C57BL/6 mice, 6–8 weeks were procured from the CSIR-Institute of Microbial Technology (IMTECH), Chandigarh, India.

All the experiments were approved by the Institutional Animal Ethics Committee of the IMTECH and performed according to the National Regulatory Guideline issued by Committee for the Purpose of Supervision of Experiments on Animals (No. 55/1999/CPCSEA), Ministry of Environment and Forest, Government of India.

Fecal samples (200–300 mg) were collected aseptically in 1 ml of 1× PBS. Samples were homogenized and supernatant were collected after centrifuging samples at 2000 rpm for 2 min to pellet down debris. Later, serial dilutions were made and supernatants (100 μl) were plated on different media to culture both aerobic and anaerobic microbes. To cultivate anaerobic microbes, plates were kept in vacuum tight container in the presence of anaerobic gas pack (Himedia, Mumbai, India) for overnight.

To assess the bacterial diversity, supernatant from fecal samples was plated on different media to culture both aerobic and anaerobic microbes for overnight, as described above. The identification of colony morphotypes was carried out using four parameters: colony size, form, color, and texture. A phenotypic variant was considered when it differed in at least one of the referred morphological parameters. Diversity was calculated by counting the total number of different colonies of bacteria grown on different media plates. Total different colonies from the feces of healthy animals were considered as a reference number. Decrease in the bacterial diversity was calculated as (reference number − total different colonies calculated from infected or pre-antibiotics or post-antibiotics) × 100/reference number.

Mice were pre-treated with vancomycin (100 mg/l), polymixin B (60 mg/l), carbenicillin (50 mg/l), trimethoprim (20 mg/l), and amphotericin B (50 mg/l) ad libitum in drinking water for 21 days. The water containing antibiotics were replaced on every 3 days. Control mice were fed antibiotics free water. After 21 days, mice were challenged with Mtb (H37Rv) with deposition of 100 CFU in the lungs. The animals were administered antibiotics in the water for additional 21 days. Later, the mice were sacrificed, and tissues were harvested aseptically for immunological, microbiological, and histopathological studies. Pictorial representation of methodology is embedded in Supplementary Material.

Mice were challenged with Mtb with the deposition of 100 CFU in the lungs. After 21 days, mice were treated with antibiotics for subsequent 21 days with replacement of water every 3 days. Later, mice were sacrificed, and tissues were harvested aseptically for immunological, microbiological, and histopathological studies. Pictorial representation of methodology is embedded in Supplementary Material.

Antibiotics-treated mice underwent FT to reconstitute the gut composition. Fecal samples (200–300 mg) from five healthy mice were collected aseptically in five separate tubes in 1 ml of 1× PBS. Samples were homogenized and supernatant was collected after centrifuging samples at 2000 rpm for 2 min to pellet down debris. Supernatant slurry was collected and pooled together, and 100 μl was gavaged into mice within 15–20 min of excretion. Five doses with a gap of 3 days interval were orally administered to mice prior to 15 days of sacrificing animals. Pictorial representation of methodology is embedded in Supplementary Material as Methodology 2.

Spleen was harvested, and single-cell suspension was prepared. Briefly, lymphocytes from spleen were prepared by lysing RBCs with ACK lysis buffer (NH₄Cl 0.15M, KHCO₃ 10 mM, EDTA 88 mM), washed 3× with PBS, and resuspended in RPMI-1640-FBS-10%. Viability of the cells was assessed by trypan blue dye-exclusion method. The experiments were performed to detect intracellular cytokines and Foxp3 expression on T cells. To detect cytokines, splenocytes were restimulated in vitro with purified protein derivative (PPD). Splenocytes stimulated with PPD (20 μg/ml) were cultured for 48 h at 37°/CO₂ (5%). During last 4 h, cells were incubated with phorbol 12-myristate 13-acetate (PMA; 20 ng/ml) and ionomycin (1 μM) plus brefeldin A (5 μg/ml) for 2 h. The cells were stained with anti-CD4 Abs. After surface staining, cells were washed and resuspended in permeabilization-fixation solution (BD Cytofix/Cytoperm kit; BD Pharmingen, San Diego, CA, USA), and intracellular cytokine staining was performed with fluorescence-labeled Abs to TNF-α, IFN-γ, according to manufacturer’s protocol. FoxP3 staining was performed (Foxp3/Transcription Factor Staining Buffer Set, Ebioscience) according to manufacturer’s instructions. Data were collected using FACS Aria and analyzed using the BD DIVA software.

Statistical analysis was done using unpaired Student’s t-test and one-way ANOVA for group analysis using the Graph pad prism software 6.

Recent studies have shed new light on an impact of antibiotics on the gut microbes, which eventually may affect the severity of disease. Hence, we were curious to explore the influence of altered gut microbiota on the progression of TB. To test this hypothesis, mice were fed with broad spectrum antibiotics, prior to Mtb infection (pre-antibiotics) (for detailed methodology, please see Supplementary Material). Broad spectrum antibiotics were selected for the study. These antibiotics exhibited no impact on Mtb recovery in a Middlebrook 7H11 agar formulation (11). Animals treated with antibiotics for initial 5 days showed significant (p < 0.001) decrease in the CFU of the gut microbiota (Figure 1A). In contrast, when the antibiotics treatment was prolonged for 42 days, significant (p < 0.001) elevation in the number of microbes was observed (Figure 1A). Similar results (p < 0.01) were observed in the case of mice prior challenged with Mtb and then treated with antibiotics (post-antibiotics) (Figure 1B). Results suggest that initially antibiotics sensitive bacteria were eliminated, which resulted in the decline of CFUs (Figure 1A). However, later antibiotics resistant microbes predominated by over proliferation. However, we observed significant (p < 0.01) decrease in the diversity of the microbial taxa (Figure 1C), since the majority of the colonies were of sister clones, as identified through their morphology. Furthermore, we noticed enlargement in the size of cecum (Figure 1D). This change in the intestine was due to the decreased ability of antibiotics-treated mice to digest the food, as it was apparent by the presence of indigested food in the intestine. Similar results were observed in post-antibiotics-treated Mtb animals (Figure 1D).

The aim of the study was to assess the consequence of antibiotics driven alteration in the gut microbiota on Mtb survival in Mtb challenged mice. Interestingly, we observed that disruption of microbiota in pre-antibiotics (p < 0.05) and post-antibiotics (p < 0.01) treatment, significantly enhanced the growth of Mtb in the lungs of the infected animals (Figures 2A,B). This information was further corroborated with histopathological analysis of the lungs (Figure 2C). Mtb-infected mice on pre-antibiotics treatment showed larger and greater number of granulomas in their lungs, compared to control (Mtb infected, but not treated with antibiotics) (Figure 2C, upper panel). In addition, we observed significant increase in the granulomatous or tuberculous region (p < 0.01) in lungs of pre-antibiotics Mtb-infected animals (Figure 2D). Similar trend was observed in the post-antibiotics-treated animals as shown in histopathological images (Figure 2C, lower panel).

Dissemination of Mtb from lungs to other parts of the body is a probable factor responsible for extra-pulmonary infection. Interestingly, significant increase in the bacterial load of Mtb was observed in the spleen (p < 0.01) and liver (p < 0.05) of pre-antibiotics Mtb-infected model (Figure 3A). Similarly, post-antibiotics infected mice showed greater Mtb burden in the spleen (p < 0.01) and liver (p < 0.01), as compared to controls (Figure 3B). These data suggest that gut the microbiota play a crucial role in restricting the proliferation and dissemination of Mtb.

Before sacrificing animals, both pre-antibiotics and post-antibiotics-treated groups were fecal transplanted orally. Interestingly, we observed significant (p < 0.001) decline in the number of gut microbes in pre-antibiotics model after fecal transplantation and their number was comparable with normal mice (Figure 4A). Furthermore, there was significant (p < 0.001) but partial restoration in the microbial diversity, as noticed through morphology of microbes in FT group (Figure S1 in Supplementary Material). The variation was not resilient, i.e., did not return to its original percentage. It suggests that lowering of microbial diversity by antibiotics treatment could not be fully restored to its original frequency after FT for 15 days. It is reported that antibiotics treatment decreases the number of beneficial microbes (12). Therefore, we have confirmed the alteration in gut microbiota due to antibiotics treatment, by identifying microbes in the fecal samples by RT-qPCR. We observed significant increase in the number of Enterococcus (p < 0.05) but decline in the level of Bifidobacterium (p < 0.01), Lactobacillus (p < 0.05), Campylobacter (p < 0.05), and Bacteroides (p < 0.05) (Figure 4B). These results further support the partial reconstitution of gut microbial composition through FT. This information was further corroborated by PCR (Figure S2 in Supplementary Material). As compared to control animals (2.3 cm), antibiotics-treated group showed increase in the size of cecum (3.2 cm). Interestingly, FT restored the size of cecum to near normal (2.6) (Figure 4C). Furthermore, as compared to controls, the histopathological studies conducted on the ileum of pre-antibiotics-treated animals exhibited distorted structure of microvilli. The microvilli structure of ileum was reinstated to normal in the FT animals (Figure 4D). Similar results were observed in post-antibiotics model (Figure S3 in Supplementary Material). It was interesting observation that FT recovers antibiotics associated abnormities.

Since, we observed significant augmentation in the growth of Mtb in the lungs of pre-antibiotics (p < 0.05) and post-antibiotics (p < 0.01) groups (Figures 2 and 3), therefore we thought to study the influence of FT on pre-antibiotics and post-antibiotics animals infected with Mtb. Interestingly, FT significantly reduced the bacterial load in the lungs (p < 0.001) and spleen (p < 0.01) of both pre- and post-antibiotics mice exposed to Mtb (Figures 5A,B).

We also examined the progression of disease by studying the histopathological changes in the lungs. As compared to control group, larger size and number of granulomas in the lungs of pre- as well as post-antibiotics group were noted (Figures 5C,D). It was observed that antibiotics-treated mice after FT, exhibited granulomas with smaller size and number, lesser infiltration of lymphocytes, and consolidated lung architecture, compared to antibiotics group (Figures 5C,D). These results suggest that gut microbiome can contribute in controlling Mtb growth and its dissemination.

The role of Th1 cells is established in protection, whereas Tregs promote susceptibility to TB. Consequently, it was imperative to analyze the impact of antibiotics treatment on Tregs and Th1 cells and correlation in the modulation in their frequency with predisposition to TB. We observed substantial decline in the expression of IFN-γ and TNF-α upon treatment with antibiotics (Figures 6A,C). It was of interest to note the restoration of the production of IFN-γ and TNF-α in the mice with FT (Figures 2 and 3). Expression of IFN-γ in the spleen was further confirmed by RT-qPCR (Figure 6B). In contrast, antibiotics treatment augmented the frequency of Tregs, whereas FT downregulated their number as evidenced by the expression of Foxp3 (Figure 6D). This observation suggests that the antibiotics driven fluctuation in the gut microbiota can modulate the frequency of Tregs and Th1 cells, which may be responsible for proneness to TB.