Immune activation and response dynamics of human iPSC‐derived macrophages in tuberculosis infection models

Abstract

Objectives

Tuberculosis (TB) remains a global health challenge, with current antibiotic therapies being limited by long treatments, side effects and multidrug‐resistant mycobacterial strains. In addition, Mycobacterium tuberculosis (Mtb), the main causative agent of TB, employs evasion mechanisms particularly within alveolar macrophages being the primary host cells. Conventional therapies fail to modulate macrophage function or effectively target host immunity, which is crucial in TB pathogenesis. Emerging evidence points to induced pluripotent stem cell‐derived macrophages (iMacs) with enhanced bactericidal activity as a promising cell‐based approach for TB treatment. Therefore, this study aimed to compare iMacs with blood monocyte‐derived macrophages (MDMs) in response to Bacillus Calmette–Guérin (BCG), the live attenuated TB vaccine and heat‐killed Mtb (HKMT).

Methods

iMacs and MDMs were challenged with BCG and HKMT to assess their functional responses. Key parameters evaluated included cell migration, phagocytosis kinetics, levels of autophagy‐ and apoptosis‐related proteins, and cytokine production profiles following infection.

Results

iMacs displayed enhanced migration, faster phagocytosis and increased expression of autophagy‐ and apoptosis‐related proteins compared with MDMs. Moreover, iMacs showed a stronger pro‐inflammatory cytokine response and rapid return to baseline cytokine levels post‐infection.

Conclusion

These findings support the potential of iMacs as an immunocompetent model for studying mycobacterial infections and as a tool for cell‐based TB immunotherapies.

In this study, we analysed the immune responses of blood‐derived monocyte macrophages and iPSC‐derived macrophages following BCG/HKMT infection. We found that iPSC‐derived macrophages displayed a markedly stronger activation profile, including faster migration, increased reactive oxygen species production, elevated apoptosis marker expression and enhanced pro‐inflammatory cytokine secretion. These results highlight iMacs as an additional robust model for studying enhanced innate immune reactivity upon tuberculosis infection.

Introduction

Tuberculosis (TB) remains a major global health problem, with 10.8 million new cases and ¹ , ² 1.09 million deaths reported in 2023 alone, making it the leading cause of death among infectious diseases. Despite extensive efforts, including the WHO End TB Strategy, TB‐related mortality has declined by only 23% since 2015—far below the 75% reduction target set for 2025. ³ A key factor in the pathogenesis of TB is the interaction between Mycobacterium tuberculosis (Mtb) and macrophages, which play a dual role acting as the first line defence and as host cells for intracellular bacterial propagation. ⁴ Macrophages utilise multiple anti‐bacterial mechanisms against Mtb ⁵ including phagocytosis and subsequent bacterial degradation, and the production of reactive oxygen species (ROS). In addition, autophagy facilitates the elimination of bacteria that evade direct killing, ⁶ , ⁷ while macrophage apoptosis contributes to pathogen eradication. ⁸ , ⁹ Cytokine secretion and intercellular signalling further regulate the immune response, thus influencing the overall infection outcome such as pathogen clearance, latency or active TB. Although TB is both preventable and treatable, current TB therapies face significant challenges. ¹⁰ , ¹¹ The standard 6‐month antibiotic regimen is often hampered by poor patient adherence because of the complexity of treatment and associated side effects, contributing to the rise of antibiotic‐resistant Mtb strains. ¹² , ¹³ Furthermore, immunodeficiencies such as HIV infection or TNF deficiencies, whether genetic or because of anti‐TNF therapies, significantly increase susceptibility to TB by impairing the body's ability to control Mtb infection, further increasing the risk of uncontrolled bacterial spread. ¹⁴ , ¹⁵ As a result, novel TB treatment strategies and preventive measures remain urgently needed. Recent clinical studies have set the focus on macrophage‐based cell therapies, as evidenced by the successful therapeutic application against liver fibrosis ¹⁶ , ¹⁷ and cancer. ¹⁸ Furthermore, the therapeutic use of macrophages has been expanded to the field of bacterial infections, demonstrating improved anti‐bacterial efficacy against both Gram‐positive ¹⁹ and Gram‐negative ²⁰ bacteria in preclinical studies.

Traditionally, primary macrophages such as blood monocyte‐derived macrophages (MDMs) have been the main source for macrophage‐based therapies, but recent research has shifted the focus towards induced pluripotent stem cells (iPSCs) as a standardised platform for generating macrophages (iMacs) in vitro. iMacs have demonstrated significant anti‐pathogenic potential in bacterial and viral infections including Staphylococcus aureus, ¹⁹ Pseudomonas aeruginosa, ²⁰ Chlamydia trachomatis ²¹ and Salmonella Typhimurium and S. Typhi, ²² underlining their utility as a valid model system, revealing their anti‐microbial activity compared to conventional cell lines (e.g. THP‐1) and primary macrophages. Recent studies have further expanded the therapeutical use of iMacs in mycobacterial research, particularly in TB, demonstrating their robust anti‐mycobacterial activity and suitability as translational disease model. ¹⁵ , ²³ , ²⁴ , ²⁵ Comparative studies between iMacs and THP‐1‐derived macrophages have revealed functional differences in key anti‐mycobacterial responses such as induction of apoptosis and TNF secretion. ²⁵ Additionally, studies comparing iMacs with MDMs have identified substantial differences in gene expression and key signalling pathways, ¹⁹ , ²⁶ emphasising the need for further investigation into functional variations among macrophage subtypes in TB. Moreover, a recent study successfully utilised iMacs as a high‐throughput drug‐screening platform to identify compounds that modulate TB host–pathogen interactions. ²³ Furthermore, preclinical studies using iMacs in murine bacteria infection models or hereditary pulmonary alveolar proteinosis (hPAP) have demonstrated functional adaptation, and disease‐modifying potential following intrapulmonary transfer of cells, making them a promising candidate for cell‐based therapies in different lung diseases. ¹⁹ , ²⁷ , ²⁸ Based on this background, we aimed to further elucidate the molecular and cellular mechanisms of iMacs as anti‐mycobacterial defence, particularly in comparison with primary macrophage models, to fully establish iMacs as a reliable TB model system and potential therapeutic application. A deeper understanding of host–pathogen interactions between different macrophage subtypes will also be crucial for the development of robust in vitro disease models, providing greater insight into TB pathogenesis and facilitating the exploration of novel prevention strategies.

Therefore, in this study, we conduct a comprehensive comparative analysis of the immune response of iMacs and MDMs to assess the potential of macrophage‐based cell therapies for TB treatment. For initial infection studies, we used Bacillus Calmette–Guérin (BCG), an attenuated strain from Mycobacterium bovis, a closely related species to Mtb, with Mycobacterium bovis sharing over 99% genetic identity with Mtb, ²⁹ triggering similar immune responses. While Mycobacterium bovis is primarily responsible for TB in cattle and deer, its attenuated derivative BCG can only infect and develop infectious disease in immunocompromised humans. ³⁰ For example, patients with Mendelian Susceptibility to Mycobacterial Disease (MSMD) are particularly vulnerable to BCG infections, often developing severe or widespread disease after exposure. ³¹ , ³² Although Mycobacterium bovis BCG shares many antigens with Mtb and elicits comparable immune responses, it is missing several crucial virulence factors—particularly those encoded within the Esx‐1 region—that are essential for full pathogenicity. ³³ These genetic deletions account for its attenuation and make it a safe model organism. As a result, BCG serves as a useful surrogate for investigating host–pathogen interactions and immune responses related to tuberculosis, thereby supporting the advancement of more effective preventive and therapeutic approaches. ³⁰ , ³⁴ , ³⁵ Following stimulation with a reporter BCG or the heat‐killed Mtb strain H37Ra (HKMT), our findings reveal that iMacs exhibit a more robust immune response than MDMs. This is characterised by faster phagocytosis, the induction of proteins related to autophagy and apoptosis, and a pronounced pro‐inflammatory cytokine response, suggesting their potential as a model for studying host‐mycobacteria interactions and potential cell therapy applications. These results further provide new insights into the unique defence mechanisms of iMacs and MDMs during mycobacteria infection.

Results

Generation and characterisation of iPSC‐ and blood monocyte‐derived macrophages

To investigate and compare the immune response of macrophages from two different cell sources upon mycobacterial infection, we generated macrophages either derived from iPSCs using a feeder‐free culture (iMacs) or blood‐derived monocytes isolated from buffy coat blood (MDMs) and performed infection studies using either BCG or HKMT (Supplementary figure 1a). To emphasise the conclusions and account for donor‐to‐donor variability, we analysed the immune response of multiple healthy MDM donors and two distinct healthy iPSC clones (Selene and NIH). Because of the similarity in outcomes, the iMac results are shown in a combined format in the following functional assays. Feeder‐free iMac production efficiency was comparable for both iPSC clones with an average total number of 7.71 × 10⁶ for Selene iMacs and 7.92 × 10⁶ for NIH iMacs per harvest of three plates, being stable for up to 10 harvests (Supplementary figure 1b). In addition, both generated macrophage cell types exhibited comparable basal anti‐bacterial phagocytic activity using E. coli bioparticles conjugates (Supplementary figures 1c and 2c). Furthermore, iMacs and MDMs shared a similar macrophage phenotype, characterised by a highly homogeneous population of CD45⁺CD11b⁺CD14⁺CD163⁺ cells, a similar expression of toll‐like receptors (TLRs) 2 and 4, important for the recognition of BCG and a comparable mixed expression profile of M1 and M2 markers, with surface marker expression levels exceeding 90% in both iMacs and MDMs (Supplementary figures 1d and 2d). To further validate the robustness and reproducibility of iMac differentiation, we compared the two iPSC clones (Selene and NIH) in more detail (Supplementary table 1). Cytokine secretion assays demonstrated comparable IL‐6 release upon IFN‐γ or LPS stimulation, confirming similar pro‐inflammatory responsiveness and reaction to bacterial mimetics (Supplementary figure 1e, f). Likewise, flow cytometric analysis showed similar expression of anti‐inflammatory marker CD206 after stimulation with IL‐4, revealing no significant differences between Selene and NIH iMacs (Supplementary figure 1g), indicating similar capacities for pro‐inflammatory (M1) and anti‐inflammatory (M2) polarisation. Finally, viability analysis of terminally differentiated iMacs using Zombie Aqua staining showed no significant differences between the two iPSC‐derived macrophage lines, confirming similar survival rates post‐terminal differentiation (Supplementary figure 1h).

Together, these results demonstrate that iMacs generated from independent iPSC clones display highly consistent differentiation efficiency, phenotype, functional polarisation capacity, and viability, supporting their reliability and reproducibility as a model for macrophage‐related infection studies.

iMacs demonstrate faster phagocytosis and increased pro‐inflammatory cytokine secretion upon BCG infection

To comprehensively examine the anti‐mycobacterial immune responses of iMacs and MDMs, we initially performed infection studies using BCG, an attenuated strain from Mycobacterium bovis. As a first step, we assessed the impact of BCG infection on two critical stages of phagocytosis—phagosomal acidification and maturation—both of which are essential for effective anti‐bacterial defence and are known targets of Mtb immune evasion strategies. Both Mtb and BCG are known to promote their survival within macrophages by arresting phagosomal acidification and maturation. Therefore, we first analysed the process of phagosomal acidification following BCG infection by assessing the recruitment and accumulation of vATPses at different time points by confocal microscopy (Figure 1a). Already as early as 5 min post infection, successful phagocytosis of BCG was observed in both iMacs and MDMs, as indicated by colocalisation of macrophages and BCG. After 30 min infection, iMacs exhibited a strong increase of phagocytosed BCG, along with a remarkable recruitment and accumulation of vATPases, which colocalised with the bacteria. In contrast, MDMs displayed a delayed bacterial uptake and acidification of vacuoles, persisting at 30 min. To complement these qualitative observations, we quantified the colocalisation of internalised BCG with vATPase. The analysis showed that the Pearson's correlation coefficient was significantly higher in iMacs than in MDMs 5 min post infection, and remained slightly higher at 30 min (Figure 1a, right panel). A higher colocalisation of BCG with vATPase indicates more rapid engagement of the phagosomal acidification machinery in iMacs, suggesting a faster initiation of the antimicrobial response.

Figure 1.

iMacs demonstrate enhanced phagocytosis and cytokine production upon BCG infection. Microscopic analysis of immunofluorescent images stained for (a) vATPase and (b) LAMP‐1 immunolabelling after infection with BCG (MOI 10:1) for various time points, using a Leica TCS SP5 confocal microscope (100× oil objective, scale bar = 10 μm). Right panels (a) + (b) show Pearson correlation coefficient demonstrating correlation between (a) BCG (green) and v‐APTase (red) accumulation and (b) BCG (green) and LAMP‐1 (red) accumulation at different time points post‐infection. The data are representative of n = 3 independent experiments and individual cells were quantified according to the program. (c) IL‐6, (d) TNF and (e) IL‐1β secretion by iMacs and MDMs 6 h and 24 h post BCG infection with an MOI of 10:1, compared to non‐infected cells as controls measured by ELISA. Data (c)–(e) are represented as the mean ± SD of n = 4–5 biological replicates measured in duplicates. Statistical significance was determined using Mann–Whitney U test (* indicates P < 0.0332, ** indicates P < 0.0021); n.i., non‐infected control cells.

To further quantify phagosomal acidification, Western blot analysis was performed to assess vATPase recruitment and accumulation. The results demonstrated a tendency toward higher vATPase levels in iMacs, with an average overall increase of 2.2‐fold, compared to a 1.1‐fold increase in MDMs, aligning with the respective confocal microscopy findings (Supplementary figures 2a and 1f). Despite these differences in kinetics, both cell types exhibited intact phagosomal acidification, with no apparent interference by BCG. Next, the phagosomal maturation was investigated, by staining for lysosome‐associated membrane protein 1 (LAMP‐1) as a recruited marker for late phagosomes and lysosomes, comparing iMacs and MDMs (Figure 1b). Similar to acidification kinetics, iMacs showed a faster uptake of BCG and an increased accumulation of LAMP‐1, with a well‐defined centralisation over time, compared to the delayed reaction observed in MDMs, both after 5 and 30 min post infection. Quantitative colocalisation analysis of BCG and LAMP‐1 revealed significantly higher colocalisation in iMacs than MDMs 5 min post infection, while no significant difference was detected at 30 min (Figure 1b, right panel). This higher early colocalisation of BCG with LAMP‐1 in iMacs suggests accelerated phagosomal maturation, which may contribute to more efficient bacterial processing. Nevertheless, LAMP‐1 expression in both macrophage types confirmed an intact phagosome maturation process, which is crucial for effective bacterial clearance. To further characterise the immune response, the cytokine secretion profile was analysed in both macrophage types, following BCG infection at an MOI of 10:1. After 6 h, both iMacs and MDMs showed elevated levels of IL‐6 secretion with an average of 6648 pg/mL (± 738.90 pg/mL) and 1485 pg/mL (± 968.70 pg/mL), respectively, confirming an effective cytokine response following BCG uptake (Figure 1c). However, 24 h post infection, iMacs displayed a markedly greater increase in IL‐6 secretion, reaching an average of 13 680 pg/mL (± 594 pg/mL), compared to 4425 pg/mL (± 35.36 pg/mL) in MDMs, suggesting a more rapid and robust anti‐bacterial response. A similar pattern was observed for TNF secretion (Figure 1d). Both iMacs and MDMs exhibited a strong increase in TNF levels after 6 h of BCG infection. However, while MDMs maintained TNF levels at 24 h, iMacs displayed a marked reduction in TNF secretion, suggesting a rapid self‐regulation mechanism. Given that TNF plays a critical role in macrophage activation, the induction of cell death, TB control and host defence, ¹⁵ , ³⁶ its modulation is essential for preventing excessive immune activation and potential granuloma formation. These findings suggest that iMacs exhibit a tightly controlled inflammatory response, rapidly returning to baseline activation following BCG uptake.

To gain a broader perspective on the secretome, a Luminex bead‐based multiplex assay was performed to quantify a range of pro‐ and anti‐inflammatory cytokines. In agreement with the ELISA results, at 24 h post BCG infection, iMacs exhibited higher levels of key pro‐inflammatory cytokines, including TNF, IL‐6, IL‐1β, IL‐12 and IL‐23, all of which are known to play crucial roles in host defence against bacterial infections ³⁷ , ³⁸ , ³⁹ (Supplementary figure 2b). Additionally, iMacs demonstrated a stronger increase of anti‐inflammatory cytokine levels such as IL‐10 and IL‐33, which are essential for resolving inflammation and preventing excessive immune activation, ⁴⁰ , ⁴¹ which is in line with the regulated TNF secretion observed in iMacs.

In summary, our findings demonstrate that both iMacs and MDMs effectively internalise BCG and undergo normal phagosomal acidification and maturation, with no major disruptions because of BCG interference. However, iMacs exhibit a more rapid and robust immune response, characterised by accelerated phagocytosis, enhanced pro‐inflammatory cytokine secretion and efficient self‐regulation, while MDMs display a comparatively delayed immune activation.

BCG infection potently stimulates rapid induction of autophagy‐ and apoptosis‐related proteins in iMacs

To further investigate the cellular responses upon BCG infection, we first examined the ability of iMacs and MDMs to induce xenophagy‐related proteins, which is a selective form of autophagy, an important intracellular degradation mechanism, leading to the suppression of intracellular mycobacterial survival. ⁶ Both cell types were infected with an MOI of 10:1 BCG for various time periods, and the expression of two key autophagy‐associated proteins, Beclin‐1 and microtubule‐associated protein 1A/1B light chain 3B (LC3B), was analysed. Beclin‐1 regulates autophagosome formation, ⁴² while LC3B marks autophagy progression. ⁴³ Densitometric analyses of Western blot assays revealed a rapid induction of Beclin‐1 expression in iMacs, detectable as early as 5 min post infection, peaking at a threefold increase at 30 min, before gradually declining over time relative to non‐infected controls. In contrast, MDMs exhibited a delayed and more modest Beclin‐1 response, reaching a maximum 1.7‐fold increase at 60 min post infection (Figure 2a; Supplementary figure 1a). Similar trends were observed in LC3B expression (Figure 2b; Supplementary figure 1b). Analysis of total LC3B expression revealed an early induction in both iMacs and MDMs at 5 min post infection, with an average increase of 1.7‐fold (± 0.49) and 1.1‐fold (± 0.1), respectively (Supplementary figure 2c). However, while LC3B expression in iMacs continued to rise, peaking at 3.4‐fold at 120 min, MDMs exhibited a progressive decline in LC3B levels over time. A comparable pattern was observed in the assessment of autophagosome accumulation, analysing the expression levels of LC3BII (Figure 2b). LC3BII is a lipidated form of LC3B that integrates into autophagosomal membranes, serving as a marker to evaluate differences in autophagosome levels. While MDMs displayed only a modest increase over time, iMacs demonstrated a significant increase in autophagosome accumulation already 5 min post infection with an average of 2.4‐fold (± 1.20), further increasing over time, peaking at a maximum of 5.5‐fold.

Figure 2.

BCG infection leads to increased expression of autophagy‐ and apoptosis‐related proteins in iMacs. Representative sample of Western blot analyses and respective densitometric analysis investigating expression of (a) Beclin‐1, (b) LC3B, (c) p38 phosphorylation, (d) c‐Jun N‐terminal kinase (JNK) phosphorylation and (e) Bax expression in iMacs and MDMs following BCG infection with an MOI of 10:1 at various time points. Data (a)–(e) are presented as mean ± SD of at least three independent experiments, presented as fold change relative to non‐infected cells as control. Statistical significance was determined using Mann–Whitney U test (* indicates P < 0.0332).

These cells were then further examined for the mitogen‐activated protein kinase (MAPK) pathway, a key regulator of intracellular signalling cascades in immune responses. ⁴⁴ To assess pathway activation, we investigated the phosphorylation of p38 mitogen‐activated protein kinase (p38) and c‐Jun N‐terminal kinase (pSAPK/JNK) via Western blot analysis at various time points post‐BCG infection, relative to non‐infected controls. Phosphorylation of p38 was highly upregulated in iMacs, reaching an average 27.33‐fold (± 18.08) increase at 30 min post infection, whereas MDMs exhibited a lower response, with an average 9.83‐fold (± 5.03) increase (Figure 2c; Supplementary figure 1c). In addition, pSAPK/JNK expression followed a similar pattern in both iMacs and MDMs, peaking at 30 min before declining over time (Figure 2d; Supplementary figure 1d). To evaluate the induction of proteins related to apoptosis, a critical programmed cell death mechanism involved in bacterial clearance, we analysed the expression of the pro‐apoptotic protein Bax using Western blot assays (Figure 2e; Supplementary figure 1e). Our findings indicated a tendency towards stronger upregulation of Bax expression in iMacs, reaching an average 2.60‐fold (± 2.26) increase at 120 min post infection, whereas MDMs exhibited a decrease in Bax levels over time, reaching an average 0.77‐fold (± 0.30) at 120 min (Figure 2e).

In conclusion, these findings reveal distinct differences in the autophagic‐ and apoptotic‐associated pathways in iMacs and MDMs following BCG infection. iMacs exhibit a more rapid and robust activation of key anti‐mycobacterial pathways, including autophagy‐ and apoptosis‐related proteins, compared with MDMs, suggesting an enhanced capacity for intracellular pathogen control.

Enhanced immune response of iMacs to heat‐killed Mycobacterium tuberculosis compared to blood monocyte‐derived macrophages

As a second candidate within the mycobacterial group, we evaluated the immune response of iMacs and MDMs to heat‐killed Mycobacterium tuberculosis (HKMT), an avirulent H37Ra strain that lost its replicative capacity upon the heat‐killing process. Despite its inability to replicate, HKMT remains biologically active and retains the ability to bind to and activate immune cells. Initial microscopic evaluation of HKMT‐stimulated iMacs and MDMs revealed differences in cell migration and cluster formation following HKMT exposure (Figure 3a'). Whereas non‐infected NIH and Selene iMacs displayed uniform distribution and an elongated morphology in cell culture, HKMT stimulation led to an active migration and subsequent cluster formation of bacteria and macrophages within 24 h of stimulation. Live‐cell imaging further demonstrated a dynamic targeted migration by iMacs toward the bacteria, pseudopodia formation and progressive clustering over time (Supplementary video 1). Similarly, MDMs displayed cluster formation 24 h after HKMT stimulation, but iMacs showed a more pronounced and rapid response compared with MDMs. To assess whether the observed cluster formation corresponded to enhanced phagocytic activity, the percentage of phagocytic macrophages was quantified over a time course ranging from 5 min to 24 h. HKMT was pre‐labelled with a live red dye for accurate tracking within the macrophages (Figure 3b; Supplementary figure 2a). iMacs exhibited markedly faster phagocytosis rates than MDMs during the initial 5 to 30 min post stimulation with a significantly higher average of 45.31% (± 23.92%) phagocytosed HKMT at 5 min, compared to 8.42% (± 7.81%) by MDMs. However, MDMs achieved comparable levels of phagocytosis after 1 h, remaining stable over subsequent time points.

Figure 3.

iMacs exhibit a beneficial immune response against HKMT. (a) Representative phase‐contrast microscopy images of Selene iMacs (left), NIH iMacs (middle) and MDMs (right) non‐stimulated or post‐HKMT stimulation using a Leica TCS SP5 confocal microscope (scale bars = 100 μm top and 200 μm bottom) (n = 2). (b) Flow cytometric quantification of the percentage of phagocytic macrophages at different time points post‐HKMT stimulation (n = 8–13). (c) ELISA quantification of IL‐6 and (d) TNF secretion 24 h post HKMT stimulation compared with non‐stimulated cells as controls (n = 4–20). (e) Cytokine bead assay analysis of additional pro‐ and anti‐inflammatory cytokines 24 h post HKMT stimulation (n = 3–6). (f) Flow cytometric quantification of ROS production upon HKMT stimulation. Statistical significance is indicated using a two‐tailored t‐test (n = 9–13). (g) RT‐qPCR analysis of Bax expression, (h) Cathepsin B expression and (i) LC3B expression in iMacs and MDMs post‐HKMT stimulation, presented as fold change relative to non‐infected cells as control (n = 3–6). All data are presented as mean ± SD. All statistical significance was determined using a Mann–Whitney U test (* indicates P < 0.0332, ** indicates P < 0.0021, *** indicates P < 0.0002 and **** indicates P < 0.0001). n.s., non‐stimulated control cells.

As a next step, to further determine the downstream effects of HKMT stimulation, cytokine secretion was assessed. ELISA measurements of IL‐6 secretion at 24 h post stimulation revealed elevated basal levels in both non‐stimulated iMacs and MDMs, with a highly significant increase (P < 0.0001) following HKMT stimulation reaching an average of 1206 pg/mL (± 250.9 pg/mL) for iMacs, which was markedly higher (P = 0.08) than the moderate, yet significant increase (P < 0.0332) observed in MDMs (1021 pg/mL ± 215.1 pg/mL) (Figure 3c). TNF secretion showed a similar trend, with low basal expression in non‐stimulated cells and a significant increase post‐stimulation. Similar to IL‐6 secretion, the increase of TNF was more pronounced in iMacs compared to MDMs (P < 0.0021) with a significant increase (P < 0.0001) to an average of 2275 pg/mL (± 1181 pg/mL) after HKMT‐stimulation, in comparison with MDMs with an average of 797.5 pg/mL (± 497 pg/mL), which did not reach statistical significance (P = 0.09) compared to the non‐stimulated control (Figure 3d). To gain a more comprehensive understanding of the macrophage immune response, a cytokine bead assay was conducted to assess additional pro‐ and anti‐inflammatory cytokines (Figure 3e). Consistent with the ELISA findings, iMacs exhibited higher levels of TNF, IL‐6 and other pro‐inflammatory cytokines, including IL‐1β, IL‐12 and IL‐23, at 24 h post infection than MDMs. Notably, only iMacs displayed significantly increased levels of IL‐10, an anti‐inflammatory cytokine that plays a crucial role in preventing excessive immune activation and maintaining immune homeostasis. ⁴⁰ This suggests that iMacs may facilitate a more rapid resolution of inflammation than MDMs, which exhibited a delayed and prolonged activation state. To further elucidate the response dynamics, the production of ROS was examined, a critical anti‐microbial mechanism enabling macrophages to eliminate pathogens through oxidative damage. ⁴⁵ To be specific, we used DHR 123, which reacts mainly with H₂O₂ (hydrogen peroxide) in the presence of peroxidases. Following HKMT stimulation, iMacs exhibited a significant increase in ROS production, averaging 1.8‐fold, compared with MDMs that demonstrated a reduction in ROS levels to an average of 0.88‐fold. This observation suggests that iMacs possess a greater capacity to induce ROS production, a crucial defence mechanism to fight mycobacteria (Figure 3f; Supplementary figure 2b).

Subsequently, quantitative PCR (qPCR) analysis of Bax expression was performed to evaluate proteins associated with apoptosis induction following HKMT stimulation, revealing a response pattern consistent with that observed during BCG infection. Bax expression was significantly increased in iMacs after 30 min and 1 h post stimulation, reaching a 2.4‐fold increase after 2 h, whereas MDMs exhibited a time‐dependent decrease in Bax expression (Figure 3g). A similar trend was observed in inflammasome activation, as assessed by Cathepsin B expression (Figure 3h). In iMacs, Cathepsin B expression was rapidly upregulated within 5 min of HKMT stimulation, reaching a peak 1.8‐fold increase and maintaining elevated levels over 60 min, before gradually declining. In contrast, MDMs exhibited a progressive decrease in Cathepsin B expression over the same time period. This differential response extended to the induction of autophagy‐related genes upon HKMT stimulation, as measured by LC3B expression (Figure 3i). Both iMacs and MDMs exhibited LC3B expression at 5 min post stimulation. However, whereas iMacs sustained higher expression levels over time, MDMs showed only a modest initial elevation in LC3B expression, with minimal changes throughout the observed period.

Discussion

Tuberculosis, the currently deadliest infectious disease that is caused by Mycobacterium tuberculosis (Mtb), remains a major global health threat. Current treatment and prevention strategies are often limited by adverse side effects or inefficacy, highlighting the urgent need for alternative therapeutic strategies. Given their critical role as both primary host cells and key immune regulators in TB infection, macrophages represent a promising target for investigation. ⁴⁶ This study aimed to compare immune responses between iMacs and MDMs upon mycobacterial exposure, using BCG and HKMT as pathogens.

iMacs and MDMs showed comparable differentiation efficiency, maintaining a mature macrophage phenotype (CD45⁺CD11b⁺CD14⁺CD163⁺), consistent with previous reports, demonstrating the phenotypic and functional resemblance of iMacs to MDMs ⁴⁷ and monocytic cell lines. ²⁵ iMacs further demonstrated stable production over multiple harvests, supporting their potential as a sustainable macrophage source for infection studies, as seen in previous studies. ⁴⁸ , ⁴⁹ In addition, iMacs demonstrated faster mycobacterial uptake and enhanced recruitment of vATPase and LAMP‐1, suggesting accelerated phagosome acidification and maturation. Efficient phagosome acidification is crucial for mycobacterial clearance, ⁵⁰ supporting the hypothesis that iMacs possess an innate advantage in pathogen processing. ⁴⁷ , ⁴⁹ Conversely, MDMs showed delayed phagosomal maturation and acidification, consistent with the known immune evasion strategies of Mtb. ⁵¹ Both macrophage subtypes displayed early upregulation of pro‐inflammatory cytokines upon infection, with iMacs exhibiting an overall stronger IL‐6 and TNF response. In humans, a deficiency in TNF confers a high susceptibility to TB, because of its key role in immune defence. ¹⁵ TNF is essential for activating macrophages and supporting granuloma formation, which helps controlling an Mtb infection. Individuals with genetic TNF deficiencies or those receiving anti‐TNF therapy for conditions such as rheumatoid arthritis are at an increased risk of TB reactivation and widespread infection. Interestingly, in comparison with the IL‐6 secretion, TNF levels did not further increase over time, rather remained stable in both macrophage types. While elevated TNF levels are beneficial during the acute phase of Mtb infection, excessive and prolonged TNF can contribute to increased tissue damage and exacerbate TB pathology. Therefore, the regulated cytokine responses observed in both macrophage types suggests a physiologically balanced and regulated cytokine response that prevents excessive inflammation. Such controlled TNF dynamics are advantageous, as they support macrophage activation and granuloma formation while minimising the risk of immune‐mediated tissue damage. Together, these findings suggest that both iMacs and MDMs achieve an effective equilibrium between pro‐inflammatory activity and immune regulation, with iMacs exhibiting a stronger initial responsiveness that could enhance early mycobacterial containment, which may be advantageous for potential anti‐TB therapeutic applications. ³⁶ Cytokine profiling further revealed that iMacs show a trend to higher levels of IL‐1β, IL‐12 and IL‐23, which are also critical for bacterial clearance, ³⁷ , ³⁸ , ³⁹ while simultaneously increasing anti‐inflammatory cytokines such as IL‐10 and IL‐33, indicating a rapid return to immune homeostasis. ⁴⁰ , ⁴¹ The ability of iMacs to transition efficiently between activation and resolution phases suggests a distinct regulatory mechanism that may be crucial for mitigating excessive immune activation and preventing tissue damage. These findings are consistent with transcriptome analyses from previous infection studies with Staphylococcus aureus ¹⁹ and Pseudomonas aeruginosa, ²⁸ demonstrating that iMacs balance inflammatory responses more effectively than MDMs, and thus, further enhance bacterial clearance while preventing chronic inflammation. ⁵² This regulatory mechanism could be particularly advantageous for TB research and future therapeutic applications, as uncontrolled inflammation may otherwise exacerbate disease progression. This makes iMacs a promising model for studying host–pathogen interactions and macrophage‐mediated responses in TB. Macrophage activation upon TB infection involves key signalling pathways, including p38 MAPK and SAPK/JNK, which regulate cytokine production, autophagy and apoptosis, orchestrating a balance between pathogen clearance and host cell survival. ⁴⁴ Indeed, we observed that both macrophage types exhibited activation of these pathways with iMacs showing overall stronger phosphorylation responses, suggesting enhanced immune regulation. Interestingly, only iMacs showed increased Bax expression, indicating higher rates of apoptosis‐related proteins upon infection consistent with earlier reports that link apoptosis to efficient mycobacterial control in these cells. ²⁵ Autophagy, another key antimicrobial process, remains a subject of debate in TB pathogenesis. ⁶ While some studies suggest that only virulent Mtb suppresses autophagy through specific virulence factors, others report that even attenuated or heat‐killed strains can modulate autophagic flux. ⁵³ In our study, iMacs displayed a more sustained activation of autophagy‐regulating proteins, including Beclin‐1 and LC3B, suggesting a greater capacity for intracellular bacterial degradation. This aligns with prior observations of persistent LC3B⁺ autophagosome formation in iMacs during mycobacterial infection. ²⁴ In contrast, MDMs exhibited weaker and transient responses, followed by a decline, indicating autophagic arrest. Together, these findings suggest that iMacs maintain a more active autophagic programme, which could facilitate bacterial clearance and provide a useful platform for identifying autophagy‐targeting TB therapeutics. The rapid and robust immune response of iMacs to mycobacterial infection likely stems from their embryonic‐like phenotype, which resembles tissue‐resident alveolar macrophages, which are key players in pulmonary immunity. Their primitive, foetal‐like state enhances pathogen recognition, inflammatory signalling and transcriptional activation because of a distinct epigenetic landscape that maintains immune genes in a more accessible state, rendering iMacs particularly responsive to mycobacterial infections. In the context of cell‐based therapies, iMacs offer key advantages over MDMs, including homogeneity and reproducibility. Unlike donor‐derived MDMs, which exhibit variability, iMacs provide an unlimited and standardised source of macrophages, as confirmed by the use of two distinct iPSC clones in the present study. For therapeutic applications, iMac production can be upscaled using bioreactors and optimised protocols to generate large quantities of macrophages for pulmonary or systemic administration. ⁴⁹ , ⁵⁴ , ⁵⁵ This enables off‐the‐shelf therapies, overcoming donor‐related logistical and ethical challenges. Unlike MDMs, which require fresh isolation and differentiation, iMacs can be cryopreserved and expanded, ensuring long‐term feasibility for research and clinical applications. ⁵⁶

In summary, the enhanced immune responses in iMacs position them as a promising model for host–pathogen interactions in TB research, where strong macrophage activity is crucial for infection control. Their distinct autophagy‐associated signalling, cytokine secretion and apoptosis‐regulating protein profiles highlight their potential as an immunocompetent model for mycobacterial infections. Given their enhanced responsiveness, iMacs also serve as a valuable in vitro platform for screening TB therapeutics targeting macrophage pathways. Future studies should investigate their transcriptional and epigenetic profile to elucidate the underlying immune mechanisms, while in vivo research could further assess their therapeutic potential in TB models. Moreover, future work should include analyses of the trained immunity potential of iMacs in comparison with primary MDMs, to determine whether iMacs are capable of exhibiting memory‐like responses upon repeated pathogen exposure. While direct studies on intrapulmonary macrophage transplantation for TB are limited, research in other lung diseases provides valuable insights. Studies have demonstrated that intrapulmonary transfer of iMacs can effectively and sustainably treat Hereditary Pulmonary Alveolar Proteinosis (herPAP), a rare lung disease, characterised by the accumulation of surfactant because of dysfunctional alveolar macrophages. In a humanised murine model of herPAP, transfered iMacs exhibited long‐term pulmonary engraftment and differentiated into functional alveolar macrophages, leading to significant disease amelioration. ²⁷ Furthermore, research has shown that alveolar macrophages in a donor lung can persist in transplanted human tissue for extended periods, up to three and a half years post‐transplantation. ⁵⁷ These macrophages remain functional and exhibit minimal replacement by recipient‐derived cells, indicating their long‐term residency in lung tissue. Given that alveolar macrophages play also a critical role in host defence against Mtb, this approach may also hold promise for potential TB therapy. iMacs could potentially replace or enhance endogenous macrophages, improving bacterial clearance while mitigating excessive inflammation. These findings provide a strong rationale for further investigation into iMac transplantation as a novel therapeutic strategy for TB and other chronic pulmonary infections. The present study, however, has several limitations that should be considered when interpreting the findings. First, iMacs and MDMs were derived from genetically distinct donors, and inter‐individual genetic variation may have influenced the observed immune differences. Although iPSCs provide a renewable and standardised macrophage source, future studies using donor‐matched iMacs and MDMs would enable more direct genotype‐controlled comparisons. Second, our analyses were limited to Mycobacterium bovis BCG and heat‐killed Mtb as surrogates for infection. While safe and widely used, these models cannot fully reproduce the complexity of virulent Mtb infection. Validation of our findings using live pathogenic strains will therefore be essential to confirm the physiological relevance of the observed mechanisms. Third, we did not directly compare the anti‐mycobacterial activities of iMacs with those of MDMs. Future work should include colony‐forming unit (CFU) assays to evaluate their capacity to control and eradicate intracellular bacteria. Such functional assays will help determine whether the enhanced immune signalling of iMacs translates into improved bacterial clearance, providing critical insights for the potential development of iPSC‐derived macrophage therapies against tuberculosis.

Methods

Production of iPSC‐derived macrophages (iMacs)

iMacs were derived from iPSCs using a feeder‐free protocol. Two established iPSC lines were utilised and compared: The first iPSC line was developed in the Hannover Medical School using CD34⁺ cells from peripheral blood of a healthy female donor ⁵⁴ (hCD34iPSC1151/MHHi015‐B: https://hpscreg.eu/cell‐line/MHHi015‐B) and will further be labelled as ‘Selene’. The second iPSC line is established from a healthy male donor ⁵⁸ (LiPSC‐GR1.1) and will further be labelled as ‘NIH’. Generation of the GMP line LiPSC‐GR1.1 was supported by the NIH Common Fund Regenerative Medicine Program and reported in Stem Cell Reports. The NIH Common Fund and the National Center for Advancing Translational Sciences (NCATS) are joint stewards of the LiPSC‐GR1.1 resource (https://hpscreg.eu/cell‐line/RUCDRi002‐A). Detailed protocols for both iPSC cultivation and the subsequent differentiation into macrophages have been previously published. ⁴⁸ After 5–7 days, the fully‐differentiated iMacs were harvested by gently detaching them using phosphate‐buffered saline (PBS, CHEMSOLUTE^®/Th.Geyer, Cat. No. 8435.0100, Höxter, Germany). For all experiments, only cells collected from the third harvest onwards were utilised. Unless otherwise specified, iMacs were cultivated in RPMI 1640 medium (Thermo Fisher Scientific, Cat. No. 21875‐034, Waltham, MA, USA) supplemented with 10% FBS Superior (Sigma‐Aldrich, Cat. No. S0615, Steinheim, Germany), 1% Penicillin–Streptomycin (Gibco, Cat. No. 15140‐122) and 50 ng/mL huM‐CSF (Peprotech, Cat. No. 300‐25, Hamburg, Germany). Cultures were maintained in an incubator at 37°C with 5% CO₂.

Production of monocyte‐derived macrophages from peripheral blood (MDMs)

The use of human donor blood was approved by the MHH ethical committee (approval number: 1427‐2012). Buffy coat blood from anonymous healthy donors was commercially obtained from ‘Deutsches Rotes Kreuz, DRK’. All donors provided informed consent for the scientific use of blood products in compliance with DRK and institutional ethical guidelines. Monocytes were isolated and differentiated into macrophages following an established protocol. ¹⁹ In deviation from the cited protocol, concentrations of huIL‐3 and huM‐CSF were raised to 25 ng/mL and 50 ng/mL, respectively, for the first 7 days of differentiation. Further differentiation for 3 days was performed using RPMI 1640 medium supplemented with 10% FBS Superior, 1% Penicillin–Streptomycin and 50 ng/mL huM‐CSF. Unless stated otherwise, the specified medium was utilised for the cultivation of MDMs, with the cells maintained in an incubator at 37°C under 5% CO₂ conditions. Before utilisation, the MDMs were washed with PBS (CHEMSOLUTE^®/Th.Geyer, Cat. No. 21875‐034) and treated with 1:3‐diluted (1.33 g/mL) Trypsin (Gibco, Cat. No. 27250‐018) for 5–7 min (min) at 37°C. Afterwards, cell culture medium was added and cells were gently detached using a cell scraper (TPP, Cat. No. 99002, Trasadingen, Switzerland).

Phagocytosis assay using pHrodo E. coli bioparticles

MDMs and iMacs were seeded in 12‐well tissue culture plates (TPP, Cat. No. 92412), at a density of 2 × 10⁵ cells per well. After a settling period of at least 24 h (h), the medium was replaced with 2 mL of RPMI 1640 medium supplemented with 10% FBS Superior, 2 mM L ‐Glutamine, 1% Penicillin–Streptomycin, 2% HEPES (Sigma‐Aldrich, Cat. No. H3537) and 50 ng/mL huM‐CSF. To assess phagocytic activity, 20 μL of pHrodo™ Deep Red E. coli BioParticles™ Conjugate for Phagocytosis (Cat. No. P35360, Invitrogen, Carlsbad, CA, USA) were added to the cultures and incubated for 2 h at 37°C, 5% CO₂. Subsequently, the cells were washed with 1 mL of Dulbecco's phosphate‐buffered saline (DPBS) (Gibco, Cat. No. 14190094, Paisley, GB). The amount of actively phagocytosed BioParticles was assessed using a CytoFLEX S flow cytometer (Beckman Coulter, Brea, California, USA).

Flow cytometry

For the analysis of surface marker expression and the phagocytosis assays, a CytoFLEX S flow cytometer (Beckman Coulter, Brea, CA, USA) was used. The following antibodies were used in this study: hCD45‐eFluor450 (Cat. No. 48‐0459‐41), hCD11b‐PE‐Cy7 (Cat. No. 25‐0118‐42), hCD14‐PE (Cat. No. 12‐0149‐41), hCD163‐APC (Cat. No. 25‐0118‐42), hCD86‐PE (Cat. No. 12‐0869‐41), hHLA‐DR‐APC (Cat. No. 307609), hCD206‐BV421 (Cat. No. 321126), hTLR2/CD282‐PE (Cat. No. 12–9922‐41) and hTLR4/CD284‐PE‐Cy7 (Cat. No. 25‐9917‐42) (all from eBioscience, only HLA‐DR from BioLegend, both San Diego, CA, USA). Prior to 20 min antibody staining at RT according to the manufacturer's instructions, the cells were blocked for 5 min at RT using human Fc‐Block TruStain FcX™ (Biolegend, Cat. No. 422302). For all markers, 50 000 events were acquired and subsequent analysis was performed using FlowJo10.

BCG culture and infection

The culturing of Mycobacterium bovis strain Bacillus Calmette–Guérin (BCG, Danish strain 1331) was carried out according to the previously published protocol. ⁵⁹ , ⁶⁰ For the majority of experiments, green fluorescent protein (GFP)‐expressing BCG (GFP‐BCG) was used. ⁶¹ , ⁶² The GFP‐BCG strain was constructed by transforming BCG with the dual reporter plasmid pSMT3L × EGFP. ⁶² For infection of iMacs and MDMs, BCG was freshly thawed and kept in culture on a shaker at 120 rpm at 37°C in Erlenmeyer flasks including 10 mL Middlebrook 7H9 Broth supplemented with glycerol (BD Biosciences, Heidelberg, Germany) and 50 g/mL Hygromycin B to maintain reporter gene expression. For infection experiments, BCG was collected by centrifugation for 10 min at 880 × g with subsequent resuspension in HEPES/saline buffer (H/S). To achieve proper separation of live and dead bacteria, BCG was vortexed for 5 min at highest speed, bath‐sonication was performed for 5 min at 4°C and the bacteria were passed several times through a syringe with a needle of 0.8 mm in diameter. The bacterial density was calculated by measuring the optical density (OD) and a standard curve. In addition to the OD measurement, CFU counts were determined for representative batches to verify the OD‐CFU correlation and ensure consistency of the multiplicity of infection (MOI) between experiments. For the infection of cells, iMacs and MDMs were kept in Minimal Essential Medium (MEM, Gibco, Paisley, GB) supplemented with 10 mM HEPES and were infected with an MOI of 10:1 BCG for the depicted time points. To enhance bacterial–host cell interactions and establish reproducible infection conditions, the bacteria were centrifuged onto the cells at 55 × g for 8 min.

For the LEGENDplex assay, RFP‐expressing Mycobacterium bovis BCG (Pasteur strain) was used. BCG was cultured under the same conditions as described above. To ensure proper bacterial dispersion, BCG was vortexed at maximum speed for 5 min using glass beads. For infection experiments, 5 × 10⁴ iMacs or MDMs were seeded in a 48‐well tissue culture plate. The following day, cells were infected with BCG at an MOI of 1:1. At 24 h post infection, the supernatant was collected, centrifuged at 300 × g for 5 min to remove residual cells, and stored at −20°C until the LEGENDplex assay was performed.

Preparation of serum‐opsonised heat‐killed Mycobacterium tuberculosis (HKMT)

For the experiments utilising HKMT, the strain H37Ra was used (Cat. No: tlrl‐hkmt‐1, InvivoGen, San Diego, CA, USA). HKMT were centrifuged at 4000 × g for 5 min at 4°C, and the supernatant was discarded. For serum opsonisation, 500 μL of DPBS and 500 μL of human AB serum (Sigma‐Aldrich, Cat. No. H4522, Steinheim, Germany) were added to the HKMT suspension. After an initial incubation for 10 min at 37°C, the tubes containing the HKMT were inverted and incubated for an additional 10 min at 37°C. The bacteria were then centrifuged at 4000 × g for 5 min at 4°C, the supernatant was discarded and the bacteria pellet was resuspended in 1 mL of DPBS. For proper washing, this process was repeated twice. Afterwards, the HKMT were resuspended in 3 mL DPBS to achieve a final concentration of 3.3 mg/mL. The prepared HKMT vials were stored at −80°C in 200 μL aliquots until further use.

Immune fluorescence microscopy

Immunofluorescence staining was performed following the protocol previously described. ⁵⁹ In short, 1.2 × 10⁵ iMacs or MDMs were seeded on round coverslips (12 mm in diameter) and infected with an MOI of 10:1 BCG. The cells were fixed in 1% paraformaldehyde (PFA; Sigma‐Aldrich, Steinheim, Germany) diluted in PBS for 15 min at RT, washed once with HEPES/Saline (H/S, 200 mM HEPES, 1.32 M NaCl, 10 mM CaCl₂, 7 mM MgCl₂, 8 mM MgSO₄, 54 mM KCl, pH 7.3) solution before being permeabilised with 0.1% Triton X‐100 (Sigma‐Aldrich) diluted in H/S for 10 min at RT. Next, the cells were washed with H/S with 0.05% Tween‐20 (Sigma‐Aldrich, Steinheim, Germany), followed by a blocking step for 15 min at RT using H/S solution supplemented with 0.05% Tween‐20 and then incubated for 45 min with the respective antibodies diluted in 5% FCS/H/S (FCS; Thermo Fisher Scientific). The following primary antibodies were used: Lamp1 (Cat. No. SC5570) and vATPase (Cat. No. SC28801), both purchased from Santa Cruz, CA, USA. As a second antibody, an IgG rabbit antibody was used. After an additional three times of washing with H/S with 0.05% Tween‐20, followed by a single wash with H/S, the infected cells and the respective non‐infected controls were mounted on glass slides using Mowiol (Kuraray Specialities Europe GmbH, Frankfurt, Germany). All images were taken with an inverted Leica TCS SP5 microscope using the 100x (oil immersion) objective (Leica Microsystems, Wetzlar, Germany) and analysed with the Leica LCS software. The Pearson correlation coefficient was calculated to assess the degree of colocalisation between fluorescent signals using the JaCoP (Just Another Colocalisation Plugin) in ImageJ. Briefly, regions of interest (ROIs) were defined on the images, and background subtraction was applied prior to analysis.

Western blot analysis

For the determination of autophagy‐ and apoptosis‐related markers after BCG infection, Western blot analysis was performed as previously described. ⁶⁰ In short, 2 × 10⁵ iMacs or MDMs were seeded in a 12‐well adherent tissue culture plate overnight (o.n.). The cells were then infected with BCG using an MOI of 10:1. After 0 min, 5 min, 30 min, 60 min and 120 min, cells were washed twice and proteins were extracted using lysis buffer supplemented with 0.1% SDS and 10 mg/mL Aprotinin/Leupeptin. 20–30 μg of the samples were loaded and separated by 10–15% NuPage Bis‐Tris gradient gels (Life Technologies) and transferred to nitrocellulose membranes (GE Lifesciences). After an 1 h blocking step at RT using Pierce Starting Block Solution (Thermo Fisher Scientific), the membrane was incubated with the following primary antibodies at 4°C o.n.: LC3B (Sigma Aldrich L7543, rabbit IgG), Beclin‐1 (Cell Signaling 3495, rabbit polyclonal), phospho‐p38 MAPK (Thr180/Tyr182) (Cell Signaling 9211, rabbit polyclonal), phospho‐SAPK/JNK (Thr183/Tyr185) (Cell Signaling 9251, rabbit polyclonal) and Bax (Cell Signaling 2772, rabbit polyclonal). After thoroughly washing for at least six times in TBS/Tween (Tris‐buffered saline supplemented with 0.1% Tween 20), an anti‐rabbit IgG Alkaline‐phosphatase‐(AP)‐conjugated secondary antibody was used (Abcam, Cat. No. ab97048, Cambridge, UK) for 1 h at RT using TBS/T. Chemiluminescence signal was determined using CDP Star substrate (PerkinElmer, Cat. No. NEL616001KT, Rodgau, Germany). Subsequent densitometric analysis was performed on scanned images using ImageJ^® software, and values were normalised to actin expression and non‐infected control cells of each experiment.

Phagocytosis assay using stained HKMT

5 × 10⁵ MDMs or iMacs were seeded into a 12‐well format tissue culture plate (TPP, Cat. No. Z707775‐126EA) using 1.5 mL of respective cell culture medium and were allowed to adhere o.n. HKMT was stained with BactoView™ Live Fluorescent Bacterial Stains in ‘live red’ (Biotium, Cat. No. 40101‐T) according to the manufacturer's instructions. Subsequently, 50 μL of the stained HKMT was added to each well and incubated for the designated time periods. Following incubation, cells were washed, and the extent of HKMT uptake was quantified using the CytoFLEX S flow cytometer (Beckman Coulter).

Cytokine secretion

To assess cytokine production, 1 × 10⁵ MDMs or iMacs were seeded in a 48‐well tissue culture plate (TPP, Cat. No. 92448) using 400 μL of the respective cell culture medium as described in Generation and characterization of iPSC‐ and blood monocyte‐derived macrophages and iMacs demonstrate faster phagocytosis and increased pro‐inflammatory cytokine secretion upon BCG infection. Each condition was prepared in duplicates. For infection with HKMT, 20 μL were added to each well, followed by incubation for 24 h. Supernatants were collected and centrifuged for 3 min at 1500 rpm using the Eppendorf centrifuge 5425 R. The supernatants were stored at −20°C until further analysis. ELISAs were performed using the Human IL‐6 DuoSet ELISA (R&D Systems, Cat. No. DY206) and the TNF‐alpha DuoSet ELISA (R&D Systems, Cat. No. DY210), following the manufacturer's instructions.

For infection with BCG, 1.2 × 10⁵ iMacs or MDMs were seeded in a 24‐well tissue culture plate o.n. The cells were infected with BCG using an MOI of 10:1 or 50:1 BCG in the respective cell culture medium. After 6 h and 24 h, supernatants were collected, centrifuged at 1000 rpm for 5 min and analysed using the human IL‐6 and TNF Quantikine ELISA Kit (R&D Systems, #D6050 and #DTA00D, respectively) according to the manufacturer's instructions. In addition, for BCG and HKMT, two and three biological replicates were analysed for cytokine/chemokine secretion using a LEGENDplex™ Human Inflammation Panel 1 Standard Assay (BioLegend, Cat. No. 740811).

Gene expression analysis using RT‐qPCR

5 × 10⁵ MDMs or iMacs were seeded in a 12‐well tissue culture plate (TPP, Cat. No. Z707775‐126EA). Each condition was prepared in duplicates. After the cells settled o.n., the infection was performed by adding 50 μL of HKMT per well. RNA was extracted using the RNeasy Micro Kit (Qiagen, Cat. No. 74004, Hilden, Germany), and an on‐column DNase digest was performed according to the manufacturer's instructions. Subsequently, RNA was transcribed into cDNA by adding 1 μL Oligo(dT)18 primer (Thermo Fisher Scientific, Cat. No. SO132), 2 μL 10 mM dNTP mix (Thermo Fisher Scientific, Cat. No. R0181), 1 μL RiboLock RNAse Inhibitor (Thermo Fisher Scientific, Cat. No. EO0381), 1 μL RevertAid Reverse Transcriptase and 4 μL 5× reaction Buffer for RT (Thermo Fisher Scientific, Cat. No. EP0442) per 1 μL RNA. Incubation was performed at 42°C for 60 min followed by 5 min of incubation at 70°C. Each sample was mixed with 7.5 μL SYBR Green PCR Master Mix (Applied Biosystems, Cat. No. 4309155), H₂O and 1 μL of the respective primer to a total volume of 15 μL. Primers were purchased from QuantiTect, and the following primers were used: Hs_MAP1LC3B_1_SG (Cat. No. QT00055069), Hs_CTSB_1_SG (Cat. No. QT00088641) and Hs_BAX_1_SG (Cat. No. QT00031192). The 7500 Fast Real‐Time PCR System (Applied Biosystems, Darmstadt, Germany) was used to measure all samples.

Reactive oxygen species (ROS) production

A total of 1 × 10⁶ MDMs or iMacs were seeded in a 6‐well tissue culture plate (TPP, Cat. No. Z707767‐72EA) and allowed to adhere o.n. For infection, the culture medium was replaced, and 50 μL of HKMT was added per well. At 24 h post infection, cells were washed once with PBS, harvested by centrifugation (5 min at 1500 rpm), and resuspended in 500 μL HBSS⁺⁺. Cells were then incubated for 5 min at 37°C with shaking (300 rpm). The cells were divided into three experimental conditions: (1) unstained, (2) stained unstimulated and (3) stained stimulated (as positive control). To achieve this, the cells were distributed into three microcentrifuge tubes and adjusted to 500 μL HBSS⁺⁺ per condition. In the stimulated condition (3), 10 μL of PMA (20 μg/mL) was added, followed by incubation for 5 min at 37°C. Subsequently, 25 μL of dihydrorhodamine 123 (DHR 123) (10 μg/mL) was added to Conditions 2 and 3, and samples were incubated for an additional 15 min at 37°C. Following incubation, samples were placed on ice, and ROS production was quantified using the FITC channel on a CytoFLEX S flow cytometer within 30 min.

Microscopy and live‐cell imaging

A total of 5 × 10⁵ iMacs were seeded into a 12‐well tissue culture plate in 1.5 mL of the respective cell culture medium. Following an incubation period of at least 24 h to allow for cell adherence, the medium was replaced with 1 mL of RPMI 1640 medium supplemented with 10% FBS, 2 mM L‐Glutamine, 1% P/S, 2% HEPES and 50 ng/mL huM‐CSF. To evaluate migratory activity, 50 μL of HKMT was added to the culture, and cells were incubated at 37°C in 5% CO₂. Bright‐field microscopy images were acquired using a ZEISS Axio Vert.A1 microscope equipped with 10× and 20× objectives (Carl Zeiss, Oberkochen, Germany). For the assessment of active migration over time, live‐cell imaging was performed under the same seeding conditions. Time‐lapse videos were taken using a ZEISS Axio Observer Z1 microscope with a 10× objective and analysed using the ImageJ software.

Macrophage polarisation

iMacs were seeded at a density of 2.5 × 10⁵ cells per well in 24‐well tissue culture plate and terminally differentiated for 5 days in RPMI 1640 medium supplemented with 50 ng/mL huM‐CSF. For polarisation, cells were washed once with PBS and starved overnight in cytokine‐free XVIVO medium. Subsequently, iMacs were stimulated for 24 h with either 10 ng/mL recombinant human IL‐4 (Peprotech) to induce M2 polarisation or 25 ng/mL recombinant human IFN‐γ (Peprotech) to induce M1 polarisation. Supernatants from IFN‐γ–stimulated cells were collected for IL‐6 quantification using a human IL‐6 ELISA kit (R&D Systems), following the manufacturer's instructions. After stimulation, cells were washed once with PBS and stained with anti‐HLA‐DR (M1 marker) or anti‐CD206 (M2 marker) antibodies for flow‐cytometric analysis on a Cytoflex S (Beckman Coulter). For LPS stimulation, iMacs were seeded and differentiated as described above and then stimulated with 100 ng/mL LPS (Sigma) in RPMI medium containing M‐CSF for 24 h without prior starvation. The supernatant was collected after 24 h and analysed using the same human IL‐6 ELISA kit (R&D Systems).

Cell viability

For viability assessment, 2.5 × 10⁵ iMacs were seeded and differentiated for 5 days under the same conditions. Cells were washed once with PBS, harvested and stained with Zombie Aqua (1:1000 dilution in PBS; BioLegend, Cat. No. 423101) for 30 min at in the dark. Samples were immediately analysed using a Cytoflex S flow cytometer (Beckman Coulter).

Statistical analysis

Statistical analyses were conducted using Prism version 9.4.1 (Graph Pad Software). Error bars throughout the paper is denoted in 95% confidence intervals of the mean. * indicates P < 0.0332, ** indicates P < 0.0021, *** indicates P < 0.0002 and **** indicates P < 0.0001.

Author contributions

Conceptualisation, DP, HS, HG and NL; methodology, DP, HS, HG, AR, BC, TG, ES, JD, AHHN, ES, TB, A‐LN, JB, HT and AP; investigation, DP, HS, HG and NL; resources, HG, EG, NL and UK; writing—original draft preparation, DP, HS and NL; writing—review and editing, DP, HS, HG, UK and NL; visualisation, DP, HS and HG; supervision, HG, NL and UK.; project administration, HG, GH and NL; funding acquisition, NL, HG, EG and UK. All authors have read and agreed to the published version of the manuscript.

Conflict of interest

The authors declare no conflict of interest.

Supporting information

Supplementary information 1

Supplementary information 2

Supplementary figure 1

Supplementary figure 2

Supplementary table 1

Supplementary video 1

Data availability statement

The data that support the findings of this study are available from the corresponding author upon reasonable request.