Bacille Calmette-Guerin (BCG) strain Pasteur 1173P2 and M. tuberculosis strain H37Ra were purchased from Shanghai Gene-Optimal Science and Technology (Gene-Optimal, China), and cultured on Middlebrook 7H11 agar plate or in 7H9 medium (BD Biosciences, USA) supplemented with 10% BBL Middlebrook OADC enrichment (BD Biosciences, USA). Six- to eight-week-old female specific-pathogen-free (SPF) C57BL/6 mice were purchased from the Experimental Animal Center of Sichuan University (Sichuan University, China) and housed under pathogen-free conditions with independent ventilation at West China Hospital, Sichuan University, China. The protocol of animal experiment was reviewed and approved by the Institutional Animal Care and Use Committee (IACUC), West China Hospital of Sichuan University (No. 20220228076).

The ag85B-rv2660c-mpt70 (arm) DNA segment was amplified from previously constructed pGAPZαA-arm recombinant plasmids (57) using specific primers (5’-CCGGAATTCCGTCCAGGTTTGCCAG-3’ and 5’-CCGCTCGAGAGCTGGTGGCATCAAAA-3’) that ligating to the incorporated EcoRI and XhoI restriction enzyme sites. The amplification employed PrimeSTAR Max DNA Polymerase (Takara, Japan) and followed the manufacture’s instruction. The amplified product was purified with a TIANgel Purification Kit (TIANGEN, China) and inserted into bacterial expression vector pET-28a (+) using non-ligase-dependent ClonExpress II One Step Cloning Kit (Vazyme, China) after digestion of EcoRI and XhoI (Takara, Japan). Recombinant pET-28a (+)-arm plasmids were confirmed by Sanger sequencing (Tsingke, China) and transformed into E. coli BL21(DE3) cells (Tsingke, China) by heat shock. Protein expression was induced with 0.5 mM IPTG for 4 hours at 37°C, yielding recombinant proteins as inclusion bodies. Precipitate was washed and dissolved in 8 M urea buffer (Buffer A) before purification using Ni²+ affinity columns (Bio-Rad, USA). Purified proteins were dialyzed against storage buffer (25 mM NaH₂PO₄, pH 8.0, 10% glycerol, 150 mM NaCl, 0.05% Tween20), quantified, and analyzed for purity using SDS-PAGE and western blot.

C57BL/6 mice were infected with 1 × 10⁷ CFU of M. tuberculosis H37Ra in 0.1 mL phosphate buffered saline (PBS) via tail vein injection. Uninfected mice were inoculated with the same volume of PBS as a naive control. Four weeks later, the infected mice were randomly divided into three groups. The PBS group received 2 doses of 100 μL PBS with 30 μg CpG ODN 1826 (Invivogen, USA) intramuscularly at two-week intervals; The ARM group received 2 doses of 10 μg ARM protein in 100 μL PBS with 30 μg CpG ODN 1826 intramuscularly at two-week intervals; The BCG group received 1 dose of 1 × 10⁶ CFU BCG in 100 μL PBS subcutaneously along the midline on the back. The intramuscular injection site was at the posterior thigh blank muscle. The body weight was monitored weekly and reported as the percentage of initial weight before the infection. To assess bacterial loads, mice were sacrificed at indicated time intervals. The lung and spleen were removed and homogenized in 1 mL PBS using gentleMACS M tube (Miltenyi, Germany). The homogenized tissue suspension was serially 10-fold diluted and plated on 7H11 agar plates supplemented with 10% OADC. Colony-forming units (CFU) were counted after incubation at 37°C for three weeks and reported as log₁₀ CFU.

Lung and spleen were collected from mice and processed into single-cell suspensions using GentleMACS dissociators and 70-μm cell strainer (Miltenyi, Germany). The mononuclear cells were isolated using Ficoll (Cytiva, USA) gradient centrifugation. After washing, cells were seeded into U-bottom 96-well plate (Thermofisher, USA) at 2×10⁶ cells/well and cultured with 20 μg/mL ARM or 25 μg/mL H37Rv whole-cell lysate (WCL, Gene-Optimal, China) in the presence of 1 μg/mL anti-CD28/CD49d (eBioscience, USA) at 37°C in 5% CO₂ for 3 hours. The culture was added with GolgiStop (eBioscience, USA) at final concentration of 1 μg/mL and incubated for another 11 hours. Cells were harvested and washed twice in cold Dulbecco’s modified phosphate-buffered saline (DPBS), then stained with surface marker antibodies diluted in 50% brilliant stain buffer (BD Horizon, USA) and fixable viability stain 700 (BD Biosciences, USA) (1:1000) at 4 °C for 20 min. After washing, cells were fixed and permeabilized with the Cytofix/Cytoperm Fixation/Permeabilization Kit (BD Biosciences, USA) according to manufacturer’s instructions, followed by intracellular cytokine staining for IFN-γ, TNF-α, IL-2, and IL-17A at 4 °C for 30 min. Data were collected using a FACSymphon A5 (BD Biosciences, USA) and analyzed using FlowJo v10.6.2 (BD FlowJo, USA). The following fluorochrome-conjugated antibodies were used for flow cytometric assessment: CD45-FITC (BioLegend, clone: 30-F11), CD3-APC/Cy7 (BD Biosciences, clone: 145-2C11), CD4-BV605 (BD Biosciences, clone: RM4-5), CD8-PerCP/Cy5.5 (BD Biosciences, clone: RPA-T8), CD44-BUV395 (BD Biosciences, clone: IM7), CD62L-APC (BioLegend, clone: MEL-14), CD69-BV510 (BioLegend, clone: H1.2F3), CD103-BV421 (BioLegend, clone: 2E7), IFN-γ-PE (BioLegend, clone: XMG1.2), TNF-α-PE/Cy7 (BioLegend, clone: MP6-XT22), IL-2-BV711 (BioLegend, clone: JES6-5H4), and IL-17A-BV785 (BioLegend, clone: TC11-18H10.1).

For antibody assay, polystyrene 96-well plates (Corning, USA) were coated with 1 μg/mL ARM or WCL protein in 100 μL Carbonate-Bicarbonate buffer (pH 9.6) overnight at 4°C. The plates were then washed three times with PBS containing 0.1% Tween-20 (PBST) and blocked with 3% BSA in PBST at 37°C for 2 hours. A volume of 100 μL serum sample in 5-fold serial dilutions, starting from 1:50, was added to the plate followed by incubation at 37°C for 1 hour. After washing, the plates were incubated with 1:5000 diluted HRP-conjugated antibodies (Proteintech, China) in 100 μL PBST for 1 hour and developed with 100 μL TMB substrate (Solarbio, China) for 5 minutes. The reaction was stopped with 100 μL 0.5 M H2SO4. The optical density (OD) was measured at 450 nm using a SpectraMax i3x microplate reader (Molecular Devices, USA). Endpoint titers were defined as the highest dilution with OD ≥ 2× negative control. For cytokine assay, serum TNF-α, IFN-γ, IL-2, and IL-10 were quantified using mouse cytokine ELISA kits (Ruixin, China) following the manufacture’s instruction.

H37Ra cells at density of OD₆₀₀ 0.5-1.0 in 1 mL were transferred to 100 mL 7H9 culture medium supplemented with 10% OADC (BD) and cultured for 2 weeks at 37°C. After centrifugation at 3000×g for 10 minutes, the pellet was resuspended in 10% glycerol and washed twice by centrifugation. Finally, the pellet was resuspended in 2 mL of 10% glycerol to prepare H37Ra competent cells. The H37Ra competent cells in 100 μL were loaded into a 2 mm Gene Pulser cuvette (Bio-Rad, USA) and mixed with 10 μL of pMV-261-eGFP plasmid (Gene-Optimal, China). Transformation was performed using the Gene Pulser Xcell™ (Bio-Rad, USA) at 2.5 kV/cm, 25 µF, 1000 Ω. Colonies were grown on 7H11 agar plates (BD Biosciences, USA) supplemented with 10% OADC enrichment, 0.5% glycerol, and 50 µg/mL kanamycin at 37°C for 35 days. Positive colonies expressing eGFP were verified by fluorescence microscopy and Ziehl-Neelsen acid-fast staining.

Murine macrophage Raw264.7 cells were suspended in DMEM medium (Gibco, USA) supplemented with 10% heat-inactivated fetal bovine serum (FBS, Gibco, USA), seeded into 96-well tissue culture plates at density of 1×10⁵ cells/well, and incubated at room temperature for 3 hours. The plate was gently washed with the culture medium to remove untouched cells and incubated for another 2 hours at 37°C in DMEM medium supplemented with 10% heat-inactivated mouse serum samples in duplicate wells. H37Ra-eGFP was added at a multiplicity of infection of 20 into each well and incubated for 3 hours. The plate was washed with PBS and dissociated the monolayer cell with 0.04% EDTA-PBS. The Raw264.7 cells with phagocytosed H37Ra-eGFP were detected with by flow cytometry.

For assessing inhibitory effect of macrophages on mycobacteria, Raw264.7 cells were cultured with H37Ra with a MOI of 10 in the presence of mouse serum samples as described above and incubated for 3 hours. After washing, the Raw264.7 cells were treated with 100 µg/mL gentamycin for 1 hour to eliminate extracellular bacteria. For cells harvested on day 1, 0.05% SDS was added to lyse the cells, and the lysates were serially 10-fold diluted and plated on Middlebrook 7H11 agar plates. For cells harvested on day 4, 100 µL of DMEM medium containing 10% mouse serum was added to each well after un-engulfed bacteria were removed on day 1. Supernatants were pooled, and the cells were lysed and plated on 7H11 agar plates on day 4. Colony-forming units (CFUs) were counted after incubating the plates at 37°C for 3 weeks, with a second count performed at 4 weeks to check for additional visible colonies.

Lung tissues were fixed in 4% paraformaldehyde, embedded in paraffin and sectioned. The section samples were stained with hematoxylin and eosin and examined under microscope (Olympus, Japan). Histopathological changes in lung were evaluated by blinded investigators using a scoring system (62) based on inflammatory cell infiltration (scale 0-3), alveolar structure destruction (scale 0-3), hemorrhage or vascular congestion (scale 0-2), and fibrosis (scale 0-2). The sum of respective scales with a total of maximum 10 represents severity of pathology.

GraphPad Prism version 9.0 was used for statistical analysis and graphical presentation of data. Data were displayed as mean ± SD. One-way and two-way ANOVA were used for group and subgroup analysis respectively. P values ≤ 0.05 were considered significant.

Our previous study reported the construction of a nucleic acid segment encoding the recombinant Mycobacterium tuberculosis Ag85B, Rv2660c, and MPT70 fusion protein (ARM) and its immunogenicity in mice (57). In this study, we constructed a pET-28a (+)-arm expression vector and expressed the recombinant ARM protein using E. coli BL21 (DE3). SDS-PAGE and western blot analyses confirmed the recombinant product as a 59 kDa fragment with specific 6×His-tag binding, consistent with predicted ARM characteristics, and the purity exceeding 94.5% ( Supplementary Figure S1A ). To establish a mouse model of Mtb infection, female SPF mice (6–8 weeks old) were infected with 10⁶, 10⁷, or 10⁸ CFU of H37Ra via tail vein injection. Weight changes were monitored weekly, and bacterial loads in the lungs and spleen were assessed on days 1, 7, 28, 42, and 84 post-infection. After infection, mice exhibited dose-dependent weight loss that persisted for at least 84 days without recovery compared to controls ( Supplementary Figure S1B ). Bacterial loads in the lungs and spleen peaked on day 42 and subsequently declined, correlating positively with the inoculation dose ( Supplementary Figure S1C ). Based on weight loss severity and pathogen persistence, 10⁷ CFU was selected as the optimal dose for the mouse model.

In this model, on day 28 post-infection (exposure), mice were treated with either PBS+30 μg CpG, 10 μg ARM+ 30 μg CpG, or 10⁶ CFU BCG, all in 100 μL PBS as described in the methods ( Figure 1A ). Mice treated with ARM and BCG stopped losing weight by day 35, one week after treatment. In contrast, PBS-treated mice continued to lose weight until day 56 post-infection. ARM-treated mice regained weight comparable to uninfected control, while BCG-treated mice maintained a lower but stable weight compared to both the uninfected control and ARM-treated mice ( Figure 1B ). Regarding mycobacterial load, both ARM- and BCG-treated mice showed significantly lower H37Ra loads in the lungs and spleen than PBS-treated mice. ARM-treated mice exhibited significantly lower H37Ra loads in the lungs on all tested days and in the spleen on day 56 compared to BCG-treated mice. Additionally, ARM-treated mice demonstrated lower H37Ra loads in the spleen on days 70 and 84, although these reductions were not statistically significant compared to BCG group ( Figure 1C ).

On day 56 post-infection, lung histological analysis of PBS-treated mice revealed widespread alveolar collapse, mononuclear cell infiltration in the parenchyma, thickened alveolar septa, inflammatory cell aggregates adjacent to bronchioles and vasculature, and patchy hemorrhage in the alveolar space. In contrast, post-infection treatment with ARM and BCG markedly alleviated pathological inflammation, as indicated by reduced alveolar destruction, decreased inflammatory infiltration, thinner alveolar septa, and the absence of hemorrhage. Among these groups, ARM-treated mice exhibited less severe pathological inflammation than the BCG group ( Figure 2A ). These differences became more pronounced on day 84 relative to day 56 ( Figure 2B ). Pathological severity was quantitatively evaluated by blinded investigators using an arbitrary scoring system ( Figure 2C ).

T cell responses, particularly CD4+ T cell response, are crucial for controlling M. tuberculosis infection (16). To investigate the impact of ARM on Mtb-specific T cell responses, mononuclear cells were isolated from the lungs and spleen on day 28 post-immunization (day 56 post-infection) and stimulated in vitro with H37Rv whole-cell lysate proteins. Cytokine expression was detected using intracellular cytokine staining and quantified by flow cytometry. Activated CD44+ T cells were gated and the cytokine production was assessed as shown in Supplementary Figure S2A . The frequency of CD44+ cells among total CD4+ T cells was significantly higher in ARM-treated mice compared to both PBS and BCG groups. While the proportion of CD44+ cells within the CD8+ T cell population was comparable to that in the BCG group but remained significantly higher than in the PBS group. ( Supplementary Figure S2B ).

Upon stimulation with H37Rv whole-cell lysate (WCL), lung CD4+CD44+ T cells from both BCG- and ARM-treated mice demonstrated significantly elevated frequencies of IFN-γ-, TNF-α-, IL-2-, and IL-17A-producing cells compared to the PBS group ( Figure 3A , left). Notably, ARM vaccination induced a significantly higher proportion of IFN-γ+ and TNF-α+ CD4+ T cells compared to BCG, suggesting a stronger Th1-type immune response. In terms of polyfunctional T cell responses, ARM-treated mice exhibited markedly increased frequencies of multi-cytokine-producing CD4+ T cells, especially those simultaneously expressing two or three cytokines, while BCG treatment mainly promoted the generation of single-cytokine-producing T cells ( Figure 3C ). The most common combination among CD4+ T cells was IFN-γ+TNF-α+, indicating coordinated cytokine responses ( Figure 3B , left). For CD8+CD44+ T cells ( Figure 3A , right), both BCG and ARM immunizations significantly enhanced the production of IFN-γ, TNF-α, IL-2, and IL-17A, with ARM again inducing a higher proportion of TNF-α+ CD8+ T cells than BCG. Consistent with CD4+ T cell data, ARM treatment also led to increased frequencies of polyfunctional CD8+ T cells, particularly double cytokine combinations, compared to PBS ( Figure 3D ). The most frequent combination in this subset was IL-2+IL-17A+ ( Figure 3B , right), highlighting a distinct cytokine co-expression pattern that may contribute to enhanced pulmonary immunity.

Likewise, spleen CD4+CD44+ T cells from both BCG- and ARM-treated mice exhibited significantly elevated frequencies of IFN-γ+, TNF-α+, and IL-2+ cells compared to the PBS group ( Figure 4A , left), with ARM induced a significantly higher proportion of IFN-γ+ and TNF-α+ CD4+ T cells than BCG, again indicating of a stronger Th1-type immune response. Further analysis of polyfunctional CD4+ T cell subsets ( Figures 4C ) demonstrated that both ARM and BCG increased the frequency of triple and double cytokine-producing cells compared to PBS, with ARM eliciting the highest levels of triple cytokine CD4+ T cells. Among these, the most prominent combinations included IFN-γ+TNF-α+IL-2+, IFN-γ+TNF-α+, and TNF-α+IL-2+ ( Figure 4B , left), underscoring coordinated Th1 cytokine co-expression. In the CD8+CD44+ T cell population ( Figure 4A , right), both BCG and ARM immunization significantly increased the overall frequency of cytokine-producing cells compared to PBS, although IL-2+ and IL-17A+ subsets remained relatively low and comparable among all groups. In line with CD4+ T cell trends, ARM and BCG also increased the frequency of polyfunctional CD8+ T cells, especially double cytokine producers ( Figure 4D ). While single cytokine-producing cells predominated across all groups, their frequencies were markedly elevated in both vaccinated groups, with the highest levels observed in ARM-treated mice for CD4+ T cells. These findings collectively suggest that ARM immunization enhances both the magnitude and functional quality of systemic T cell responses.

To evaluate the systemic cytokine milieu following immunization, serum levels of key Th1 cytokines (IFN-γ, TNF-α, IL-2) and the regulatory cytokine IL-10 were measured over time using ELISA ( Figure 5 ). Both BCG and ARM treatments significantly increased the concentrations of IFN-γ, TNF-α, and IL-2 compared to the PBS control across all time points. Notably, ARM treatment led to an earlier peak in IFN-γ levels at day 56, which was significantly higher than both the PBS and BCG groups. In contrast, the BCG-induced IFN-γ response peaked later, at day 70, but remained elevated through day 84. For IL-2, both BCG and ARM groups exhibited significantly elevated serum levels from day 56 onward, maintaining high concentrations through day 84. The PBS group displayed only a modest increase in IL-2 and remained significantly lower than both vaccine groups throughout the study period. Regarding IL-10, both BCG and ARM immunizations triggered a marked increase by day 56 compared to PBS. However, while BCG maintained high IL-10 levels over time, ARM-treated mice demonstrated a gradual decline in IL-10 concentrations post-day 56. This decline suggests that ARM may promote the resolution of inflammation more efficiently than BCG. Importantly, the temporal kinetics of serum Th1 cytokine production closely mirrored the dynamics of antigen-specific T cell responses observed in the lungs and spleens, particularly at day 56. The coordinated rise in serum cytokines and the frequency of cytokine-producing T cells suggests that ARM immunization not only enhances cellular immune responses but also modulates systemic cytokine profiles, potentially contributing to a balanced and effective protective immunity.

Although the protective role of humoral immunity against intracellular pathogens remains under debate (23, 24), accumulating evidence supports antibody-mediated immunity against Mtb (25, 63–65). To investigate this, we assessed the humoral immune responses in Mtb-exposed mice subjected to different immunizations ( Figure 6 ). ARM immunization led to a robust increase ARM-specific IgG titers, with a sharp rise following the first and second doses and peaking around day 49 ( Figure 6A ). ARM immunization markedly increased IgG (predominantly IgG2c) and IgM levels ( Supplementary Figure S3A ). Endpoint titers revealed that Ag85B-specific IgG was the most abundant ( Supplementary Figure S3B ). In contrast, neither PBS nor BCG induced detectable ARM-specific antibodies.

To evaluate the broader antibody response against Mtb, H37Rv-WCL-specific IgG levels were measured ( Figure 6B ). Although WCL-specific IgG levels peaked in all groups at day 56, the ARM group exhibited significantly higher titers compared to BCG, indicating an enhanced humoral response. To determine the functional capacity of these antibodies, we performed antibody-dependent phagocytosis assays using day 56 sera. RAW264.7 cells incubated with sera from ARM-treated mice exhibited a substantial increase in mean fluorescence intensity (MFI) of H37Ra-eGFP ( Figure 6C ), indicating enhanced phagocytosis. This effect was markedly diminished upon Fcγ receptor (FcγR) blockade, highlighting the crucial role of the antibody-FcγR pathway in promoting Mtb phagocytosis ( Figure 6D ). We next assessed the impact of antibody responses on bacterial killing. Sera from ARM-treated mice led to significantly higher Mtb uptake at day 1, as shown by increased CFU counts ( Figure 6E ), consistent with enhanced phagocytosis. Growth rates, calculated as CFU (day 4-day 1)/day 1, showed that sera from ARM-treated mice significantly inhibited Mtb growth compared to BCG and PBS sera ( Figure 6F ).

Given the complex composition of serum from Mtb-exposed mice, we conducted additional functional assays using sera from non-Mtb-exposed mice immunized with PBS, BCG, or ARM at the same time points to further validate these findings. Phagocytosis assays revealed that both BCG and ARM sera enhanced Mtb uptake in a dose-dependent manner, with ARM showing superior activity ( Figure 6G ) CFU assays showed that ARM sera significantly restricted Mtb growth ( Figures 6H–I ), reinforcing the notion that ARM immunization generates functional antibodies capable of promoting phagocytosis and limiting intracellular bacterial replication.

Phagocytosis followed by phagosome–lysosome (P–L) fusion is a critical step in macrophage-mediated elimination of Mtb, facilitating bacterial degradation within the phagolysosome (66). To further investigate the mechanisms underlying enhanced bacterial clearance, we assessed P–L fusion in RAW264.7 macrophages treated with sera from PBS-, BCG-, or ARM-immunized mice. Confocal microscopy revealed markedly increased colocalization of H37Ra-eGFP (green) with LysoTracker-labeled lysosomes (red) in the ARM group, as evidenced by the pronounced yellow signal in the merged images ( Figure 7A ). Quantitative analysis confirmed a significantly higher percentage of colocalization in the ARM group compared to both PBS and BCG controls ( Figure 7B ), indicating more efficient P–L fusion. These results suggest that antibodies induced by ARM immunization enhance both Mtb uptake and subsequent lysosomal fusion, contributing to improved intracellular bacterial clearance.