Abstract

   Staphylococcus aureus (S. aureus) is a major zoonotic pathogen, with
   mammary gland infections contributing to mastitis, a condition that
   poses significant health risks to lactating women and adversely affects
   the dairy industry. Therefore, understanding the immune mechanisms
   underlying mammary infections caused by S. aureus is essential for
   developing targeted therapeutic strategies against mastitis. This study
   identified hydroxycarboxylic acid receptor 2 (HCAR2) as a potential
   regulator of S. aureus infection in mammary glands. It is demonstrated
   that HCAR2 deficiency exacerbates the inflammatory response and
   disrupts the blood‐milk barrier in the mammary gland during S. aureus
   infection, with NLRP3 inflammasome‐mediated pyroptosis playing a
   central role. Activation of HCAR2, on the other hand, suppressed CMPK2
   expression, thereby mitigating mitochondrial damage and pyroptosis in
   mouse mammary epithelial cells (mMECs) induced by S. aureus.
   Additionally, mitochondrial DNA (mtDNA) released from S.
   aureus‐infected mMECs activates the cGAS/STING signaling pathway in
   macrophages, impairing their bactericidal activity. In conclusion, this
   study highlights the critical role of HCAR2 in S. aureus infection of
   the mammary gland and provides a theoretical basis for identifying
   potential therapeutic targets for such infections.

   Keywords: CMPK2, HCAR2, mastitis, pyroptosis, S. aureus
     __________________________________________________________________

   In the mammary gland infected with S. aureus, the activation of
   hydroxycarboxylic acid receptor 2 (HCAR2) in mammary epithelial cells
   (mMECs) leads to a reduction in CMPK2 expression, which in turn
   mitigates cell pyroptosis. The attenuation of pyroptosis in mMECs
   reduces the release of mitochondrial DNA (mtDNA) into the extracellular
   space, thereby alleviating the suppression of macrophage activity
   induced by S. aureus via the cGAS/STING‐dependent pathway.

   graphic file with name ADVS-12-2411947-g004.jpg

1. Introduction

   Staphylococcus aureus (S. aureus) is a major zoonotic pathogen
   responsible for extensive clinical infections in both humans and
   livestock, posing a significant threat to public health due to its
   infectious nature and pathogenic potential.^[ [40]^1 , [41]^2 ^]
   Mammary gland infections caused by S. aureus are a primary contributor
   to mastitis, often progressing to purulent mastitis, which severely
   disrupts the structural integrity of the mammary gland. If not promptly
   and effectively treated, mastitis can lead to systemic infectious
   complications, endangering the health of lactating women and negatively
   impacting the dairy industry.^[ [42]^3 , [43]^4 , [44]^5 ^] Currently,
   S. aureus exhibits resistance to many commonly used antibiotics, and
   even strains classified as sensitive can still cause severe infections
   in the host.^[ [45]^6 , [46]^7 ^] Therefore, understanding the immune
   mechanisms involved in S. aureus‐induced host infections and developing
   strategies to bolster the host's defense mechanisms is critical. This
   approach is essential for preventing further resistance development and
   minimizing the impact of broad‐spectrum antibiotics on beneficial
   microbiota, thereby enabling more effective targeted treatment
   strategies for managing infections.

   The innate immune system plays a pivotal role in the early detection
   and elimination of invading S. aureus, utilizing pattern recognition
   receptors (PRRs) to identify pathogen‐associated molecular patterns
   (PAMPs) and initiate appropriate defense responses within host cells.^[
   [47]^8 ^] Mammary epithelial cells (mMECs), as the functional units of
   the mammary gland, are primarily responsible for synthesizing,
   metabolizing, and maintaining the integrity of the blood‐milk barrier.
   Loss of functionality in these cells can lead to irreversible
   disruption of the blood‐milk barrier.^[ [48]^9 , [49]^10 ^] During
   mammary infection, these epithelial cells act as sentinel cells,
   initially detecting components of S. aureus and contributing to the
   host's defense mechanisms. However, S. aureus can induce various forms
   of programmed cell death (PCD) in mMECs, including pyroptosis, a highly
   inflammatory type of cell death. This mode of cell death not only
   compromises the blood‐milk barrier but also facilitates the recruitment
   of diverse immune cell populations.^[ [50]^11 , [51]^12 ^] Although
   recruited macrophages and neutrophils are essential for pathogen
   clearance and inflammatory response regulation, their excessive
   pro‐inflammatory activation is a key factor in mammary tissue damage.^[
   [52]^13 ^] Studies suggest that excessive epithelial cell death can
   trigger a dysregulated inflammatory response, impairing the
   functionality of immune cells, including macrophages.^[ [53]^14 ^] Upon
   cell death, damage‐associated molecular patterns (DAMPs) and cellular
   debris are released, which are subsequently engulfed and degraded by
   macrophages through phagocytosis. However, excessive cell death, along
   with varying modes of PCD, can directly affect the functional capacity
   and efficiency of macrophages.^[ [54]^15 , [55]^16 ^] Consequently,
   host responses during pathogen elimination may lead to detrimental
   inflammation, exacerbating clinical symptoms. A comprehensive
   understanding of the innate immune mechanisms employed during S. aureus
   mammary infections is essential for identifying novel prevention and
   treatment strategies.

   G protein‐coupled receptors (GPCRs) are integral to various
   physiological processes, including immune regulation. Endogenous
   peptides, short‐chain fatty acids, and proteases serve as natural
   ligands for GPCRs.^[ [56]^17 ^] This study identifies hydroxycarboxylic
   acid receptor 2 (HCAR2) as a key player in S. aureus infection within
   mammary tissue. HCAR2 is expressed in mMECs and macrophages, with
   endogenous compounds such as nicotinic acid (Nia) acting as ligands for
   HCAR2.^[ [57]^18 , [58]^19 ^] Additionally, research by Guo et al.
   demonstrated that HCAR2 activation can mitigate sepsis, highlighting
   its role in infectious diseases.^[ [59]^20 ^] Despite this, the precise
   function and underlying mechanisms of HCAR2 in S. aureus infection
   remain unclear. This study elucidates the role of HCAR2 in modulating
   S. aureus infection in the mammary gland and provides novel insights
   into potential therapeutic targets for S. aureus‐related diseases.
   These findings offer significant scientific value in developing
   strategies to reduce the incidence and progression of such infections.

2. Results

2.1. HCAR2 Exhibits a Significant Correlation with Mammary Infections caused
by S. aureus

   To identify the key target involved in S. aureus‐induced mammary
   infection, a mouse model of mastitis was established using S. aureus.
   Infection with S. aureus resulted in significant swelling and redness
   of the mammary glands, alveolar degeneration, and inflammatory cell
   infiltration within the alveolar cavity (Figure [60]1a,b). Furthermore,
   S. aureus infection led to a marked increase in MPO content, bacterial
   load, and pro‐inflammatory factors within the mammary gland
   (Figure [61]1c–g). Transcriptome sequencing was performed to analyze
   gene expression profiles in the mammary tissues of the NT and S. aureus
   groups. The analysis revealed a total of 2056 differentially expressed
   genes between the two groups (Figure [62]1h). Gene Ontology (GO)
   enrichment analysis indicated significant differences in several
   pathways, including innate immune response, pyroptosis, and various
   pathways related to GPCRs (Figure [63]1i). Additionally, the Kyoto
   Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis
   identified alterations in nicotinic acid metabolism. Further clustering
   of differentially expressed genes revealed a significant upregulation
   of the G protein‐coupled receptor HCAR2, activated by Nia, in the S.
   aureus‐stimulated group (Figure [64]1j,k). Gene set enrichment analysis
   (GSEA) further highlighted the upregulation of the NOD‐like receptor
   signaling pathway and alterations in nicotinic acid and nicotinamide
   metabolism in mammary tissue following S. aureus stimulation
   (Figure [65]1l,m). To validate the results of the transcriptome
   sequencing, HCAR2 expression was examined. The results confirmed that
   S. aureus stimulation significantly induced the transcription and
   translation of GSDMD and HCAR2 in mammary tissue, with
   immunohistochemistry (IHC) results consistent with the protein
   expression data (Figure [66]1n–q). These results suggest that HCAR2 is
   involved in mammary infections induced by S. aureus.

Figure 1.

   Figure 1
   [67]Open in a new tab

   HCAR2 exhibits a significant correlation with mammary infections caused
   by S. aureus. a) Images of mouse mammary glands and H&E staining; red
   arrows indicate inflammatory cell infiltration, green arrows highlight
   thickened acinar walls, and Scale bar represents 100 µm. b) Mammary
   injury score. c) MPO content in mammary tissue. d) Bacterial burden in
   mammary tissue. e–g) ELISA quantification of IL‐1β, IL‐6, and TNF‐α
   protein levels in mammary tissue. h) Transcriptome volcano plot of
   mammary gland tissues. “NoDiff” represents no significant difference in
   gene expression between groups, “Sig‐Up” indicates upregulation
   compared to the NT group, and “Sig‐Down” indicates downregulation
   compared to the NT group. i) GO enrichment analysis. j) Cluster
   analysis of differentially expressed genes. k) KEGG enrichment
   analysis. l, m) GSEA analysis for Nicotinate and Nicotinamide
   metabolism and NOD‐like receptor signaling pathway. n) mRNA expression
   of HCAR2, NLRP3, and GSDMD in mammary tissues. o) Western blot analysis
   of GSDMD and HCAR2 protein levels in mammary tissues. p, q)
   Immunohistochemistry (IHC) staining for N‐GSDMD and HCAR2 in mouse
   mammary tissue. Scale bar = 50 µm. Results are presented as mean ± SD,
   n = 6.

2.2. HCAR2 Deficiency Exacerbates the Severity of S. aureus Infection in
Mouse Mammary Glands

   To elucidate the role of HCAR2 in S. aureus infection of the mammary
   gland, experiments were conducted using HCAR2 knockout (HCAR2^−/−)
   mice. Initial confirmation of HCAR2 deletion in the mammary glands of
   these mice was performed (Figure [68]S1, Supporting Information).
   Subsequently, a mastitis model was established in both wild‐type (WT)
   and HCAR2^−/− mice by infecting them with S. aureus. No noticeable
   differences were observed between the mammary glands of WT and
   HCAR2^−/− mice under baseline conditions. However, after infection,
   HCAR2^−/− mice exhibited significantly more pronounced redness and
   swelling in the mammary glands compared to the NT group, along with an
   increased number of necrotic foci within the acini. This resulted in a
   higher tissue damage score for HCAR2^−/− mice than for WT mice (Figure
   [69]2a,b). Furthermore, HCAR2^−/− mice showed a higher bacterial load
   in the mammary glands, although no significant difference in MPO
   content was observed between HCAR2^−/− and WT mice (Figure [70]2c,d).
   Elevated levels of IL‐1β and IL‐6 were also detected in the mammary
   glands of HCAR2^−/− mice compared to WT mice (Figure [71]2e–g). These
   results suggest that HCAR2 deficiency exacerbates S. aureus infection
   and induces a more intense inflammatory response in the mammary gland.

Figure 2.

   Figure 2
   [72]Open in a new tab

   HCAR2 deficiency exacerbates the severity of S. aureus infection in
   mouse mammary glands. a) Images of mouse mammary glands and H&E
   staining; red arrows indicate inflammatory cell infiltration, green
   arrows highlight thickened acinar walls, blue arrows point to tissue
   necrosis, and the scale bar represents 100 µm. b) Mammary injury
   scores. c) Bacterial burden in mammary tissue. d) MPO content in
   mammary tissue. e–g) ELISA quantification of IL‐1β, IL‐6, and TNF‐α
   protein levels in mammary tissue. h) TUNEL staining of mammary glands;
   red arrows mark TUNEL‐positive cells, scale bar represents 50 µm. i)
   Transmission electron microscopy of mitochondrial integrity; red arrows
   indicate mitochondria, and the scale bar represents 2 µm. j) Western
   blot analysis of GSDMD protein expression in mammary tissue. k) IHC of
   N‐GSDMD in mammary tissue, scale bar represents 50 µm. l) FITC‐albumin
   fluorescence in mammary tissue; green fluorescence indicates
   FITC‐albumin, blue fluorescence marks nuclei, and red arrows indicate
   FITC‐albumin diffusion in acini, scale bar represents 50 µm. m) Western
   blot analysis of ZO‐1 and Occludin protein expression in mammary
   tissue. Results are presented as mean ± SD, n = 6.

   Additionally, S. aureus infection led to an increase in cell death in
   the mammary glands of both WT and HCAR2^−/− mice. Notably, the number
   of dead cells was significantly higher in HCAR2^−/− mice than in WT
   mice (Figure [73]2h). Given the close association between cell death
   and mitochondrial damage, mitochondrial integrity in mMECs was
   assessed. Results revealed significantly more severe mitochondrial
   damage in HCAR2^−/− mice compared to WT mice (Figure [74]2i). S. aureus
   is known to induce pyroptosis, exacerbating tissue damage.^[ [75]^21 ^]
   Our results showed that S. aureus infection led to a marked increase in
   GSDMD activation in the mammary glands of both WT and HCAR2^−/− mice,
   with the latter, showing a more pronounced effect (Figure [76]2j,k).
   The blood‐milk barrier plays a critical role in maintaining
   intramammary stability, and epithelial cell pyroptosis accelerates its
   disruption. FITC‐albumin diffusion assays revealed that S. aureus
   infection induced albumin leakage in the mammary glands of both WT and
   HCAR2^−/− mice. However, the leakage was significantly more pronounced
   in HCAR2^−/− mice than in WT mice (Figure [77]2l). Additionally, S.
   aureus infection caused a reduction in tight junction (TJ) protein
   expression, with lower levels of these proteins observed in HCAR2^−/−
   mice compared to WT mice (Figure [78]2m). These results indicate that
   HCAR2 deficiency exacerbates pyroptosis and the disruption of the
   blood‐milk barrier during S. aureus infection in the mammary gland.

2.3. HCAR2 Deficiency enhances NLRP3 Inflammasome Activation in Mammary
Glands of S. aureus‐Infected Mice

   To further explore the regulatory mechanisms of HCAR2 in S.
   aureus‐induced mammary gland infection, transcriptomic analysis was
   performed to compare gene expression changes in the mammary glands of
   WT and HCAR2^−/− mice following S. aureus infection. The analysis
   revealed 527 upregulated and 892 downregulated genes in the mammary
   glands of HCAR2^−/− mice compared to WT mice (Figure [79]3a). GO
   enrichment analysis indicated that these differential genes are
   involved in bacterial defense responses, macrophage chemotaxis, and
   activation of GPCRs (Figure [80]3b). KEGG enrichment analysis
   highlighted significant differences in pathways such as S. aureus
   infection, Toll‐like receptor signaling, and NOD‐like receptor
   signaling (Figure [81]3c). The gene‐pathway enrichment network diagram
   demonstrated a close association between the Gram‐positive bacterial
   defense response, positive regulation of macrophage chemotaxis, and the
   NF‐κB signaling pathway with the NOD‐like receptor signaling pathway
   (Figure [82]3d). Given the pivotal role of the NOD‐like receptor
   signaling pathway in regulating pyroptosis, the activation of the NLRP3
   inflammasome was assessed. The results showed that in HCAR2^−/− mice,
   S. aureus stimulation led to significantly higher activation of the
   NLRP3 inflammasome compared to WT mice (Figure [83]3e). Clustering
   analysis of differential genes related to innate immune responses
   revealed significantly higher expression levels of NLRP3, TLR2, and
   Mefv in the mammary glands of HCAR2^−/− mice than in WT mice
   (Figure [84]3f). Furthermore, this study identified genes involved in
   regulating the NLRP3 inflammasome and validated several candidates
   (Figure [85]3g). The results demonstrated that in S. aureus‐infected
   HCAR2^−/− mice, the mRNA expression levels of CMPK2, Mefv, Irf1, and
   TLR2 were significantly elevated compared to those in WT mice
   (Figure [86]3h–l).

Figure 3.

   Figure 3
   [87]Open in a new tab

   HCAR2 deficiency enhances NLRP3 inflammasome activation in mammary
   glands of S. aureus‐infected mice. a) Transcriptome volcano plot of
   mammary gland tissues. “NoDiff” indicates no significant difference in
   gene expression between the two groups, while “Sig‐Up” and “Sig‐Down”
   represent upregulated and downregulated genes, respectively, in
   comparison to the NT group (b) GO enrichment analysis. c) KEGG
   enrichment analysis. d) Gene‐pathway enrichment network diagram. e)
   Western blot detection of NLRP3, ASC, and caspase‐1 protein levels in
   mammary tissue. f) Cluster analysis of differential genes associated
   with innate immune response. g) Protein‐protein interaction (PPI)
   analysis. h–l) mRNA expression levels of Mefv, Ncf1, CMPK2, Irf1, and
   TLR2 in mammary tissues. Results are presented as mean ± SD, n = 3.

2.4. Nia exerts a Significant Alleviating Effect on S. aureus Infection in
the Mammary Glands of Mice

   Based on the results above, the potential alleviating effect of HCAR2
   activation on S. aureus‐induced mastitis was explored. Nia treatment
   significantly ameliorated the pathological changes in mammary gland
   tissue induced by S. aureus stimulation (Figure [88]4a,b). Moreover,
   Nia treatment led to a marked reduction in MPO content, bacterial
   burden, and pro‐inflammatory cytokine levels in the mammary tissue
   (Figure [89]4c–g). Additionally, Nia alleviated the downregulation of
   TJ proteins caused by S. aureus infection in the mammary glands
   (Figure [90]4h,i). Furthermore, Nia treatment notably reduced
   FITC‐albumin diffusion within the mammary ducts, a hallmark of barrier
   disruption induced by S. aureus (Figure [91]4j). Further analysis
   revealed that Nia inhibited the increase in dead cells triggered by S.
   aureus infection in the mammary gland (Figure [92]4k). Nia also
   significantly decreased the up‐regulation of N‐GSDMD expression in
   mammary tissue induced by S. aureus (Figure [93]4l,m). Furthermore, Nia
   treatment inhibited the activation and initiation of the NLRP3
   inflammasome induced by S. aureus in the mammary glands
   (Figure [94]4n). These results suggest that HCAR2 activation, through
   Nia, has a substantial alleviating effect on S. aureus infection in the
   mammary glands of mice.

Figure 4.

   Figure 4
   [95]Open in a new tab

   Nia has a significant alleviating effect on S. aureus infection in the
   mammary glands of mice. a) Images of mouse mammary glands and H&E
   staining; red arrows indicate inflammatory cell infiltration, green
   arrows highlight thickened acinar walls, blue arrows point to tissue
   necrosis, and the scale bar represents 100 µm. b) Mammary injury score.
   c) Bacterial burden in mammary tissue. d) MPO content in mammary
   tissue. e–g) ELISA detection of IL‐1β, IL‐6, and TNF‐α protein levels
   in mammary tissue. h) IHC of ZO‐1 and Occludin in mammary tissue, scale
   bar represents 50 µm. i) Western blot detection of ZO‐1 and Occludin
   protein expression in mammary tissue. j) FITC‐albumin fluorescence in
   mammary tissue, with green fluorescence representing FITC‐albumin, blue
   fluorescence representing the nucleus, and red arrows indicating
   FITC‐albumin diffusion in the acini, scale bar representing 50 µm. k)
   TUNEL staining in the mammary gland, with red arrows indicating
   TUNEL‐positive cells, the scale bar represents 50 µm. l) IHC of N‐GSDMD
   in mammary tissue, scale bar represents 50 µm. n) Western blot
   detection of NLRP3, ASC, and caspase‐1 protein expression in mammary
   tissue. m) Western blot detection of ZO‐1, Occludin, and Claudin 3
   protein expression in mammary tissue. Results are presented as mean ±
   SD, n = 6.

2.5. Activation of HCAR2 Mitigates Pyroptosis and Tight Junction Damage
Induced by S. aureus in mMECs

   The mMECs, neutrophils, and macrophages interact to maintain immune
   homeostasis within the mammary gland. To investigate how HCAR2
   regulates cell‐cell interactions and influences the mammary immune
   response, S. aureus was injected into the mammary glands of WT and
   HCAR2^−/− mice at various time points. The results revealed that,
   before the substantial recruitment of immune cells, mMECs in HCAR2^−/−
   mice underwent pyroptosis earlier than those in WT mice (Figure [96]S2,
   Supporting Information). To further examine the role of HCAR2 in mMECs
   infected with S. aureus, Nia and siHCAR2 were used to activate or knock
   down HCAR2, respectively (Figures [97]1, [98]5). Nia treatment
   inhibited the upregulation of pro‐inflammatory factors induced by S.
   aureus through HCAR2 (Figure [99]S3a–c, Supporting Information).
   Additionally, Nia enhanced the expression of TJ proteins in S.
   aureus‐infected mMECs via HCAR2 (Figure [100]5b; Figure [101]S3d,e,
   Supporting Information), suggesting that HCAR2 activation mitigates the
   inflammatory response and TJ disruption induced by S. aureus in mMECs.

Figure 5.

   Figure 5
   [102]Open in a new tab

   Activation of HCAR2 mitigates pyroptosis and tight junction damage
   induced by S. aureus in mMECs. a) Schematic representation of the S.
   aureus infection model in vitro. b) Immunofluorescence images of ZO‐1
   and Occludin in mMECs, with scale bar representing 100 µm. c) TUNEL
   staining of mMECs, with red fluorescence indicating TUNEL‐positive
   cells, scale bar represents 200 µm. d) Quantitative analysis of
   TUNEL‐positive cells. e) LDH release assay to measure LDH levels in
   mMECs. f) Transmission electron microscopy (TEM) images showing
   cellular morphology of mMECs, with red arrows indicating mitochondria
   and blue arrows indicating cell membranes. The scale bar represents
   5 µm. g) Western blot analysis of GSDMD expression in mMECs. h) Western
   blot analysis of NLRP3, ASC, and caspase‐1 expression in mMECs. i)
   Immunofluorescence staining for CK18 in primary mammary epithelial
   cells, with scale bar representing 100 µm. j) Western blot analysis of
   HCAR2 expression in primary mMECs. k) Western blot analysis of GSDMD
   expression in primary mMECs. l) TUNEL staining of primary mMECs, with
   red fluorescence indicating TUNEL‐positive cells, scale bar represents
   200 µm. m) Quantitative analysis of TUNEL‐positive cells in mMECs. n)
   LDH release assay to measure LDH levels in primary mMECs. Results are
   presented as mean ± SD, n = 3.

   In vitro TUNEL staining results showed that Nia reduced cell death
   induced by S. aureus through HCAR2 activation. Furthermore, Nia
   inhibited the release of LDH from mMECs following S. aureus infection
   via HCAR2 (Figure [103]5c–e). Morphological analysis of the cells
   revealed that S. aureus infection induced swelling and bubble‐like
   protrusions in mMECs, which were significantly ameliorated by HCAR2
   activation (Figure [104]5f). GSDMD expression and activation were
   significantly increased in mMECs upon S. aureus stimulation, and Nia
   reduced GSDMD expression and activation through HCAR2 (Figure [105]5g).
   Additionally, Nia significantly decreased the expression of NLRP3, ASC,
   caspase‐1, and cleaved caspase‐1 in mMECs through HCAR2
   (Figure [106]5h). Co‐localization analysis showed that S. aureus
   stimulation induced co‐localization of ASC and caspase‐1 in mMECs,
   whereas Nia inhibited this co‐localization via HCAR2 (Figure [107]S3f,
   Supporting Information). Finally, primary mMECs isolated from WT and
   HCAR2^−/− mice were characterized (Figure [108]5i,j). It was
   demonstrated that Nia also alleviated S. aureus‐induced pyroptosis in
   primary mMECs through HCAR2 activation (Figure [109]5k–n). The
   detection of pro‐inflammatory factors, tight junctions, and NLRP3
   inflammasome components in primary mMECs confirmed the results observed
   in previous experiments (Figure [110]S4a–h, Supporting Information).
   These results further support that HCAR2 activation mitigates
   pyroptosis and tight junction damage induced by S. aureus in mMECs.

2.6. HCAR2 Inhibits Pyroptosis and Tight Junction Damage in mMECs Induced by
S. aureus through Downregulation of CMPK2 Expression

   Given that in vivo experiments revealed exacerbated mitochondrial
   damage induced by S. aureus upon HCAR2 deletion, the mitochondrial
   status and function in mMECs were further evaluated. S. aureus induced
   the loss of mitochondrial cristae and rupture of the mitochondrial
   membrane in mMECs, whereas Nia mitigated this mitochondrial damage
   through HCAR2 activation (Figure [111]6a). Furthermore, Nia restored
   the mitochondrial membrane potential reduced by S. aureus via HCAR2
   (Figure [112]6b). Similarly, results from primary mMECs suggested that
   activating HCAR2 aids in repairing mitochondrial damage caused by S.
   aureus (Figure [113]S4i, Supporting Information). Mitochondrial damage
   typically leads to the release of mtDNA, which functions as a DAMP to
   activate the NLRP3 inflammasome within the cell. To assess this, dsDNA
   and mitochondria were stained, revealing that Nia reduced the release
   of mitochondrial DNA (mtDNA) from mitochondria in S. aureus‐infected
   mMECs through HCAR2 activation (Figure [114]6c). Furthermore,
   disparities in the dsDNA content across groups led to the investigation
   of mtDNA replication. Subsequent experiments demonstrated that S.
   aureus stimulates enhanced mtDNA replication in mMECs, while Nia
   suppressed mtDNA replication via HCAR2 activation (Figure [115]6d,e).
   Previous research has shown that CMPK2 regulates mtDNA replication and
   contributes to inflammatory diseases by promoting this process.^[
   [116]^22 ^] Based on in vivo sequencing results, CMPK2 levels were
   assessed in mMECs, revealing that HCAR2 activation effectively
   suppressed S. aureus‐induced CMPK2 overexpression in both mMECs and
   primary mMECs (Figure [117]6f,g; Figure [118]S4j, Supporting
   Information).

Figure 6.

   Figure 6
   [119]Open in a new tab

   HCAR2 inhibits pyroptosis and tight junction damage in mMECs induced by
   S. aureus through downregulation of CMPK2 expression. a) Transmission
   electron microscopy was used to observe the mitochondrial morphology in
   mMECs, with red arrows indicating mitochondria and a scale bar of 2 µm.
   b) The mitochondrial membrane potential of mMECs was assessed using the
   JC‐1 membrane potential assay kit, with a scale bar of 100 µm. c) dsDNA
   antibodies and Mitotracker were used to label dsDNA and mitochondria,
   followed by imaging with a laser confocal microscope. Red arrows
   indicate co‐localization of dsDNA and mitochondria, with a scale bar of
   2 µm. d, e) qPCR was performed to detect specific primers for mtDNA
   (D‐loop, ND2) and nDNA (Tert), and to determine the relative content of
   mtDNA. f) qRT‐PCR was used to measure mRNA levels of CMPK2 in mMECs. g)
   Western blot analysis was employed to detect CMPK2 protein expression
   in mMECs. h) TUNEL staining of mMECs, with red fluorescence
   representing TUNEL‐positive cells and a scale bar of 200 µm. i)
   Statistical analysis of TUNEL‐positive cells. j) LDH release was
   assessed using the LDH assay kit in mMECs. k) Western blot analysis was
   used to measure GSDMD protein expression in mMECs. l) dsDNA antibodies
   and Mitotracker were used to label dsDNA and mitochondria, followed by
   imaging with a laser confocal microscope. Red arrows indicate
   co‐localization of dsDNA and mitochondria, with a scale bar of 2 µm. m)
   Immunofluorescence of ZO‐1 and Occludin in mMECs, with a scale bar of
   100 µm. Results are presented as mean ± SD, n = 3.

   Subsequently, the involvement of CMPK2 in S. aureus‐induced mMECs was
   investigated. The effects of siCMPK2 and OE‐CMPK2 on mMECs were
   evaluated, revealing that OE‐CMPK2 significantly increased CMPK2
   expression in S. aureus‐stimulated mMECs, while siCMPK2 markedly
   reduced CMPK2 expression (Figure [120]S5a–e, Supporting Information).
   Moreover, siCMPK2 and OE‐CMPK2 modulated pyroptosis in mMECs, with
   siCMPK2 mitigating and OE‐CMPK2 exacerbating pyroptosis induced by S.
   aureus (Figure [121]6h,i). LDH release assays corroborated these
   observations (Figure [122]6j). Additionally, OE‐CMPK2 promoted GSDMD
   activation, whereas siCMPK2 inhibited its activation (Figure [123]6k).
   In S. aureus‐stimulated mMECs, OE‐CMPK2, and siCMPK2 respectively
   enhanced and suppressed mtDNA release (Figure [124]6l). Both inhibition
   and overexpression of CMPK2 significantly decreased or increased mtDNA
   replication, respectively (Figure [125]S5f,g, Supporting Information).
   Moreover, OE‐CMPK2 and siCMPK2 exacerbated and ameliorated TJ loss in
   mMECs induced by S. aureus (Figure [126]6m; Figure [127]S5h,i,
   Supporting Information). Additionally, OE‐CMPK2 and siCMPK2 upregulated
   and downregulated the activation of the NLRP3 inflammasome in S.
   aureus‐stimulated mMECs, respectively (Figure [128]S5j, Supporting
   Information). These results collectively demonstrate that activation of
   HCAR2 can inhibit CMPK2 over‐expression, thereby alleviating pyroptosis
   and tight junction damage caused by S. aureus in mMECs.

2.7. The Release of mMEC‐Derived Factors Following S. aureus Infection
Modulates Macrophage Function and Activity via the cGAS/STING Pathway

   Given that pyroptosis in mMECs precedes immune cell recruitment into
   the acinar lumen, it was hypothesized that pyroptosis in mMECs might
   also influence the functionality of native immune cells. Primary
   peritoneal macrophages and neutrophils were isolated from WT mice
   (Figure [129]7a; Figure [130]S6a, Supporting Information). To replicate
   the immune microenvironment in which macrophages and neutrophils
   migrate to a mammary gland infected with S. aureus, macrophages were
   exposed to the supernatant from mMEC cultures infected with S. aureus.
   The results revealed that the S. aureus group exhibited decreased
   bactericidal activity compared to the NT group, while the S. aureus +
   Nia group demonstrated enhanced bactericidal ability relative to the S.
   aureus group. However, the bactericidal ability of the S. aureus + Nia
   + siHCAR2 group was lower than that of the S. aureus + Nia group
   (Figure [131]7b). No significant difference was observed in the
   bactericidal ability of neutrophils among the groups (Figure [132]S6b,
   Supporting Information). In contrast to the bactericidal ability, the
   release of pro‐inflammatory factors from macrophages showed an opposite
   trend (Figure [133]7c–e). Given that macrophage function is closely
   linked to cellular activity, the activity of each macrophage group was
   assessed. The results showed that cell activity in the S. aureus group
   was significantly lower than that in the NT group, whereas cell
   activity in the S. aureus + Nia group was higher than in the S. aureus
   group. The S. aureus + Nia + siHCAR2 group exhibited lower cell
   activity than the S. aureus + Nia group (Figure [134]7f,g).
   Interestingly, neutrophil activity displayed a similar trend to that of
   macrophages (Figure [135]S6c,d, Supporting Information). Further
   analysis revealed that the expression of clv‐caspase‐1, p‐MLKL, and
   N‐GSDMD was significantly upregulated in the S. aureus group compared
   to the NT group, while expression was downregulated in the S. aureus +
   Nia group compared to the S. aureus group. In contrast, expression was
   up‐regulated in the S. aureus + Nia + siHCAR2 group compared to the S.
   aureus + Nia group (Figure [136]7h–j). These results suggest that
   co‐stimulation can trigger extensive PCDs in macrophages and that
   reducing mMEC pyroptosis via HCAR2 may alleviate PCDs in co‐stimulated
   macrophages.

Figure 7.

   Figure 7
   [137]Open in a new tab

   The release of mMEC‐derived factors following S. aureus infection
   modulates macrophage function and activity via the cGAS/STING pathway.
   a) The purity of primary peritoneal macrophages was assessed using
   F4/80 and CD11b labeling. After 6‐h incubation with supernatants from
   various mMEC groups, macrophages were subsequently infected with S.
   aureus for an additional 6 h. b) Macrophage bactericidal activity
   against S. aureus was measured by plate counting. c–e) ELISA was
   employed to quantify IL‐1β, IL‐6, and TNF‐α levels in the macrophage
   culture supernatant. f) Flow cytometry was performed to assess cell
   viability. g) The proportion of viable macrophages was determined in
   each group. h–j) Western blot analysis was used to detect the protein
   expression of cleaved caspase‐1, p‐MLKL, MLKL, and GSDMD. k) Western
   blot analysis was used to detect the protein expression of cGAS and
   STING. l) Immunofluorescence staining for STING in macrophages was
   visualized, with red arrows indicating STING localization (scale bar =
   200 µm). Macrophages were pre‐incubated with 5 µm H‐151 for 1 h,
   followed by exposure to supernatants from the S. aureus‐treated mMEC
   group for 6 h, and subsequent infection with S. aureus for 6 h. m) Flow
   cytometry was again used to measure cell viability. n) The percentage
   of viable macrophages was quantified in each group. o) The bactericidal
   capacity of macrophages against S. aureus was assessed by plate
   counting. Results are presented as mean ± SD, n = 3.

   Additionally, significant alterations were observed in the cGAS/STING
   signaling axis of macrophages and neutrophils following co‐stimulation,
   with changes correlating with the expression of PCD‐related proteins
   (Figure [138]7k,l; Figure [139]S6e, Supporting Information). To further
   investigate this, the STING antagonist H‐151 was used to suppress STING
   signaling in macrophages and measured associated proteins. The results
   showed that H‐151 significantly inhibited the secretion of
   pro‐inflammatory factors in the S. aureus group (Figure [140]S7a–c,
   Supporting Information). Macrophage activity and bactericidal ability
   in the S. aureus group were significantly improved by H‐151
   (Figure [141]7m–o). Additionally, H‐151 reduced the expression of
   clv‐caspase‐1, p‐MLKL, and N‐GSDMD in the S. aureus group (Figure
   [142]S7d,e, Supporting Information). These results underscore the role
   of STING in inducing widespread PCDs in macrophages through
   co‐stimulation.

2.8. Extracellular mtDNA is Critical for Activating the cGAS/STING Signaling
Axis in Macrophages, with HCAR2 Deficiency Rendering them More Sensitive to
mtDNA

   Macrophage purity was assessed, revealing no difference between
   macrophages isolated from HCAR2^−/− and WT mice (Figure [143]S8a,
   Supporting Information). Subsequently, macrophages were stimulated with
   S. aureus and the supernatants from mMEC cultures to evaluate their
   bactericidal activity (Figure [144]S8b, Supporting Information). The
   results indicated that the bactericidal capacity of macrophages from
   HCAR2^−/− mice was comparable to that of WT mice, but the bactericidal
   ability in all other groups was significantly lower than that of WT
   mice, except the NT group (Figure [145]8a). In addition, the secretion
   of pro‐inflammatory factors was significantly elevated in macrophages
   from all groups except the NT group, compared to the WT group (Figure
   [146]S8c–e, Supporting Information). Assessment of cell viability
   revealed a marked reduction in macrophage activity in HCAR2^−/− mice in
   the S. aureus, S. aureus + Nia, and S. aureus + Nia + siHCAR2 groups,
   relative to WT mice (Figure [147]8b,c). cGAS/STING signaling changes in
   macrophages from HCAR2^−/− mice mirrored those in WT mice, but the
   expression levels of cGAS and STING were significantly lower in
   macrophages from HCAR2^−/− mice in all groups, except the NT group
   (Figure [148]8d). These results suggested that HCAR2‐deficient
   macrophages are more susceptible to the supernatant from S.
   aureus‐infected mMEC cultures.

Figure 8.

   Figure 8
   [149]Open in a new tab

   Extracellular mtDNA is critical for activating the cGAS/STING signaling
   axis in macrophages, with HCAR2 deficiency rendering them more
   sensitive to mtDNA. Macrophages were incubated with the supernatants
   from mMECs of various groups for 6 h, followed by infection with S.
   aureus for an additional 6 h. a) Macrophage bactericidal activity
   against S. aureus was assessed using a plate counting method. b) Cell
   viability was analyzed by flow cytometry. c) The percentage of normally
   active macrophages in each group was determined. d) Western blot
   analysis was performed to measure the protein expression of cGAS and
   STING. e, f) The copy number of D‐loop and ND2 in the supernatants of
   each group of mMEC cultures was quantified. Following incubation of
   macrophages with varying concentrations of mtDNA for 12 h, further
   experiments were carried out. g) Western blot was used to assess cGAS
   and STING protein levels. h) Immunofluorescence staining of STING in
   macrophages. The red arrow indicates STING, with a scale bar of 200 µm.
   Statistical significance is indicated as follows: ## p < 0.01 compared
   to the NT group; ^* and ^** p < 0.05 and p < 0.01, respectively,
   compared to the S. aureus group; & and && p < 0.05 and p < 0.01,
   respectively, compared to the S. aureus + Nia group; $ and $$ p < 0.05
   and p < 0.01, respectively, compared to WT mice. Results are expressed
   as mean ± SD, n = 3.

   Building on previous studies indicating that HCAR2 activation reduces
   CMPK2 expression in mMECs infected with S. aureus, leading to decreased
   mtDNA replication and pyroptosis, it was hypothesized that the
   variation in mtDNA released from mMECs could alter cGAS/STING signaling
   in macrophages. The data demonstrated that mtDNA copy numbers were
   lower in the S. aureus+Nia group compared to the S. aureus group, while
   the S. aureus+Nia+siHCAR2 group exhibited higher mtDNA levels than the
   S. aureus group (Figure [150]8e,f). This suggests that HCAR2 activation
   in S. aureus‐infected mMECs reduces mtDNA release. Further
   investigation, wherein mtDNA was extracted from mouse mammary tissue
   and used to stimulate macrophages at various concentrations, revealed
   that increasing mtDNA concentrations upregulated the protein expression
   of cGAS and STING in macrophages. Macrophages lacking HCAR2 were more
   responsive to mtDNA stimulation (Figure [151]8g). Immunofluorescence
   analysis of STING also confirmed these findings (Figure [152]8h).
   Moreover, mtDNA stimulation induced the secretion of pro‐inflammatory
   factors by macrophages, with HCAR2‐deficient macrophages producing
   higher levels of these factors (Figure [153]S6f–h, Supporting
   Information). Additionally, in vivo, data demonstrated that the absence
   of HCAR2 enhances the activation of the cGAS/STING pathway and promotes
   widespread PCDs in the mammary glands of S. aureus‐infected mice
   (Figure [154]S9, Supporting Information). Collectively, these results
   indicate that mtDNA released by S. aureus‐infected mMECs activates the
   cGAS/STING signaling axis in macrophages, impairing their function and
   activity.

2.9. CMPK2 Mediates the Alleviating Effect of Nia on S. aureus Infection in
the Mammary Gland of Mice

   Based on the previous findings, alterations in CMPK2 expression within
   mammary tissue were further investigated, revealing that Nia
   significantly inhibited the protein levels of CMPK2, cGAS/STING, and
   key executive proteins involved in PCDs in S. aureus‐induced mastitis
   in mice (Figure [155]S10, Supporting Information). To determine whether
   HCAR2 mediates the resolution of mastitis caused by S. aureus in the
   mammary gland via CMPK2, CMPK2 was reintroduced into the mammary gland
   using adenovirus vectors, and the effects on mastitis amelioration by
   the HCAR2 agonist were assessed. Upon injection of two adenoviruses,
   AAV‐NC and AAV‐CMPK2, into the mammary ducts of mice, in vivo imaging
   demonstrated robust fluorescence signals in the mammary gland of the
   AAV9‐treated group, compared to the negative control (0.9% NaCl),
   confirming successful colonization of the mammary glands by the
   adenovirus vector (Figure [156] 9a,b). Further analysis showed stable
   expression of both AAV9‐NC and AAV9‐CMPK2 in mammary tissues, with
   AAV9‐CMPK2 significantly increasing CMPK2 protein expression in the
   gland (Figure [157]9c,d). Next, Nia was used to activate HCAR2, and a
   mastitis model was established with S. aureus (Figure [158]9e). The
   findings indicated that while Nia alleviated the symptoms of S.
   aureus‐induced mastitis and reduced the bacterial load in the mammary
   gland, this improvement was significantly reversed upon CMPK2
   overexpression (Figure [159]9f–h). Additionally, MPO and
   pro‐inflammatory factor assays showed that CMPK2 overexpression in the
   mammary glands of S. aureus‐infected mice diminished the inhibitory
   effect of Nia on MPO and pro‐inflammatory factors (Figure [160]9i–l).
   Further analysis of the blood‐milk barrier revealed that CMPK2 weakened
   Nia's protective effects on this barrier (Figure [161]9m,n). Lastly,
   assessments of cell pyroptosis in mammary tissue showed that CMPK2
   compromised Nia's ability to reduce cell pyroptosis and weakened Nia's
   inhibitory effect on NLRP3 inflammasome activation (Figure [162]9o–q).
   These results collectively demonstrate that CMPK2 mediates the
   alleviating effects of Nia on S. aureus infection in the mammary gland.

Figure 9.

   Figure 9
   [163]Open in a new tab

   CMPK2 mediates the alleviating effect of Nia on S. aureus infection in
   the mammary gland of mice a) Flowchart illustrating intra‐mammary
   inoculation with adenovirus. b) In vivo imaging of the mammary gland
   following injection of 0.9% NaCl or adenovirus. c) Intramammary
   fluorescence detection of AAV9‐NC and AAV9‐CMPK2 after inoculation into
   the mammary gland, with green fluorescence indicating adenovirus and
   blue fluorescence representing the nucleus; scale bar represents 50 µm.
   d) Western blot analysis of CMPK2 protein expression in mammary tissue.
   e) Flowchart of the animal experimental procedure. f) Representative
   images of mouse mammary gland and H&E staining; red arrows indicate
   inflammatory cell infiltration, green arrows indicate thickening of the
   acinar wall; scale bar, 100 µm. g) Mammary injury score. h) Bacterial
   load in mammary tissue. i) MPO content in mammary tissue. j–l) ELISA
   analysis of IL‐1β, IL‐6, and TNF‐α protein levels in mammary tissue. m)
   Western blot analysis of ZO‐1 and Occludin protein expression in
   mammary tissue. n) IHC for ZO‐1 and Occludin in mammary tissue; scale
   bar, 50 µm. o) IHC for N‐GSDMD in mammary tissue; scale bar, 50 µm. p)
   Western blot analysis of GSDMD protein expression in mammary tissue. q)
   Western blot analysis of NLRP3, ASC, and caspase‐1 protein expression
   in mammary tissue. Results are presented as mean ± SD, n = 6.

3. Discussion

   The innate immune system, composed of epithelial cells and macrophages,
   plays a critical role in the early detection and elimination of
   pathogens that invade mammary tissue. However, epithelial cell death or
   an exaggerated immune response from macrophages can exacerbate
   pathological progression during infection.^[ [164]^23 , [165]^24 ^]
   This study demonstrates that activation of HCAR2 in S. aureus‐infected
   mMECs leads to a reduction in CMPK2/mtDNA expression, which in turn
   inhibits NLRP3 inflammasome activation and mitigates cell pyroptosis.
   The attenuation of pyroptosis in mMECs reduces the release of mtDNA
   into the extracellular space, thereby alleviating the suppression of
   macrophage activity induced by S. aureus via the cGAS/STING‐dependent
   pathway. These findings offer new insights into the pathogenesis of S.
   aureus mastitis and highlight potential therapeutic targets for
   managing S. aureus‐associated infections.

   GPCRs are a crucial class of cell surface receptors with significant
   regulatory roles in bacterial infections across various organs.^[
   [166]^25 ^] In the respiratory tract, the activation of GPCRs,
   particularly OXGR1 expressed on respiratory epithelial cells, is
   essential for maintaining epithelial cell function and regulating the
   innate immune response during bacterial infection.^[ [167]^26 ^]
   Similarly, research by Naina Gour et al. has shown that Mrgpra1, a GPCR
   located on the surface of neutrophils, plays a critical role in
   modulating excessive neutrophil responses during acute lung infections
   by regulating neutrophil polarization.^[ [168]^27 ^] To investigate
   GPCRs with immune regulatory functions during S. aureus infection,
   transcriptomic analysis revealed significant upregulation of HCAR2
   expression in the mammary gland during infection. Previous studies by
   Guo et al. have demonstrated that the absence of HCAR2 exacerbates
   sepsis progression, while its immunoregulatory function has been
   confirmed in inflammatory conditions, such as enteritis and
   neuroinflammation.^[ [169]^20 , [170]^28 , [171]^29 ^] Our results
   indicate that HCAR2 deficiency exacerbates the inflammatory response,
   compromises barrier integrity, and promotes increased bacterial
   colonization within the mammary gland. In contrast, administration of
   HCAR2 agonists alleviated these pathological changes.

   HCAR2 is highly expressed not only in macrophages and neutrophils but
   also in mMECs, where it plays a critical role in regulating milk fat
   composition.^[ [172]^19 , [173]^30 ^] As the predominant cell type in
   the breast, mMECs are integral to both milk secretion and the formation
   of the blood‐milk barrier, contributing significantly to immune
   regulation and the maintenance of the mammary gland's internal
   environment.^[ [174]^31 ^] Disruption or dysfunction of these cells
   compromises the blood‐milk barrier, alters the mammary gland's internal
   environment, and adversely affects immune cell function.^[ [175]^11 ,
   [176]^32 ^] Numerous studies have confirmed that the death of mMECs
   during bacterial infections accelerates the destruction of the
   blood‐milk barrier, leading to tissue damage.^[ [177]^33 , [178]^34 ^]
   Findings from both in vivo and in vitro experiments in this study
   indicate that HCAR2 activation can restore the integrity of the
   blood‐milk barrier damaged by S. aureus. Previous research has
   highlighted the role of pyroptosis in combating immune evasion by
   intracellular bacteria^[ [179]^35 ^]; however, excessive pyroptosis can
   result in irreversible tissue damage and systemic inflammatory
   consequences.^[ [180]^36 ^] Recent studies by Chao Wei have emphasized
   the importance of endothelial cell pyroptosis in the disruption of the
   inflammatory blood‐brain barrier,^[ [181]^37 ^] while Xiaozhou Wang
   et al. demonstrated that S. aureus triggers pyroptosis in mMECs.^[
   [182]^12 ^] This study shows that HCAR2 activation significantly
   mitigates S. aureus‐induced pyroptosis in mMECs.

   The NLRP3 inflammasome can be activated by intracellular S. aureus,
   leading to the assembly of a protein complex comprising NLRP3, ASC, and
   caspase‐1. This complex subsequently activates caspase‐1 to cleave
   caspase‐1 and GSDMD, producing the N‐terminal fragment of GSDMD,
   ultimately resulting in pyroptosis.^[ [183]^38 ^] In this study, it was
   found that in mice with S. aureus‐induced mammary gland infections, the
   deletion of HCAR2 enhances the activation of the NLRP3 inflammasome,
   while HCAR2 activation mitigates NLRP3 activation triggered by S.
   aureus. Mitochondrial damage plays a pivotal role in the induction of
   pyroptosis and inflammation. Mitochondrial components and metabolic
   byproducts, released as DAMPs into the extracellular space, further
   propagate inflammation in infectious diseases.^[ [184]^39 ^] Upon
   mitochondrial damage, mtDNA undergoes oxidation and translocates to the
   cytoplasm, where it directly interacts with NLRP3 to activate the
   inflammasome.^[ [185]^40 ^] Suyuan Liu et al. demonstrated that
   inhibiting mitochondrial DNA synthesis effectively modulates
   inflammatory responses mediated by the NLRP3 inflammasome.^[ [186]^41
   ^] Similarly, Xing Gao et al. showed that S. aureus induces
   mitochondrial damage in mMECs, leading to elevated mitochondrial
   reactive oxygen species (mtROS) levels and electron transport chain
   (ETC) dysfunction.^[ [187]^42 ^] This study further demonstrates that
   HCAR2 activation, both in vivo and in vitro, alleviates S.
   aureus‐induced mitochondrial damage and significantly reduces the
   upregulation of mtDNA duplication in S. aureus‐stimulated mMECs.

   CMPK2, a mitochondrial nucleotide monophosphate kinase essential for
   deoxyribonucleoside triphosphate (dNTP) biosynthesis,^[ [188]^43 ,
   [189]^44 ^] has been shown to regulate mtDNA synthesis and inhibit
   NLRP3 inflammasome activation.^[ [190]^45 ^] Previous studies have
   indicated that targeting CMPK2 to reduce mtDNA synthesis can
   effectively suppress NLRP3 inflammasome activation, thus mitigating
   pneumonia and sepsis infections.^[ [191]^22 ^] In this study, CMPK2 was
   identified as a downstream target of HCAR2, and modulation of CMPK2
   expression in mMECs was found to influence both NLRP3 inflammasome
   activation and pyroptosis in these cells. Meanwhile, CMPK2 mediated the
   alleviating effect of Nia on S. aureus infection in the mammary gland
   of mice. In mMECs stimulated with S. aureus, CMPK2 expression was
   closely associated with mitochondrial stability, possibly linked to
   CMPK2's role in regulating GSDMD activation. Rui Miao et al.
   demonstrated that N‐GSDMD interacts with the outer mitochondrial
   membrane, leading to mitochondrial damage within cells.^[ [192]^37 ^]
   Nevertheless, the precise regulatory mechanism by which HCAR2
   influences CMPK2 remains to be elucidated. This will also serve as the
   subsequent experimental direction for this study.

   This study demonstrates that both WT and HCAR2^−/− mice exhibit
   pyroptosis in the mammary glands following S. aureus infection, with
   the latter group displaying a more pronounced degree of pyroptosis.
   Additionally, the bacterial load in the mammary glands of HCAR2^−/−
   mice is significantly elevated, suggesting that HCAR2 may regulate
   epithelial pyroptosis and influence the bactericidal capacity of immune
   cells. Previous research has indicated that the interaction between
   alveolar epithelial cells and macrophages can impair macrophage
   phagocytic function in chronic obstructive pulmonary disease.^[
   [193]^46 ^] Similarly, hepatocyte pyroptosis exacerbates acute liver
   injury by enhancing macrophage‐mediated inflammatory responses.^[
   [194]^47 ^] A significant correlation between mMEC apoptosis and
   macrophage activity has also been reported, particularly highlighting
   M2 macrophages as key contributors to epithelial cell death during
   postpartum breast tissue remodeling.^[ [195]^48 ^] However, the role of
   the interaction between mammary epithelial cell death and macrophages
   in breast immunity remains largely unexplored. While previous studies
   suggest that pyroptosis in epithelial or endothelial cells can enhance
   macrophage bactericidal functions by preventing bacterial immune
   evasion,^[ [196]^49 ^] our findings confirm that pyroptosis of mMECs
   exacerbates inflammatory responses in S. aureus‐infected macrophages,
   reducing their activity and bactericidal capacity. Furthermore, this
   study shows that HCAR2 modulates pyroptosis in mMECs to alleviate the
   reduction in macrophage activity induced by S. aureus. Research by Wang
   Y and Chen X et al. has shown that macrophage death leads to an
   increased release of pro‐inflammatory mediators, impairing their
   bactericidal capacity, whereas promoting macrophage survival can reduce
   inflammation and restore cellular function.^[ [197]^50 , [198]^51 ^]
   Therefore, the diminished bactericidal capacity of macrophages in
   HCAR2^−/− mice is likely linked to reduced macrophage activity, which
   may also explain the observed increase in both bacterial load and
   inflammatory responses in the mammary glands of these mice.

   Due to its ability to be activated by both endogenous and exogenous
   DNA, combined with the predominant expression of STING in immune cells,
   the cGAS‐STING signaling pathway plays a critical role in mediating
   inflammatory responses and bacterial infections.^[ [199]^52 ^]
   Furthermore, the involvement of cGAS‐STING in modulating various forms
   of PCDs)in macrophages has been well‐documented.^[ [200]^53 , [201]^54
   ^] This study demonstrates that the reduction in macrophage activity
   caused by mMEC pyroptosis is regulated through the cGAS‐STING signaling
   pathway. Previous studies have shown that hepatocyte pyroptosis
   promotes the release of mtDNA into the extracellular space, thereby
   activating the cGAS‐STING pathway in macrophages.^[ [202]^47 ^] The
   present study demonstrated that pyroptosis in mMECs facilitates the
   extracellular release of mtDNA, which subsequently activates the
   cGAS‐STING signaling pathway in macrophages. Both in vivo and in vitro
   analyses revealed that HCAR2^−/− macrophages exhibit an enhanced
   capacity to activate the cGAS‐STING pathway. However, the exact
   mechanisms driving the increased susceptibility of HCAR2^−/−
   macrophages to mtDNA remain to be clarified.

   In conclusion, our findings demonstrate that the activation of HCAR2 in
   S. aureus‐infected mMECs effectively reduces CMPK2/mtDNA expression,
   thereby inhibiting NLRP3 inflammasome activation and mitigating
   pyroptosis. The attenuation of pyroptosis in mMECs leads to a decreased
   release of mtDNA, which subsequently inhibits the activation of the
   cGAS/STING signaling pathway in macrophages, alleviating the reduced
   macrophage activity induced by S. aureus in a cGAS/STING
   pathway‐dependent manner (Figure [203]10 ). These results not only
   elucidate the role of HCAR2 in enhancing resistance to S. aureus
   infection but also provide insight into the underlying mechanisms,
   establishing HCAR2 as a potential therapeutic target for S.
   aureus‐induced mastitis.

Figure 10.

   Figure 10
   [204]Open in a new tab

   HCAR2 modulates the crosstalk between mammary epithelial cells and
   macrophages to mitigate S. aureus infection in the mouse mammary gland.

4. Experimental Section

Experimental Animals and Design

   In this experiment, 9‐week‐old wild type (WT) C57BL/6 were obtained
   from Liaoning Changsheng Biotechnology Co. and HCAR2^−/−mice were
   generous gift from Dr. Martin Sager (Zentrale Einrichtung für
   Tierforschung und Tierschutzaufgaben der Heinrich‐Heine Universität
   Düsseldorf, Germany). One male and one female mouse were placed in the
   same cage and allowed to eat and drink freely. After the mice became
   pregnant, the pregnant mice were transferred to a new cage for the
   experiment. All animal experiments were carried out in strict adherence
   to the guidelines set forth by the Animal Welfare Protection and
   Utilization Committee of Jilin University (SY202409026).

   The mouse experiments were divided into four parts. 1) Within 12–14
   days postpartum, WT mice were randomly assigned to two groups: the NT
   group and the S. aureus group, with 6 mice per group. All mice were
   separated from their pups 2 h before model induction. The S. aureus
   group was used to establish the S. aureus‐induced mastitis model. 2)
   Within 12–14 days postpartum, WT mice were randomly divided into two
   groups: the WT group and the WT+S. aureus group, with 6 mice per group.
   HCAR2^−/− mice were also divided into two groups: the HCAR2^−/− group
   and the HCAR2^−/−+S. aureus group, with 6 mice per group. The WT+S.
   aureus and HCAR2^−/−+S. aureus groups were used to establish the S.
   aureus‐induced mastitis model. 3) Within 5–7 days postpartum, WT mice
   were randomly assigned to three groups: the NT group, the S. aureus
   group, and the Nia + S. aureus group, with 6 mice per group. Mice in
   the Nia + S. aureus group were administered a 50 mm nicotinic acid
   (Nia) solution dissolved in drinking water for 7 days before model
   induction. Both the S. aureus and Nia + S. aureus groups were used to
   establish the S. aureus‐induced mastitis model. 4) Within 1 day
   postpartum, WT mice were randomly assigned to four groups: the NT
   group, the S. aureus group, the Nia + S. aureus group, and the Nia + S.
   aureus + CMPK2 group, with 6 mice per group. All mice received a 50 µL
   adenovirus injection into the subcutaneous mammary tissue on the 1st
   and 7th days postpartum, with a viral titer of 1 × 10^12 V.g mL^−1.
   AAV9‐NC was injected into the NT, S. aureus, and Nia + S. aureus
   groups, while AAV9‐CMPK2 was injected into the Nia + S. aureus + CMPK2
   group. Seven days before model induction, the Nia + S. aureus and Nia +
   S. aureus + CMPK2 groups were administered a 50 mm nicotinic acid
   solution in their drinking water. The S. aureus, Nia + S. aureus, and
   Nia + S. aureus + CMPK2 groups were used to establish the S.
   aureus‐induced mastitis model. D‐luciferin sodium salt was employed as
   a luminescent substrate to observe adenovirus replenishment in the
   mammary gland tissue using an in vivo small animal 3D visible light
   imaging system (IVIS Spectrum). The Living Image 4.3 software was used
   for post‐imaging analysis.

S. aureus‐Induced Mastitis Mice Model Construction

   Mice were anesthetized by intraperitoneal injection of 45 mg kg^−1
   pentobarbital sodium, and once anesthesia was achieved, they were
   placed supine and secured. The region surrounding the fourth pair of
   mammary glands was disinfected with 75% ethanol. Using fine tweezers,
   the nipple was raised, and ≈2 mm of the tip was excised under an
   anatomical microscope. A 100 µL micropipette was employed to inject
   50 µL of a 1 × 10^8 CFU mL^−1 S. aureus suspension into the nipple (the
   S. aureus suspension was prepared in PBS at the appropriate
   concentration).^[ [205]^55 ^] In control groups, 50 µL of sterile PBS
   was injected into the mammary gland. After 24 h, all mice were humanely
   euthanized. The fourth pair of mammary glands were then exposed,
   carefully observed, and photographed. Mammary tissue samples were
   collected for subsequent analysis.

Transcriptome Sequencing

   Following the collection of mouse mammary tissues, sequencing and data
   analysis were conducted by Lianchuan Biomedical Technology Co
   (Hangzhou, China). In summary, after collecting mouse mammary tissue,
   RNA was extracted and purified, and its concentration and purity were
   assessed using the NanoDrop ND‐1000. The RNA was then
   reverse‐transcribed to generate complementary DNA (cDNA). The RNA‐DNA
   hybrid was subsequently converted into double‐stranded DNA, with the
   ends capped using specialized reagents. The double‐stranded DNA was
   digested with UDG enzyme, and a library was constructed using PCR
   amplification, targeting fragment sizes of 300 bp ±50 bp. Sequencing
   was performed using the Illumina NovaSeq 6000 (LC Bio‐Technology Co.,
   Ltd., Hangzhou, China), following standard protocols. Sequencing data
   were aligned to the Homo sapiens GRCh38 genome using HISAT2
   ([206]https://ccb.jhu.edu/software/hisat2). Differential gene
   expression analysis was conducted with the R package edgeR
   ([207]https://bioconductor.org/packages/release/bioc/html/edgeR.html).
   GO and KEGG pathway enrichment analyses were performed using DAVID
   software ([208]https://david.ncifcrf.gov/).

Hematoxylin−Eosin (H&E) Staining

   The tissue sections were deparaffinized using xylene, followed by
   rehydration with a series of ethanol solutions. The sections were
   subsequently stained with hematoxylin and eosin, dehydrated in ethanol,
   cleared in xylene, and ultimately mounted with neutral resin. The
   criteria for the mammary injury score were established based on a
   previous study.^[ [209]^56 ^]

Immunohistochemical (IHC)

   After the sections of mammary tissue were dewaxed and hydrated, immerse
   the sections in citrate buffer for antigen retrieval. Subsequently, the
   sections were rinsed with PBS and a peroxidase blocker was applied. The
   sections were inoculated with 5% donkey serum to block, followed by
   incubation with primary antibody diluted in 5% donkey serum at 4 °C
   overnight. Then the immunohistochemistry kit (Biological Technology,
   Wuhan, China) was used for immunohistochemical staining, and finally,
   neutral resin was used to mount the slides.

Immunofluorescence (IF)

   After the sections of mammary tissue were dewaxed and hydrated, the
   antigen was repaired using citrate buffer, and the endogenous
   peroxidase was blocked using a paraffin embedding compound. After the
   cells were inoculated and treated on the cell crawl, they were fixed
   with immunostaining fixative. The subsequent procedures were conducted
   in a manner consistent with the cellular protocol, involving surface
   blocking using 5% donkey serum, subsequent incubation with the primary
   antibody, and application of a fluorescent secondary antibody. Finally,
   the coverslips containing DAPI were used for staining and mounting, and
   fluorescence microscopy or laser scanning microscopy was used to
   observe the results.

Blood‐Milk Barrier Integrity Assessment via FITC‐Albumin

   FITC‐albumin (Sigma–Aldrich, USA) was dissolved in PBS, and freshly
   collected mammary tissue from each group was immersed in the
   FITC‐albumin solution for 45 min. Tissue samples were then processed
   for frozen sectioning, and the green fluorescence within the acini was
   visualized under a fluorescence microscope. Detailed experimental
   procedures could be found in a previous publication.^[ [210]^57 ^]

Detection of Myeloperoxidase (MPO) Activity

   A little mammary tissue was taken and weighed, then the solution of
   N‐2‐Hydroxyethylpiperazine‐N‐2‐Ethane sulfonic acid 5000 times the
   weight was added to the tissue with magnetic beads, and then ground
   with automatic grinder for 10 min. After centrifugation, the
   supernatant was absorbed as a sample of elisa. Then, an equal volume of
   5% cetyltrimethylammonium chloride solution was added to the HEPEs
   solution and continued grinding for 10 min. The supernatant was
   collected as the sample of MPO. Sequentially 75 µL of the sample was
   added to each well of the 96‐well plate, followed by the addition of
   75 µL of MPO substrate coloring solution. Terminate the color reaction
   after 3 min by adding 2 m H[2]SO[4] solution.

Enzyme‐Linked Immunosorbent Assay (ELISA)

   The instructions of the Elisa kit (Biolegend, San Diego, CA, USA) were
   followed for detecting IL‐1β, IL‐6, and TNF‐α in the mammary tissue of
   mice or the supernatant of macrophages.

Western Blot

   The tissue or cells were resuspended in NP40 lysate and transfer 2 mL
   into EP tubes. Following lysis, centrifuge to collect the supernatant
   as the protein sample, then utilize the BCA reagent kit (Thermo
   Scientific, USA) for protein concentration determination. Subsequently,
   aliquot the samples into separate tubes based on their protein
   concentration. Then, the 12% sodium dodecyl sulfate‐polyacrylamide gel
   electrophoresis (SDS‐PAGE) was performed. After transferring the
   membrane using the PVDF membrane, incubate the PVDF membrane with the
   primary antibody and secondary antibody. Finally, the
   electrochemiluminescence detection kit (Biyuntian, Wuhan, China) was
   used for the visualization of the protein bands by using an
   autoradiography apparatus. The information on antibodies used in this
   study is listed in Table [211]S1 (Supporting Information).

Quantitative Real‐Time Polymerase Chain Reaction (q‐RTPCR)

   The total mRNA of tissue or cell was extracted by using Trizol (Thermo
   Fisher Scientific, Shanghai, China). Subsequently, RNA reverse
   transcription was conducted utilizing the cDNA First‐Strand Synthesis
   Kit (ABclonal, Wuhan, China). Finally, the Quantitect SYBR Green RT‐PCR
   kit (Roche, Shanghai, China) was used for q‐RTPCR. The primer sequence
   is listed in Table [212]S2 (Supporting Information).

S. aureus Infection of Cells

   The mouse mammary epithelial cell line, EpH4‐Ev cells, was cultured in
   DMEM medium (Gibco, Grand Island, NY, USA) supplemented with 10% fetal
   bovine serum (Hyclone, USA). Cells in Petri dishes were counted, and S.
   aureus was added at a multiplicity of infection (MOI) of 10. After a
   2‐h incubation with the bacteria, the medium was discarded, and the
   cells were washed twice with PBS. Gentamicin was then added to the
   medium at a final concentration of 100 µg mL^−1 for 1 h, followed by
   two PBS washes. Fresh medium was added, and cells were cultured for an
   additional 4 or 6 h before the medium or cells were collected for
   further analysis.

Measurement of Total mtDNA

   Cells were collected, and total DNA was extracted using the Cell Fast
   DNA Extraction Kit (ABclonal, Wuhan, China) according to the
   manufacturer's instructions. Total mtDNA was quantified by qPCR using
   specific primers targeting the D‐loop region of mitochondria and other
   uninsured mtDNA regions. Nuclear DNA encoding Tert served as a
   normalization control. Detailed procedures could be found in previous
   studies.^[ [213]^22 ^] The primer sequences are provided in Table
   [214]S2 (Supporting Information).

Cell Viability

   One‐step TUNEL In Situ Apoptosis Kit (Green, FITC) (Elabscience, Wuhan,
   China) was used to detect dead cells in the mammary gland. One Step
   TUNEL Apoptosis Assay Kit (Biyuntian, Wuhan, China) was used to detect
   dead cells of mMECs. Lactate dehydrogenase (LDH) Cytotoxicity Assay Kit
   (Biyuntian, Wuhan, China) was used to detect LDH released by cells. The
   Annexin V‐FITC Apoptosis Detection Kit (Biyuntian, China) was employed
   for assessing cell viability. All experiments were carried out in
   accordance with the instructions provided by the kit following cell
   treatment.

RNA Interference and Overexpression

   RNA interference and overexpression were performed using Lipofectamine
   2000 (Thermo Fisher Scientific Co., Shanghai, China). Briefly, siRNA or
   overexpression plasmid and Lipofectamine 2000 were diluted in Opti‐MEM
   and then mixed for 20 min. The mixture was added to the cell culture
   dish, the medium was replaced after 6 h, and the cells were cultured
   for an additional 24 h. siHCAR2, siCMPK2, and OE‐CMPK2 were synthesized
   by Shanghai Genechem Co. Ltd. The primer sequences are listed in Table
   [215]S3 (Supporting Information).

Mitochondrial Probe Staining

   Mitotracker Red CMXRos (Beyotime, Wuhan, China) was diluted in DMSO to
   a final concentration of 200 µm. The Mitotracker Red CMXRos solution
   was added to the cell culture medium at a 1:1000 dilution, and cells
   were incubated for 20 min. Cells were then fixed with an immunofixative
   solution, followed by immunofluorescence staining and subsequent
   procedures.

Mitochondrial Membrane Potential Detection

   The mitochondrial membrane potential assay kit with JC‐1 (Biyuntian,
   Wuhan, China) was used to detect changes in cell intima potential.
   Briefly, after the cells were treated, the cells medium was discarded
   and the JC‐1 dyeing solution was mixed with the new DMEM medium at a
   ratio of 1:1, and the cells were incubated with the mixture for 20 min.
   Then the cell medium was discarded, the cells were washed with JC‐1
   staining buffer twice, and the results were observed by fluorescence
   microscopy.

Extraction of Primary Mammary Epithelial Cells

   Lactating mice were euthanized and their mammary glands were immersed
   in 75% ethanol for 30 s. The glands were dissected, and the lymph nodes
   within the tissue were removed. Mammary tissue was placed in a D‐Hanks
   solution containing antibiotics and washed at least 5 times. After
   removing the excess solution, the tissue was minced into small pieces.
   The tissue was transferred to 15 mL centrifuge tubes and incubated with
   trypsin at 37 °C for 2 h with shaking. After digestion, the samples
   were centrifuged at 1500 rpm for 10 min, the supernatant was discarded,
   and the pellet was resuspended in a 5 mL DMEM culture medium.
   Centrifugation at 1500 rpm for 5 min followed, and the supernatant was
   discarded. To eliminate red blood cells, 2 mL of Red Blood Cell Lysis
   Buffer was added at 37 °C and incubated for 3 min. Following a final
   centrifugation at 1500 rpm for 5 min, the supernatant was discarded.
   The cells were resuspended in a DMEM culture medium, filtered through a
   40 µm cell strainer, and cultured for identification.

Extraction of Primary Peritoneal Macrophages

   Thioglycolate broth (3 mL) was injected into the peritoneal cavity of
   mice 3 days and 1 day before the experiment. Following euthanasia and
   immersion in 75% ethanol for 3 min, the mice were placed on a sterile
   operating table and 3 mL of complete 1640 culture medium was injected
   into the peritoneal cavity. The peritoneal cavity was gently massaged,
   and the peritoneal fluid was aspirated. The collected fluid was
   centrifuged at 1000 rpm for 5 min, and the supernatant was discarded.
   The pellet was resuspended in red blood cell lysate and mixed
   thoroughly. After another centrifugation at 1000 rpm for 5 min, the
   supernatant was discarded, and the pellet was resuspended in a DMEM
   medium. The suspension was centrifuged at 1000 rpm for 3 min, the
   supernatant was discarded, and the pellet was resuspended in a complete
   medium. After cell counting, the cell density was adjusted to 2 × 10^6
   cells mL^−1, and the cells were seeded into 6‐well plates or 3 cm^2
   culture dishes according to experimental requirements.

Extraction of Primary Peritoneal Neutrophils

   Mice received two injections of 1 mL thioglycolate broth into the
   peritoneal cavity, with a 12‐h interval between injections. 3 h after
   the second injection, the mice were euthanized, and 3 mL of 1640
   culture medium was infused into the peritoneal cavity. The peritoneal
   cavity was gently massaged for 2 min, and the peritoneal wash fluid was
   aspirated into 15 mL centrifuge tubes, repeating this step twice. The
   tubes were centrifuged at 1000 rpm for 10 min, and the supernatant was
   discarded. The collected cells were resuspended in a 1640 culture
   medium and red blood cell lysate. After adding granulocyte separation
   liquid, the mixture was centrifuged at 1500 rpm for 20 min. The lower
   white cell layer was collected into another 15 mL centrifuge tube and
   resuspended in a 5 mL 1640 culture medium. The suspension was
   centrifuged at 500 rpm for 5 min, the supernatant was discarded, and
   the step was repeated. The collected cells were analyzed for purity
   using fluorescently conjugated antibodies, including Ly6G‐FITC and
   CD11b‐APC.

Extraction of mtDNA from Mammary Tissue

   The Mitochondrial DNA Isolation Kit (Abcam, USA) was used to isolate
   mtDNA from mammary tissue. Briefly, mammary tissue was added with an
   appropriate amount of cytosol extraction buffer. After grinding, the
   mammary tissue was incubated for 10 min, centrifuged at 700 g for
   10 min, supernatant was taken, and centrifuged at 10000 g for 30 min.
   After discarding the supernatant, the precipitation was suspended in
   the cytoplasmic extraction buffer briefly and centrifuged for 30 min.
   The supernatant was discarded and added to the mitochondrial cleavage
   buffer, followed by enzyme mix and ethanol for incubation for 10 min.
   Finally, the supernatant was discarded after centrifugation to obtain
   mtDNA precipitation.

The Bactericidal Ability of Macrophages

   Following 6 h incubation of macrophages with various culture
   supernatants of mMECs, the culture medium was replaced, and S.aureus
   (MOI = 5) was introduced to infect the macrophages for 2 h. After
   washing the cells with the culture medium twice, 100 µg mL^−1 of
   gentamicin should be added to the culture medium, followed by
   incubation for 1 h. After washing the cells with culture medium twice,
   0.25% Triton‐100 sterile water was added to the culture hole for cell
   lysis. The sterile water was used to be coated on the LB solid culture
   plate, and the number of internalized bacteria (CFU0) was determined.
   In the other wells, incubate the cells for another 6 h, add 0.25%
   Triton‐X‐100 sterile water to lyse the cells, and calculate the number
   of bacteria (CFU2) in the sterile water. The intracellular bactericidal
   activity of macrophages is expressed as (CFU0−CFU2)/CFU0×100%.

Data and Statistical Analysis

   Data were presented as the mean ± standard deviation (SD). Graphs and
   statistical analyses were performed using GraphPad Prism 8 software.
   The Shapiro–Wilk test was used to assess data distribution normality.
   The F‐test assessed the equality of variances between the two groups,
   while the paired t‐test with unequal variances was applied to compare
   the two groups. The Brown–Forsythe test evaluated variance homogeneity
   across multiple groups, and one‐way ANOVA was used to determine
   statistical differences among multiple groups. ^* indicates p < 0.05,
   representing a significant difference between groups; ^** indicates p
   < 0.01, representing a highly significant difference between groups.

Conflict of Interest

   The authors declare no conflict of interest.

Author Contributions

   X.R., J.L., and S.F. performed conceptualization. W.G. and S.F.
   performed resources. X.R., K.L., Y.L., W.G., and X.W. performed the
   methodology. X.R., B.Y., and S.F. performed visualization. B.Y. and
   J.L. performed supervision. X.R. and K.L. wrote the original draft.
   X.R., J.L., and S.F. did review & editing.

Supporting information

   Supporting Information
   [216]ADVS-12-2411947-s001.docx^ (5.2MB, docx)

Acknowledgements