dermis infections had 10⁶ parasites per gram ( Figure 2B ). Parasites were mostly concentrated in the edema sites, such as the nose tip and pre-maxillary regions ( Figures 2C, D ). Some animals in the NB group had around 10⁵ parasites in NALT ( Figure 2E ). Only one animal in the NC group showed spleen infection ( Figure 2F ). The cervical lymph nodes of NC and NB mice had 10⁶ parasites ( Figure 2G ), while the ear lymph nodes had 10⁵ parasites ( Figure 2H ). The Septum group showed parasitic load in the cervical lymph node but not the nasal cavity ( Figure 2G ). These results indicate that mucocutaneous L. amazonensis infection leads to a higher parasite content in nasal mucosa infection compared to ear dermis, with the cervical lymph node being the primary draining node.

MicroCT analysis was performed to assess nasal cavity impairment, focusing on the NC and NB groups, as the ST group showed no visible lesions or significant parasite load. NC-infected mice typically had lesions at the nose apex, while NB-infected mice had lesions affecting the mucosa and premaxilla, extending to the upper lips ( Figures 3A, B ). No nasal cavity enlargement, perforations, or deformation were observed compared to controls ( Supplementary Figure S2 ), but tissue swelling was evident in both NC and NB groups ( Figures 3A-D ). Axial sections showed noticeable edema on the outer nasal cavity ( Figures 3C, D ). These results indicate that lesion profiles depend on the inoculation site, mainly affecting the nasal tissues initial portions.

Histological analysis of the nasal cavity revealed significant inflammation and parasite presence. In the NC group, edema with a neutrophilic infiltrate and vacuolated macrophages containing amastigotes was observed ( Figures 3E , Supplementary Figure S3 ), along with respiratory epithelium loss ( Supplementary Figure S4B ), and parasitized macrophages near cartilage ( Supplementary Figure S4E ). In the NB group, similar edema and neutrophil infiltration were seen, with many parasitized macrophages ( Figure 3F ). Respiratory epithelium showed slight morphological changes, including cellular stress and detachment ( Supplementary Figures S4C, F ). These findings indicate that Leishmania infection impacts the inner nasal mucosa in this model.

To assess immune responses to different nasal mucosa infections by Leishmania, we measured antibody levels. The NC group showed the highest IgM, IgG, and IgG1 levels, followed by NB and ED ( Figures 4A-F ). IgG2a and IgG2b were highest in NB, then NC ( Figures 4G-J ). Overall, NC and NB had stronger antibody responses than ED, while ST had the lowest. These findings suggest mucocutaneous infection triggers a stronger humoral response than ear dermis infection, with antibody profiles varying by inoculation site.

Flow cytometry of draining lymph nodes showed reduced CD4+ T cell frequency in the ED group but not in nasal infections. The NC and NB groups had increased total CD4+ T cells ( Figures 5A, E ) and higher numbers of cells expressing cytotoxic markers (CD107a, granzyme B, perforin) across NC, NB, and ED groups ( Figures 5B-D ), despite unchanged frequencies ( Supplementary Figures S6 - S8 ).

Although frequencies remained unchanged ( Supplementary Figures S9 , Supplementary Figures S10 ), total IFN-γ⁺ ( Figure 5F ) and IL-17⁺ ( Figure 5G ) cells increased in NC, NB, and ED groups. CD4+ IL-10+ T cells increased only in the ED group ( Figure 5H ), with lower IL-10-producing CD4⁺ T cell frequencies in nasal infections compared to ED ( Figures 6A, B ), highlighting distinct expansion dynamics between nasal and dermal infections.

We analyzed whether changes in CD4+ IL-10+ T cells were linked to CD4⁺ CD25+ FoxP3+ Tregs. Treg numbers increased in NC, NB, and ED groups without frequency changes ( Supplementary Figure S11 ). Similar trends were observed for CD25- FoxP3+, CD25+ FoxP3-, and CD25- FoxP3- cells ( Supplementary Figures S12 - Supplementary Figure S14 ), indicating the IL-10 reduction is unrelated to regulatory T cell frequency.

We also analyzed CD8⁺ T lymphocytes during mucosal infections. NC and NB groups showed an expansion in total CD8⁺ T cells ( Figures 7A, E ) with frequencies similar to CD4⁺ T cells ( Supplementary Figure S15 ). Cytotoxic CD8⁺ T cells markers increased in NC, NB, and ED groups ( Figures 7B-D ), with NC showing higher CD8⁺ Perforin⁺ frequency ( Supplementary Figure S18 ), nut no frequency changes to CD107a and Granzyme B respectively ( Supplementary Figure S16 , Supplementary Figure S17 ). There were no IFN-γ⁺ CD8⁺ significant changes in frequency ( Figure 7F ) or number ( Supplementary Figure S19 ). IL-17⁺ CD8⁺ T cells increased in NB and ED groups ( Figure 7G ) without frequency changes ( Supplementary Figures S20 ). CD8⁺ IL-10⁺ T cells showed no numerical change ( Figure 7H ), but frequency decreased in mucosa-infected and ED groups ( Supplementary Figure S21 ).