Fourteen 3–4 months old female Pyrenean goats, from an officially tuberculosis-free herd, located in the Catalan Pyrenees (Spain), were chosen for this study. Experimental animals were transferred to the IRTA-CReSA Biosafety Level 3 (BSL-3) facility (Catalonia, Spain), where they were weighted and randomly divided into two experimental groups of seven animals each, challenged by the endobronchial (EB) and intranasal (IN) routes, respectively. According to body weights, corrections to ensure homogenous distribution between the groups were applied, and then, the animals were housed in two experimental boxes with animals 4–3 and 3–4 from the EB and IN groups in each box, respectively. The animals were fed hay, alfalfa, feed, and mineral salt and had access to water ad libitum throughout the experiment.

After 1 week of acclimatization, the animals were sedated intramuscularly with acepromazine maleate (0.05 mg/kg) and butorphanol (0.2 mg/kg) and subsequently anesthetized with intravenous administration of propofol (5 mg/kg) and midazolam (0.2 mg/kg). Afterward, the animals were challenged with 0.5 ml of a M. caprae field strain (Balseiro et al., 2017) at a suspension of 2 × 10³ CFU/ml. Animals of the EB group were challenged through the endobronchial route as previously described (Pérez de Val et al., 2011).

The animals of the IN group were challenged by using a syringe-adapted device for atomized spraying (MADgic^®, Wolfe Tory Medical, Inc., Salt Lake City, USA). This device consists of a flexible cannula measuring 21.6 cm in length and 4.82 mm in width with a nebulizer located at its end that generates droplets of 30–100 μm in diameter, according to the manufacturer's instructions. The cannula was inserted into the nasal cavity ~10–15 cm depending on the size of the animal. Each side of the nasal cavity received 0.5 ml of the same M. caprae strain at a suspension of 10³ CFU/ml without anesthesia.

After the challenge, all animals were daily monitored for clinical signs by a veterinarian to assess whether any of the animals reached the endpoint criteria. The clinical evaluation included 0–3 scoring of each of the following five parameters: body condition, mental state/behavior, weight loss, fever, and the presence of respiratory signs (0: absence of respiratory signs; 1: mild dyspnea and/or occasional cough; 2: moderate dyspnea and continued coughing; and 3: marked dyspnea, continued coughing, and/or severe nasal discharge). Any global score higher than 11 and/or the presence of qualitative criteria such as prostration, lack of movement, or severe respiratory distress resulted in euthanasia. Rectal temperature and body weight were measured each week. Heparinized blood samples were collected at weeks 0 (before challenge), 3, 5, 7, and 9 (endpoint) for immunological assays.

All the procedures with experimental animals carried out during the study were approved by the Animal Welfare Committee of the Generalitat de Catalunya (Project Ref. #10794), in accordance with the European Union legislation for the protection of experimental animals (86/609/CEE, 91/628/EEC, 92/65/EEC, and 90/425/EEC).

M. bovis tuberculin (PPD-B, 2,500 IU/ml) was obtained from CZ vaccines (Porriño, Spain). The specific recombinant antigens of MTBC ESAT-6, CFP-10, and MPB83 were obtained from Lionex (Braunschweig, Germany) at a concentration of 500 μg/ml each. ESAT-6 and CFP-10 were mixed 1:1 in an antigenic (E/C) cocktail. The P22 antigenic complex was produced by immunopurification of PPD-B (CZ vaccines) as described previously (Infantes-Lorenzo et al., 2017) and was supplied by the Carlos III Research Institute (Madrid, Spain) at a concentration of 500 μg/ml.

Approximately 10 ml of whole blood was collected at the abovementioned time points from the jugular vein using heparinized tubes. Blood samples were maintained at room temperature for <1 h when they were stimulated in 96-well cell culture plates (Eppendorf, Hamburg, Germany) with PPD-B, P22 (except at week 7), and E/C at a final concentration of 20 μg/ml each. Phosphate-buffered saline was used as an unstimulated control. The samples were incubated at 37°C and 5% CO₂ for 18 ± 2 h. Plasma supernatants were collected after centrifugation at 18 g for 10 min, and the released IFN-γ was measured by ELISA (ID Screen^® Ruminant IFN-g kit, ID.vet, Grabels, France). Optical densities of the ELISA plates were read at 450 nm (OD_450nm) using a spectrophotometer (Biotek Power Wave XS^®, Agilent, Santa Clara, USA). Antigen-specific IFN-γ responses were calculated as OD_450nm of the antigen-stimulated well minus OD_450nm of the unstimulated well (ΔOD_450nm).

Plasma samples from all experimental animals were analyzed in duplicate to follow antibody responses against MTBC after the challenge. Indirect ELISAs were used to detect total IgG against MPB83 antigen and P22 antigenic complex, respectively. The two ELISAs were performed as previously described (Pérez de Val et al., 2017; Infantes-Lorenzo et al., 2019). MPB83-IgG levels were calculated as OD_450nm of the antigen-coated well minus OD_450nm of the uncoated well (ΔOD_450nm). A sample was considered positive when ΔOD_450nm ≥ 0.05. P22-IgG levels were calculated as a percentage of ELISA (E%) = [mean OD_450nm of the antigen-coated well/(2 × mean negative control OD_450nm)] × 100. A sample was considered negative when E% ≤ 100%, non-conclusive when E% was between 110 and 150%, and positive (seroconversion) when E% ≥ 150%.

At week 9 post-challenge, all goats were sedated intramuscularly by an injection of acepromazine maleate (0.1 mg/kg) and butorphanol (0.2 mg/kg) and subsequently euthanized with an intravenous overdose of sodium pentobarbital. At necropsy, the size of TB-compatible lesions in the retropharyngeal (right and left), mediastinal (cranial and caudal), and tracheobronchial (LN) lymph nodes was measured, and the volume of lesions was inferred using the volume formula of the most similar 3D-geometrical morphology (including sphere, cylinder, or prism), as previously described (Balseiro et al., 2017). Other visible lesions in extrapulmonary tissues were also recorded and fixed in 10% buffered formalin to be confirmed by histopathology and Ziehl–Neelsen staining to detect acid-resistant bacilli. The lungs were fixed by airway instillation followed by immersion in formalin, as previously described (Pérez de Val et al., 2011), and then scanned by computed tomography (CT) using a 16-slice multidetector scanner (Brivo CT-385, General Electric Healthcare, Madrid, Spain). CT analysis was performed as previously described (Balseiro et al., 2017). In brief, total volumes of the lungs and TB lesions were measured using volume representation images with different density patterns (i.e., solid, mineralized, and cavitary lesions), and the total volume of mineralization was calculated using 100–300 Hounsfield units.

After macroscopic pathological evaluation, mediastinal and tracheobronchial LNs were collected and stored at −20°C until they were processed for bacteriology. The whole LNs were then thawed and homogenized in 10 ml of sterile-distilled water using a homogenizer (Masticator, IUL Instruments, Barcelona, Spain). An aliquot of 1 ml of each homogenate was inactivated at 75°C for 1 h for bacterial DNA quantification. In parallel, an aliquot of 100 ml of the master seed of M. caprae (~10⁸ CFU/ml), used for the challenged inoculum, was inactivated and then diluted in eight 10-fold dilutions to establish the standard curve. DNA samples were extracted using an ID Gene™ spin universal extraction kit (ID.vet) and amplified with the Mycobacterium tuberculosis complex duplex kit (ID.vet), following the manufacturer's instructions. The amplification was performed in a 7500 fast real-time PCR system (Applied Biosystems, Waltham, MA, USA). The genomic equivalents of CFU were calculated as previously described (Vidal et al., 2022).

A completely randomized design was performed to compare the effects of M. caprae infection (primary factor) by the EB and IN routes. First, all data distributions were analyzed by using the Shapiro–Wilk normality test. Temperature and body weight were compared using a unidirectional unpaired t-test. Antigen-specific immune responses (measured by IGRA, IgG-ELISA), lesion volumes, and mycobacterial DNA load were compared using the non-parametric Mann–Whitney test. GraphPad Prism version 8.0.0 (San Diego, CA, USA) was used for statistical analyses.

Mean antigen-specific IFN-γ responses in whole blood before and after the challenge for each treatment group are presented in Figure 1. An increase in IFN-γ-specific responses to PPD-B, P22, and E/C antigens was detected in both groups after the challenge (week 0, Figures 1A–C). Animals of the EB group showed a peak-up response of IFN-γ to the three antigens at weeks 3 and 5, which was significantly higher than in animals of the IN group (P < 0.01; except P < 0.05 for PPD-B at week 5). IFN-γ-specific responses in intranasally challenged animals increased progressively over weeks, while IFN-γ levels elicited after the endobronchial challenge decreased at the end of the experiment when responses in both groups tended to converge (no significant differences at weeks 7 and 9).

IgG levels to the MPB83 antigen and the P22 antigenic complex, as well as individual seroconversion, were studied using two ELISAs throughout the study (Figure 2). Endobronchial challenge induced an early increase in MPB83 IgG-specific responses (a peak-up response at weeks 5 and 7), and all EB animals seroconverted at week 5 p.c. On the contrary, only a mild response was detected in IN animals, seroconversion was not detected in 2 of them throughout the experiment, and the other five animals seroconverted between weeks 5 and 9 (Figure 2A). In comparison with MPB83, IgG responses to P22 were more progressive in the EB group, reaching a peak at the end of the experiment (week 9), but seroconversion was detected in six of seven animals. IgG responses to P22 were practically undetectable in the IN group throughout the experiment, and only one animal showed a clear seroconversion (Figure 2B).

Rectal temperature, body weight, and clinical signs of TB were weekly recorded throughout the experiment (Figure 3).

In the EB group, a peak of mean rectal temperature was observed between weeks 2 and 4 p.c., being statistically significantly higher compared with the IN group (P < 0.01, weeks 2 and 4; P < 0.05, week 3; Figure 3A). A mild peak was also observed in the IN group between weeks 5 and 7, although still remained slightly lower than the EB group, and decreased at week 8 to be significantly lower than the EB group again (P < 0.05).

The mean body weight gain was greater in the IN group compared with the EB group since week 4, being statistically significant at week 5 (P <0.05, Figure 3B), although a stagnation in body weight increase was observed from week 5 onward. In contrast, animals belonging to the EB group showed a mild increase in mean body weight from week 6 to the end of the experiment, where the difference in mean cumulative weight gain (ΔKg, Kg at weeks 9–0) between the IN and EB groups was minimal (4.2 ± 3.3 95% CI and 3.5 ± 2.9 95%CI, respectively).

The presence or absence of clinical signs, other than weight loss or fever, compatible with TB was recorded in all challenged animals until the end point of the experiment (Figure 3C). The observed signs included cough, dyspnea, eye discharge, enlarged lymph nodes, runny nose, and anorexia. An animal of the EB group showed clinical signs at week 1, 4 at week 2, and 6 from week 3 to the endpoint, whereas a 2-week delay in the onset of clinical signs was observed in the IN group: 1, 4, and 6 animals at weeks 3, 4, and from 5 to the endpoint, respectively (Figure 3C). The proportion of animals with clinical signs was significantly higher in the EB group at weeks 2 (P < 0.01), 3 (P < 0.01), and 4 (P < 0.05) compared with the IN group. All experimental animals showed clinical signs at weeks 8 and 9.

All goats showed TB lesions at necropsy. Lung lesions were quantitatively assessed by CT. All animals challenged by the EB route (7/7) had lung lesions, while only 4/7 animals of the IN group presented lung lesions, and the severity and dissemination of these lesions were very much lower compared with those found in the EB group. Figure 4 shows the extension of lung TB granulomas expressed as the volume of lung lesions, the intrapulmonary spread of TB lesions expressed as the number of affected lung lobes, and the degree of development of lung TB lesions expressed as the volume ratio between mineralization and lesion of each group. The EB group showed a significantly larger median volume of lung lesions compared with the IN group (P < 0.001, Figure 4A). In addition, the EB group showed a higher number of affected lobes compared with the IN group (P < 0.01, Figure 4B). Five out of seven EB animals showed TB lesions in all seven pulmonary lobes, while only one IN animal showed lesions in six pulmonary lobes. The EB group also showed a significantly greater median volume of lung mineralization (18.36 cm3, 95% CI: 2. 634–84.53, P < 0.001) compared with the IN group (0 cm3, 95% CI: 0–0.72), as well as higher median proportion of mineralization of lung lesions (P < 0.01; Figure 4C) indicative of a faster progression of pulmonary TB infection.

All intranasally challenged animals showed lesions in retropharyngeal (RF) LNs, while these lesions were only detected in 2/7 animals of the EB group, and the median volume of lesions in the RF LNs was significantly higher in the IN group compared with the EB group (P < 0.01, Figure 5C). In contrast, only 3/7 animals that received the IN challenge developed TB lesions in pulmonary LNs, whereas these lesions were macroscopically found in all EB-challenged ones. The median volume of TB lesions in pulmonary LN was significantly higher in the EB group compared with the IN group (P < 0.001, Figure 5A), and the bacterial DNA load in pulmonary LNs, estimated as CFU equivalents by qPCR, was also significantly higher in the EB group (P < 0.05, Figure 5B).

All extrapulmonary lesions macroscopically compatible with TB were confirmed as such by histopathological examination. In general, the IN group presented major affectation of the head, whereas more severe TB-compatible lesions in the thoracic and abdominal cavities were found in the EB group. Four out of seven animals of the IN group had an intense granulomatous inflammatory infiltrate in the nasal cavity (see Figures 6E, F), and the three animals had lesions in the retropharyngeal and/or submandibular lymph nodes.

Figures 6, 7 show examples of TB lesions developed in representative IN- and EB-challenged animals in the head and lungs, respectively. The longitudinal section of the head of intranasally challenged goats evidenced bilateral extensive multifocal to coalescent granulomatous lesions in the nasal cavity of 4/7 goats; the endoturbinates and the middle nasal concha (on the caudal aspect of the nasal cavity) were always the most affected, while the dorsal and ventral nasal concha and the nasal septum were more variably involved (Figure 6E). Microscopically, granulomas of variable extension were observed in the nasal turbinates (Figure 6F) and nasopharyngeal mucosa (Figure 6G), while the EB-challenged animals did not present visible lesions in this localization (Figures 6A, B). Lesions in the head lymph nodes were more severe in the IN group (Figure 6H) while minimal or absent in the EB group (Figure 6D). Microscopically, the lesional patterns observed were equivalent in both groups, differing only in their extension and location. On the other hand, the rendered images of the lungs obtained by CT disclosed more extensive lung lesions and a higher total volume of lung mineralization in the EB-challenged animals (Figure 7).

Figure 8 presents the distribution of TB lesions in all the animals of each group. The animals of the EB group have minimal involvement in the head structure whereas all animals of the IN group had lesions in the head (all of them in the retropharyngeal LNs). On the other hand, all animals of the EB group had lesions in the thorax but also in the abdominal cavity (especially in the spleen), indicative of early hematogenous generalization.