Abstract

   Cardiac fibrosis, a key pathological feature of cardiac remodeling, is
   a major contributor to mortality in older patients with heart failure.
   The underlying mechanisms are complex, involving alterations in
   intercellular communication and chronic inflammation. This study
   investigates the role of indole‐3‐propionic acid (IPA) in aging‐related
   myocardial fibrosis and its regulatory effects on autophagy through
   palmitoyl‐protein thioesterase 1 (PPT1). Here, plasma levels of IPA, a
   tryptophan‐derived metabolite, are found to be reduced in older
   patients with heart failure, and this reduction is associated with
   deteriorating cardiac function. Notably, IPA supplementation
   significantly attenuated aging‐related myocardial fibrosis. PPT1, a
   lysosomal enzyme involved in autophagy, is upregulated in macrophages
   during aging. IPA reversed aging‐induced increase in PPT1 expression.
   Using PPT1^flox/flox Lyz2‐cre mice, it is demonstrated that
   macrophage‐specific deletion of PPT1 significantly reduced cardiac
   inflammation and myocardial fibrosis in aged mice. Furthermore, PPT1
   silencing in macrophages reduced the expression of myocardial fibrosis
   markers in vitro. Mechanistically, IPA regulated PPT1 expression to
   modulate the PI3K‐AKT‐mTOR pathway, thereby restoring autophagic
   activity in senescent macrophages and suppressing both inflammation and
   aging‐related myocardial fibrosis. Additionally, IPA influenced the
   cGAS‐STING signaling pathway to regulate PPT1 expression. These
   findings demonstrate that IPA inhibits PPT1, activates autophagy in
   macrophages, and mitigates aging‐related myocardial fibrosis.

   Keywords: aging‐related cardiac fibrosis, autophagy, IPA, macrophages,
   palmitoyl‐protein thioesterase 1
     __________________________________________________________________

   IPA is an intestinal tryptophan metabolite whose effects decline with
   decreased heart function. Supplementing IPA can alleviate the
   aging‐related myocardial fibrosis through PPT1. PPT1 is a key protein
   localized to lysosomes, and IPA can restore macrophage autophagy
   function by regulating PPT1 expresssions, thereby reducing
   aging‐related myocardial fibrosis and inflammation.

   graphic file with name ADVS-12-e01070-g008.jpg

1. Introduction

   Heart failure (HF) remains a leading cause of morbidity and mortality
   worldwide, with its incidence and prevalence increasing consistently.^[
   [56]^1 ^] Pathological remodeling processes lead to phenotypic changes
   in cardiac fibroblasts, exacerbating extracellular matrix deposition.
   This excessive matrix accumulation reduces cardiac tissue compliance
   and ultimately contributes to the progression of advanced HF.^[ [57]^2
   ^] Aging is a significant risk factor for myocardial fibrosis, playing
   a crucial role in its development and progression.^[ [58]^3 ^] However,
   the precise mechanisms underlying aging‐related myocardial fibrosis
   have not been fully elucidated. Consequently, identifying novel
   therapeutic targets is essential for developing effective
   interventions. Emerging evidence suggests that alterations in gut
   microbiota significantly influence the pathophysiology of
   cardiovascular diseases.^[ [59]^4 , [60]^5 ^] To facilitate genetic and
   molecular analyses, we utilized a rodent model of aging. Our findings
   revealed a reduction in circulating and cardiac tissue levels of
   indole‐3‐propionic acid (IPA) during aging, a gut‐derived metabolite
   synthesized from tryptophan by Clostridium sporogenes.^[ [61]^6 ^]
   Tryptophan, an essential amino acid, plays a key role in maintaining
   physiological equilibrium. IPA has been shown to regulate mitochondrial
   activity and modulate cardiac function.^[ [62]^7 ^] Additionally, it
   has demonstrated cardioprotective effects by preserving ejection
   fraction.^[ [63]^8 ^] However, whether IPA inhibits myocardial fibrosis
   in older individuals remains unclear.

   Owing to the aging of the immune system, older individuals frequently
   experience low‐grade chronic inflammation, which is associated with an
   increased risk of cardiovascular disease.^[ [64]^9 ^] Studies have
   demonstrated a positive correlation between age and the number of
   cardiac macrophages.^[ [65]^10 , [66]^11 ^] In aging mice, the number
   of resident cardiac macrophages declines while the contribution of
   circulating blood monocytes to cardiac macrophage populations
   increases.^[ [67]^12 ^] Macrophages play a central role in promoting
   inflammation by secreting proinflammatory molecules such as interleukin
   (IL)‐6, tumor necrosis factor (TNF)‐α, and matrix metalloproteinases
   (MMPs), which collectively contribute to cardiac dysfunction.^[ [68]^13
   , [69]^14 ^] Our study investigated macrophage‐mediated effects on the
   heart to identify potential therapeutic targets.

   Palmitoyl‐protein thioesterase 1 (PPT1) is mainly located within
   lysosomes, where it regulates the degradation and clearance of
   lysosomal hydrolases, thereby influencing autophagy.^[ [70]^15 ,
   [71]^16 ^] PPT1 plays a critical role in disease pathogenesis, as it
   modulates the lysosomal degradation of nucleotide‐binding domain‐like
   receptor protein 3 (NLRP3). By reducing the degradation of NLRP3, PPT1
   enhances its activation, leading to the release of inflammatory factors
   and ultimately triggering the proinflammatory phenotype of
   macrophages.^[ [72]^17 ^] Notably, genetic defects in PPT1 or
   pharmacologic inhibition of PPT1 have been shown to reduce autoantibody
   levels and attenuate systemic inflammation.^[ [73]^18 ^] PPT1 is a
   target of transcription factor EB (TFEB), the master regulator of
   autophagy and lysosomal biogenesis, which orchestrates lysosomal
   function via PPT1 regulation.^[ [74]^19 ^] Therefore, we utilized
   PPT1^flox/flox Lyz2‐cre mice to examine the regulatory effects of PPT1
   in the context of cardiovascular aging. In addition, we explored the
   effect of IPA on reducing inflammation and fibrosis in the aged model
   by regulating PPT1. This is the first study to investigate the role of
   PPT1 in aging‐related cardiovascular diseases.

   We further explored the interaction of IPA with PPT1 by PI3K‐AKT‐mTOR
   signaling pathway. Activation of mTOR complex 1 (mTORC1) has been
   implicated in the proliferation and expansion of inflammatory
   monocytes, thereby exacerbating systemic inflammation.^[ [75]^20 ^]
   Conversely, pharmacologic inhibition of mTORC1 with rapamycin has been
   shown to mitigate inflammatory responses.^[ [76]^21 ^] The cyclic
   GMP‐AMP synthase (cGAS)‐stimulator of interferon genes (STING) pathway
   is a critical danger‐sensing mechanism involved in innate immunity.^[
   [77]^22 ^] We hypothesize that IPA modulates PPT1 through the
   regulation of the cGAS‐STING pathway. STING activation in
   cardiomyocytes triggers downstream signaling cascades, leading to
   increased production of pro‐inflammatory cytokines and transforming
   growth factor‐beta (TGF‐β), which ultimately contribute to myocardial
   fibrosis.^[ [78]^23 ^] In this study, we investigated the specific
   mechanism by which IPA inhibits PPT1 activation and induces autophagy.

   Our findings indicate that IPA is downregulated in older patients with
   HF and that IPA supplementation reduces the degree of myocardial
   fibrosis in aged rats, suggesting its potential as a therapeutic agent
   for aging‐related myocardial fibrosis. Moreover, macrophages highly
   expressing PPT1 exhibit a pro‐inflammatory phenotype significantly
   upregulated in senescent macrophages. Furthermore, IPA regulates
   macrophage PPT1 via the cGAS‐STING pathway while simultaneously
   modulating the autophagic function of macrophages through the
   PI3K‐AKT‐mTOR pathway.

2. Results

2.1. Abnormal Expression of Tryptophan and its Metabolite IPA in Aged
Patients and Rats

   To evaluate plasma metabolite expression in older patients with heart
   failure, we performed metabolomic analysis on plasma samples from young
   (n = 14), older with non‐HF (n = 30), and older patients with heart
   failure (Older with HF, n = 30) (Table [79]S1, Supporting Information).
   A metabolite difference volcano plot (Figure [80]1A) and Kyoto
   Encyclopedia of Genes and Genomes (KEGG) pathway enrichment analysis
   revealed differential metabolites in human plasma, with tryptophan
   metabolism being significantly enriched in the older with HF group
   compared to the older with non‐HF group (Figure [81]1B). IPA was
   significantly downregulated in older with HF group compared with older
   with non‐HF group (Figure [82]1C). Correlation analysis revealed a
   negative correlation between IPA concentration and age (Figure [83]1D).
   Hematoxylin and eosin (H&E) staining revealed increased inflammation in
   the colons of the aged group (Figure [84]1E). To investigate gut
   microbiota composition, we collected and analyzed rat fecal samples
   using 16S rDNA sequencing. As shown in Figure [85]1F, the
   Firmicutes/Bacteroidota ratio was decreased in aged rats, indicating
   increased inflammation. Linear discriminant analysis effect size
   (LEfSe) analysis identified a significant difference in the relative
   abundance of c_Clostridia between groups (Figure [86]1G). Previous
   studies have shown that c_Clostridium is the primary bacteria
   responsible for metabolizing tryptophan into IPA (Figure [87]1H).^[
   [88]^24 ^] Next, we performed targeted quantitative metabolomic
   analysis of rat fecal samples to measure IPA levels (Figure [89]1I).
   Venn diagram revealed that IPA, nicotinic acid (N‐acid), 2‐Aminobenzoic
   acid (2‐AA), and 2‐Ketoadipic acid (2‐KA) were common differentially
   expressed metabolites (Figure [90]1J). Furthermore, IPA levels in heart
   tissues were downregulated with left ventricular ejection fraction
   (LVEF) value (Figure [91]1K). These findings indicate that gut
   microbiota composition is altered in aged rats with heart failure and
   that IPA, a key tryptophan metabolite, is significantly reduced in
   plasma and heart tissue with aging and deterioration cardiac function.

Figure 1.

   Figure 1
   [92]Open in a new tab

   Abnormal expression of tryptophan and its metabolite IPA in tissue
   samples from elderly patients and rats. A) Volcano plot of differential
   metabolites in Older with non‐HF vs Older with HF groups of human
   plasma samples. B) KEGG pathway enrichment map of differential
   metabolites in Older with non‐HF vs Older with HF groups of human
   plasma samples. C) Box plots of IPA content in Young (n = 14), Older
   with non‐HF (n = 30) and Older with HF (n = 30) groups of human plasma
   samples (^## p < 0.01 vs the Older with non‐HF group). D) Correlation
   scatter plot between human plasma IPA concentration and age. E) H&E
   staining of rat colon samples in Young and Aged groups. (n = 3)
   Magnification: 100×, scale bar = 100 µm. F) Stacked bar plots of
   relative abundance of species at phylum level based on ASV of rat feces
   samples in Young (n = 8) and Aged (n = 8) groups. G) ASV‐based
   cladogram of rat feces samples in Young and Aged groups. H) Metabolic
   pathway of IPA in the tryptophan metabolic pathway of rat feces
   samples. I) Heat map of differential metabolites of rat feces samples
   in Young (n = 8), Aged (LVEF>70%) (n = 8), and Aged (LVEF≤70%) (n = 8)
   groups. J) Venn plot of differential metabolites of rat feces samples
   in Young, Aged (LVEF>70%), and Aged (LVEF≤70%) groups. K) Correlation
   scatter plot about rat heart IPA content and LVEF(%) value in Young (n
   = 12) and Aged (n = 24) group.

2.2. IPA Inhibits the Secretion of Pro‐Inflammatory Factors in Senescent
Macrophages and Alleviates Myocardial Fibrosis in Aged Rats

   Aged rat models were established using 18‐ to 20‐month‐old
   Sprague‐Dawley (SD) rats (Figure [93]S1A, Supporting Information).
   Echocardiography was performed to assess cardiac function (Figure
   [94]S1B, Supporting Information), measuring LVEF, left ventricular
   fractional shortening (LVFS), left ventricular internal diameter at
   diastole (LVIDd), left ventricular internal diameter at systole
   (LVIDs), left ventricular posterior wall thickness at diastole (LVPWd),
   left ventricular posterior wall thickness at systole (LVPWs),
   interventricular septal end‐diastole (IVSd), and interventricular
   septal end‐systole (IVSs) were detected (Figure [95]S1C–J, Supporting
   Information). A significant decline in cardiac function was observed in
   aged rats. H&E and Masson's staining revealed disordered cardiac muscle
   fibers and increased myocardial fibrosis in aged hearts (Figure
   [96]S1K–M, Supporting Information). Additionally, aged rats exhibited
   an upregulation of senescence markers P21 and P16, along with increased
   levels of fibrosis‐related genes COL1A1 and COL3A1 and pro‐inflammatory
   markers IL‐6 and CD86 in aged rats (Figure [97]S1N,O, Supporting
   Information). These findings confirm that aging leads to significant
   cardiac dysfunction and myocardial fibrosis.

   To investigate the therapeutic potential of IPA in aging‐related
   myocardial fibrosis, aged rats were administered IPA (20
   mg kg^−1 day^−1) by oral gavage for 8 weeks (Figure [98]2A).
   Echocardiography was used to monitor cardiac function (Figure [99]2B),
   with additional cardiac function indices measured (Figure [100]2C–J).
   Histological evaluation using H&E and Masson's staining demonstrated
   that IPA treatment mitigated myocardial fibrosis and improved cardiac
   muscle fiber organization (Figure [101]2K,L, Figure [102]2O).
   Immunohistochemistry (IHC) analysis using α‐SMA and COL1A1 antibodies
   revealed that IPA significantly reduced α‐SMA and COL1A1‐positive areas
   in aged heart tissue (Figure [103]2M,N,P,Q). COL3A1 and COL1A1 were
   significantly reduced in the IPA‐treated aged group (Figure [104]2R–T).
   The detection of the IPA concentration in the heart after intragastric
   administration is shown in Figure [105]S2A (Supporting Information).
   These findings suggest that IPA attenuates aging‐related myocardial
   fibrosis in aged rats.

Figure 2.

   Figure 2
   [106]Open in a new tab

   IPA alleviates aging‐related myocardial fibrosis in rats. A) Schematic
   of the experimental design. B) Representative echocardiographic graphs.
   C–J) Echocardiographic measurements of LVEF, LVFS, LVIDd, LVIDs, LVPWd,
   LVPWs, IVSd, and IVSs. K) H&E staining of rat hearts. Magnification:
   200×, scale bar=50 µm. L) Masson's staining of rat hearts.
   Magnification: 100×, scale bar = 100 µm. M,N) Representative images of
   IHC staining with α‐SMA or COL1A1 antibody. Magnification: 200×, scale
   bar = 50 µm. O) Statistical graphs of Masson's staining. P,Q)
   Statistical graphs of IHC staining with α‐SMA or COL1A1. R) Statistical
   graphs of RT‐qPCR of COL3A1 and COL1A1. S,T) Representative images and
   statistical graphs of COL3A1 and COL1A1 at protein levels. (n = 6–7,
   data are expressed as mean ± SEM, ^* p < 0.05, ^** p < 0.01 vs the
   Young group; ^# p < 0.05, ^## p < 0.01 vs the Aged group).

   To determine the specific cell types through which IPA exerts its
   cardioprotective effects, primary neonatal rat cardiomyocytes and
   cardiac fibroblasts were isolated and cultured. Cellular senescence was
   induced by treating these cells with 50 µm hydrogen peroxide (H[2]O[2])
   for 48h (Figure [107]2B–E). CIBERSORT analysis revealed compositional
   changes in immune cells within the ventricles of aged mice (Figure
   [108]S2F–H, Supporting Information). As shown in Figure [109]3A–F, the
   TNF‐α, CD68, and MCP‐1 positive areas were significantly reduced in the
   heart tissue of the Aged+IPA group, whereas these markers were notably
   increased in the Aged group. Immunofluorescence (IF) analysis further
   demonstrated M1 macrophages (CD68^+CD86^+) accumulated in the heart and
   bone marrow‐derived macrophages (BMDMs) with aging, but their numbers
   significantly decreased following IPA treatment (Figure [110]3G,H).
   Additionally, levels of IL‐6 and TNF‐α were increased in the Aged
   group, but IPA administration effectively reversed these increases
   (Figure [111]3I–K). Furthermore, IF staining showed a reduction in the
   intensity of iNOS in vitro (Figure [112]3L,M). To further confirm the
   anti‐inflammatory effects of IPA, we evaluated the expression of IL‐6
   and TNF‐α (Figure [113]3N–P). These findings suggest that IPA inhibits
   M1 macrophage polarization, suppresses the expression of inflammatory
   cytokines, and exerts a protective effect against cardiac fibrosis and
   inflammation in aging.

Figure 3.

   Figure 3
   [114]Open in a new tab

   IPA inhibits proinflammatory factor secretion in senescent macrophages.
   A–F) Representative images and statistical graphs of IHC staining with
   TNF‐α, CD68 and MCP‐1 antibody. Magnification: 200×, scale bar = 50 µm.
   G) Representative images of the immunofluorescence of CD68, CD86, and
   DAPI in rat heart tissue (CD68^+CD86^+ represent M1 macrophage).
   Magnification: 200×, scale bar = 20 µm. H) Representative images of the
   immunofluorescence of CD68, CD86, and DAPI in rat BMDM. Magnification:
   200×, scale bar = 20 µm. I) Statistical graphs showing RT‐qPCR results
   of IL‐6 and TNF‐α expression of rat hearts from the Young, Aged, and
   Aged+IPA groups. J,K) Representative images and statistical graphs of
   IL‐6 and TNF‐α at protein levels. L,M) Representative graphs of the
   immunofluorescence of iNOS in Control, LPS/IFN‐γ and LPS/IFN‐γ+IPA
   groups. Magnification: 200×, scale bar = 20 µm. N) Statistical graphs
   of RT‐qPCR of IL‐6 and TNF‐α in Control, LPS/IFN‐γ and LPS/IFN‐γ+IPA
   groups. O,P) Representative images and statistical graphs of IL‐6 and
   TNF‐α at protein levels in Control, LPS/IFN‐γ and LPS/IFN‐γ+IPA groups.
   (n = 3–6, data are expressed as mean ± SEM, ^** p < 0.01 vs the Young
   or Control group; ^## p < 0.01 vs the Aged or LPS/IFN‐γ group).

2.3. Knockout of Macrophage PPT1 can Reduce Cardiac Inflammatory Infiltration
and Myocardial Fibrosis in Aging Mice

   Macrophages are the predominant resident immune cells in the heart and
   play a crucial role in maintaining cardiac homeostasis.^[ [115]^25 ^]
   To investigate macrophage alterations during aging, we performed flow
   cytometry analysis, which revealed a significant increase in the M1/M2
   macrophage ratio (CD45^+CD86^+/ CD45^+CD163^+) in the Aged group
   (Figure [116]S3A,B, Supporting Information). Both protein and mRNA
   levels of P21 and TNF‐α were markedly upregulated in the aged group
   (Figure [117]S3C–F, Supporting Information), indicating enhanced
   inflammatory infiltration and a pro‐inflammatory phenotype in senescent
   macrophages. Proteomic sequencing revealed differences between the two
   groups (Figure [118]4A). KEGG pathway analysis revealed alterations in
   the lysosomal pathway in aged hearts, with a significant upregulation
   of PPT1 expression (Figure [119]4B,C). PPT1 positive area, mRNA, and
   protein levels were increased in aged heart tissue (Figure [120]4D–G).
   To analyze the inflammatory response of PPT1 in macrophages, we
   analyzed single‐cell RNA sequencing (scRNA‐seq) data from 27 healthy
   donors.^[ [121]^26 ^] Monocytes and macrophages were classified into
   seven subsets (Figure [122]4H) and the marker is shown in
   Figure [123]4I. Percentage of 5 celltypes of macrophages is shown in
   Figure [124]4J. The monocyte subset had a higher immune response, as
   demonstrated by the box plot analysis (Figure [125]4K). The M1 score
   box plot further confirmed that the Mφ‐5 subset had the highest M1
   polarization, with significantly elevated PPT1 expression in the Mφ‐5
   and monocyte subsets—identifying them as highly inflammatory cell
   subsets (Figure [126]4L,M). Subsequently, we found that PPT1^+ mice
   exhibited heightened immune responses and increased M1 scores
   (Figure [127]4N,O). IF staining confirmed a significant upregulation of
   PPT1 expression in CD68^+ macrophages in aged hearts and BMDMs
   (Figure [128]4P–R). Similarly, PPT1 levels were upregulated in aged
   macrophages (Figure [129]4S,T). Collectively, these findings indicate
   that aging is associated with an increased proportion of
   pro‐inflammatory monocytes and macrophages, with pro‐inflammatory
   PPT1^+ cells highly expressed in the heart.

Figure 4.

   Figure 4
   [130]Open in a new tab

   PPT1 expression is upregulated in senescent macrophages. A) Volcano
   plot of differentially expressed proteins of Young and Aged groups in
   rats heart. B) KEGG enrichment top 20. C) Heat map of differentially
   expressed proteins in the lysosomal pathway. D,E) Representative images
   and statistical analysis of IHC staining with PPT1 antibody.
   Magnification: 400×, scale bar = 100 µm. F) Statistical analysis of
   PPT1 mRNA level. G) Representative images and statistical analysis of
   PPT1 protein level. H) UMAP plots of heart macrophage from 27 healthy
   donors. I) A heatmap showing the markers ≈7 types of macrophages. J)
   Stacked graph of cell proportions. K) Violin plots of inflammatory
   response scores in different subgroups. L) Violin plots of the M1
   scores of different macrophage subsets. M) Violin plots of PPT1
   expression in different monocyte‐macrophage subsets. N) Inflammatory
   response score. O) M1 score. P) Representative figure of the
   immunofluorescence of CD68, PPT1, and DAPI in heart. Magnification:
   200×, scale bar = 20 µm. Q,R) Representative immunofluorescence images
   and statistical analysis of PPT1 intensity. Magnification: 200×, scale
   bar = 20 µm. S) Statistical analysis of PPT1 mRNA level in BMDM. T)
   Representative graphs and statistical analysis of PPT1 at protein level
   in BMDM. (n = 3–6, data are expressed as mean ± SEM, ^* p < 0.05, ^** p
   < 0.01 vs the Young group).

   To verify the role of PPT1 in the heart during aging, PPT1^flox/flox
   Lyz2‐cre mice (cko‐PPT1) were administered D‐galactose (D‐gal) for 12
   weeks to induce aging (Figure [131]5A). In addition, adeno‐associated
   virus (AAV)‐F4/80‐sh‐PPT1 was injected into the aged rats to induce
   PPT1 knockdown (Figure [132]S4A, Supporting Information).
   Echocardiography was performed to assess cardiac function
   (Figure [133]5B; Figure [134]S4B, Supporting Information). The results
   showed that PPT1 knockdown via either cko‐PPT1 or sh‐PPT1 attenuated
   the decline in cardiac function induced by D‐gal or aging
   (Figure [135]5C–J; Figure [136]S4C–J, Supporting Information). H&E and
   Masson's staining revealed that cko‐PPT1 or sh‐PPT1 reversed the
   inflammation, disordered muscle fibers, and excessive collagen
   deposition induced by D‐gal or aging (Figure [137]5K,L,O; Figure
   [138]S4K,L,O, Supporting Information). IHC further demonstrated that
   cko‐PPT1 or sh‐PPT1 reduced the D‐gal‐ or aging‐induced MCP‐1 and α‐SMA
   positive areas (Figure [139]5M,N,P,Q; Figure [140]S4M,N,P,Q, Supporting
   Information). Additionally, IF staining showed that macrophage PPT1
   (CD68^+PPT1^+) expression increased with aging, while cko‐PPT1 or
   sh‐PPT1 decreased the number of CD68^+PPT1^+ cells (Figure [141]5R;
   Figure [142]S4R, Supporting Information). Fibrotic and inflammatory
   markers were also downregulated in the D‐gal + cko‐PPT1 and aging +
   sh‐PPT1 groups (Figure [143]5S–U; Figure [144]S4S–U, Supporting
   Information). These findings indicate that cko‐PPT1 or sh‐PPT1 could
   reverse the aging‐induced increase in fibrosis and inflammation.

Figure 5.

   Figure 5
   [145]Open in a new tab

   Transgenic knockout of macrophage PPT1 improves cardiac inflammatory
   infiltration and myocardial fibrosis in D‐gal induce‐aged mice. A)
   Schematic of the experimental design. B) Representative
   echocardiographic graphs. C–J) Echocardiographic measurements of LVEF,
   LVFS, LVIDd, LVIDs, LVPWd, LVPWs, IVSd, and IVSs. K) H&E staining of
   left ventricles. Magnification: 200×, scale bar = 50 µm. L) Masson's
   staining of left ventricles. Magnification: 200×, scale bar = 50 µm.
   M,N) Representative images of IHC staining with MCP‐1 and α‐SMA
   antibody. Magnification: 200×, scale bar = 50 µm. O) Statistical graphs
   of Masson's staining. P,Q) Statistical graphs of IHC staining with
   MCP‐1 and α‐SMA antibody. R) Representative images of the
   immunofluorescence of CD68, PPT1, and DAPI in mice heart.
   Magnification: 200×, scale bar = 20 µm. S) Statistical analysis of
   COL3A1, COL1A1, PPT1, IL‐6, and TNF‐α mRNA level. T,U) Representative
   images and statistical analysis of COL3A1, COL1A1, PPT1, IL‐6, and
   TNF‐α at protein level. (n = 5–6, data are expressed as mean ± SEM, ^**
   p < 0.01 vs the WT group; ^# p < 0.05, ^## p < 0.01 vs the D‐gal
   group).

2.4. IPA Regulates PPT1 to Inhibit the Secretion of Pro‐Inflammatory Factors
in Senescent Macrophages

   To investigate the pro‐inflammatory effects of macrophage PPT1, we
   employed si‐PPT1 or DC661, a PPT1 inhibitor, and found that si‐PPT1#1
   had a better knockdown effect (Figure [146]S5A,B, Supporting
   Information). Subsequently, both si‐PPT1 and DC661 significantly
   reduced iNOS levels induced by LPS/IFN‐γ in vitro (Figures [147]S5C,D
   and [148]S6A,B, Supporting Information). Furthermore, we examined the
   impact of PPT1 knockdown on M1 macrophages using si‐PPT1 or DC661 and
   observed a significant reduction in their pro‐inflammatory activity
   (Figures [149]S5E,F and [150]S6C–E, Supporting Information). These
   findings suggest that the knockdown of PPT1 could significantly inhibit
   the pro‐inflammatory effects of M1 macrophages.

   To determine whether IPA mediates PPT1 regulation in macrophages, we
   assessed PPT1 expression in aged and M1 macrophages using IF and
   western blotting. Our results demonstrated that IPA reduced the
   expression of PPT1 both in vivo and in vitro (Figure [151]S7A–E,
   Supporting Information). This effect was further validated by PPT1
   overexpression (Figure [152]S7F,G, Supporting Information), followed by
   IPA treatment, which confirmed that IPA could reduce PPT1 levels
   despite its overexpression (Figure [153]S7H,I, Supporting Information).

   To further investigate whether IPA regulates senescent macrophages
   through PPT1, we first verified the anti‐inflammatory effects of IPA in
   aged macrophages, which showed a reduction in pro‐inflammatory marker
   levels (Figure [154]6A–E). Similarly, we found that IPA attenuated the
   pro‐inflammatory effects of oe‐PPT1 (Figure [155]6F–J). Collectively,
   these results provide compelling evidence that IPA could regulate the
   inflammatory response in aged macrophages through PPT1.

Figure 6.

   Figure 6
   [156]Open in a new tab

   IPA inhibits PPT1 expression and reduces the secretion of inflammatory
   factors in aging macrophages. A,B) Representative images and
   statistical graphs of the immunofluorescence of iNOS. Magnification:
   200×, scale bar = 20 µm. C) Statistical analysis of IL‐6 and TNF‐α at
   mRNA level. D,E) Representative images and statistical analysis of IL‐6
   and TNF‐α at protein level. F,G) Representative images and statistical
   graphs of the immunofluorescence of iNOS. Magnification: 200×, scale
   bar = 20 µm. H) Statistical analysis of IL‐6 and TNF‐α at mRNA level.
   I,J) Representative images and statistical analysis of IL‐6 and TNF‐α
   at protein level. (n = 3–6, data are expressed as mean ± SEM, ^** p <
   0.01 vs the Young or Vector group; ^# p < 0.05, ^## p < 0.01 vs the
   Aged or oe‐PPT1 group).

2.5. IPA Regulates PPT1 to Inhibit Senescent Macrophages from Inducing
Myocardial Fibrosis

   A previous study reported that macrophage‐fibroblast crosstalk plays an
   important role in the pathophysiology of cardiac inflammation and
   fibrosis.^[ [157]^27 ^] To further investigate the role of
   PPT1‐expressing macrophages in myocardial fibrosis, conditioned medium
   (CM) from M1 macrophages was harvested and added to cardiac fibroblasts
   (Figure [158]S8A, Supporting Information). BMDMs were extracted from
   young and aged rats, treated with si‐PPT1 or DC661, and their CM was
   collected and administered to cardiac fibroblasts. As shown in Figure
   [159]S8B,C,F,G (Supporting Information), PPT1 inhibition via si‐PPT1 or
   DC661 significantly reduced α‐SMA expression in cardiac fibroblasts.
   Western blot analysis further showed that incubation of aged BMDMs with
   si‐PPT1 or DC661 alleviated fibrosis in cardiac fibroblasts by
   decreasing the expression of COL3A1, COL1A1, and α‐SMA (Figure
   [160]S8D,E,H,I, Supporting Information). These findings indicate that
   inhibiting PPT1 in aged macrophages reduces collagen deposition in
   myocardial fibroblasts.

   Next, we assessed whether IPA regulates myocardial fibrosis through its
   effects on PPT1 in the macrophages of aged rats. We measured fibrosis
   marker expression levels in cardiac fibroblasts which influenced by
   Young, Aged, and Aged + IPA macrophages and found that Aged + IPA‐CM
   reversed the Aged‐CM‐induced upregulation of α‐SMA (Figure [161]7A–F).
   Similarly, IPA treatment reversed the oe‐PPT1‐induced increases in
   COL3A1, COL1A1, and α‐SMA levels (Figure [162]7G–K). These findings
   suggest that IPA regulates the crosstalk between macrophages and
   fibroblasts via PPT1, thereby attenuating fibroblast fibrosis.

Figure 7.

   Figure 7
   [163]Open in a new tab

   IPA inhibits PPT1 in aged macrophages and alleviates collagen
   deposition in fibroblasts. A) Schematic of the experimental design.
   B,C) Representative figures and statistical graphs of the
   immunofluorescence of α‐SMA. Magnification: 200×, scale bar = 20 µm. D)
   Statistical analysis of COL3A1, COL1A1 and α‐SMA at mRNA level. E,F)
   Representative images and statistical analysis of COL3A1, COL1A1 and
   α‐SMA at protein level. G,H) Representative Figure and statistical
   graphs of the immunofluorescence of α‐SMA. Magnification: 200×, scale
   bar = 20 µm. I) Statistical analysis of COL3A1, COL1A1 and α‐SMA at
   mRNA level. J,K) Representative figures and statistical analysis of
   COL3A1, COL1A1 and α‐SMA at protein level. (n = 3–6, data are expressed
   as mean ± SEM, ^* p < 0.5, ^** p < 0.01 vs the Young–CM or Vector–CM
   group; ^# p < 0.5, ^## p < 0.01 vs the Aged–CM or oe‐PPT1–CM group).

   While previous studies have established a correlation between
   macrophage dysfunction and aging‐related myocardial fibrosis,
   additional cell‐specific interventions, such as adoptive macrophage
   transfer experiments, are required to establish a causal relationship.
   To validate the regulatory effects of IPA on PPT1 expression in
   macrophages, we performed adoptive macrophage‐transfer experiments.
   Clodronate liposomes (Lip‐Clo) were used to selectively deplete
   macrophages in young rats. Macrophages from Young, Aged, Aged + IPA,
   and Aged + sh‐PPT1 rats were then transferred into young rats to assess
   cardiac function (Figure [164]8A). Flow cytometry confirmed a
   significant reduction in macrophage levels in the Lip‐Clo group
   (Figure [165]8B,C). Echocardiography was performed to monitor cardiac
   function (Figure [166]8D), revealing that Aged + IPA or Aged + sh‐PPT1
   macrophage transfer improved cardiac function compared with Aged
   macrophage transfer alone (Figure [167]8E–L). H&E and Masson's staining
   showed increased inflammation and collagen deposition in the Aged to
   the Young group, whereas Aged + IPA or Aged + sh‐PPT1 mitigated these
   effects (Figure [168]8M,N,Q). Furthermore, the MCP‐1 and α‐SMA‐positive
   areas were increased in the Aged to Young group, but both IPA and
   sh‐PPT1 treatment reduced these markers (Figure [169]8O,P,R,S). The
   mRNA and protein levels of fibrosis and inflammatory markers
   corroborated these findings (Figure [170]8T–V).

Figure 8.

   Figure 8
   [171]Open in a new tab

   IPA inhibits PPT1 in elderly macrophages in vitro and reduces
   myocardial fibrosis in vivo. A) Schematic of the experimental design.
   B,C) Representative graphs and statistical graph of flow cytometry. D)
   Representative echocardiographic graphs. E–L) Echocardiographic
   measurements of LVEF, LVFS, LVIDd, LVIDs, LVPWd, LVPWs, IVSd, and IVSs.
   M) H&E staining of left ventricles. Magnification: 200×, scale bar = 50
   µm. N) Masson's staining of left ventricles. Magnification: 200×, scale
   bar = 50 µm. O,P) Representative images of IHC staining with MCP‐1 and
   α‐SMA antibody. Magnification: 200×, scale bar = 50 µm. Q) Statistical
   graphs of Masson's staining. R,S) Statistical graphs of IHC staining
   with MCP‐1 and α‐SMA antibody. T) Statistical analysis of COL3A1,
   COL1A1, IL‐6, and TNF‐α at mRNA level. U,V) Representative images and
   statistical analysis of COL3A1, COL1A1, IL‐6, and TNF‐α at protein
   level. (n = 3–6, data are expressed as mean ± SEM, ^* p < 0.05, ^** p <
   0.01 vs the Young group; ^# p < 0.05, ^## p < 0.01 vs the Aged group;
   ^& p < 0.05, ^&& p < 0.01 vs the Aged+sh‐PPT1 group).

2.6. IPA Regulates Macrophage Autophagy Function Through PPT1

   Macrophage autophagy decreases with age, leading to aggravated
   inflammation.^[ [172]^28 ^] Given the regulatory effect of IPA on
   autophagy and its impact on PPT1 expression in senescent macrophages,
   further investigation was warranted. To assess autophagic flux, BMDMs
   were infected with Ad‐GFP‐mCherry‐LC3B. Live‐cell imaging of BMDM
   showed spots with both GFP and mCherry fluorescence (i.e.,
   autophagosomes) in the LPS/IFN‐γ group and spots with only mCherry
   fluorescence in the LPS/IFN‐γ+IPA group (i.e., autolysosomes) (Figure
   [173]S9A, Supporting Information). Immunofluorescence analysis
   demonstrated that LC3B‐II formed distinct spots or clumps within the
   cell in both the control and LPS/IFN‐γ+IPA groups, indicative of normal
   autophagic activity (Figure [174]S9B, Supporting Information). LC3B‐II
   and Beclin1 protein expression was upregulated, whereas P62 protein
   expression was downregulated (Figure [175]S9C, Supporting Information).
   These data demonstrate that IPA activates autophagy in M1 macrophages.
   To determine whether IPA regulates macrophage autophagy via PPT1, we
   compared autophagic flux between the oe‐PPT1 group and oe‐PPT1 + IPA
   groups (Figure [176]S10A, Supporting Information). IPA treatment
   altered autophagic flow in the oe‐PPT1+IPA group compared to oe‐PPT1
   alone, as evidenced by the formation of LC3B‐II puncta (Figure
   [177]S10B, Supporting Information). Moreover, IPA reversed the
   oe‐PPT1‐induced downregulation of LC3B‐II and Beclin1 while suppressing
   the upregulation of P62, indicating that IPA counteracts PPT1‐mediated
   autophagy impairment.

   To investigate the mechanism by which IPA enhances lysosomal function
   and regulates PPT1 activation during aging, we performed RNA‐seq to
   screen differentially expressed genes. Comparative transcriptomic
   analysis revealed distinct gene expression profiles among Young‐BMDMs,
   Aged‐BMDMs, and Aged‐BMDMs treated with IPA (Figure [178]9A–C). It
   focused on the KEGG pathway involved in the PI3K‐AKT signaling pathway.
   To further explore the role of IPA in autophagy regulation, we assessed
   autophagic flux. Our results demonstrated that IPA treatment increased
   autophagy levels in aged macrophages (Figure [179]9D). Fluorescence
   imaging of LC3B revealed distinct punctate structures in Young and Aged
   + IPA macrophages, indicating the activation of autophagy
   (Figure [180]9E). Western blot analysis further confirmed that IPA
   modulated the expression of autophagy‐associated marker proteins,
   including LC3B‐II, Beclin1, and P62, by activating the PI3K‐Akt‐mTOR
   signaling pathway (Figure [181]9F,G).

Figure 9.

   Figure 9
   [182]Open in a new tab

   IPA regulates macrophage autophagy via PI3K‐AKT‐mTOR pathway. A) Heat
   map of differentially expressed genes. B) Venn diagram of
   differentially expressed genes. C) KEGG pathway enrichment bubble
   diagram of differentially expressed genes. D) mCherry‐GFP‐LC3B
   immunofluorescence staining representative images. Magnification: 200×,
   scale bar = 20 µm. E) Representative images immunofluorescence staining
   of LC3B. Magnification: 200×, scale bar = 20 µm. F,G) Representative
   images and statistical analysis of mTOR, p‐mTOR, PI3K, p‐PI3K, AKT,
   p‐AKT, P62, Beclin1 and LC3B at protein level. (n =4–6, data are
   expressed as mean ± SEM, ^** p < 0.01 vs the Young group; ^## p < 0.01
   vs the Aged group).

2.7. IPA Promotes PPT1 Expression via the cGAS‐STING Pathway

   As an important innate immune signaling pathway, the cGAS‐STING pathway
   is activated by nucleic acids. It is an important DNA sensor associated
   with the secretion of pro‐inflammatory factors and is linked to the
   senescence‐associated secretory phenotype (SASP).^[ [183]^29 , [184]^30
   ^] To verify the pathway through which IPA affects the PPT1 protein, we
   first assessed the impact of IPA on cGAS‐STING expression. Our results
   showed that IPA treatment reduced the mRNA and protein levels of cGAS
   and STING in cardiac tissue (Figure [185]10A–C). These findings were
   corroborated by in vitro protein analysis (Figure [186]10D,E). To
   further validate the role of the cGAS‐STING pathway in PPT1 regulation,
   we used specific inhibitors: RU.521 (a cGAS inhibitor) and C176 (a
   STING inhibitor). Inhibition of this pathway led to a significant
   reduction in PPT1 expression in M1 macrophages (Figure [187]10F), as
   confirmed by immunofluorescence and western blotting
   (Figure [188]10G–L).

Figure 10.

   Figure 10
   [189]Open in a new tab

   cGAS‐STING pathway is involved in IPA‐induced inhibition of PPT1. A)
   Statistical graph of cGAS and STING mRNA levels in macrophages. B,C)
   Representative images and statistical graph of cGAS and STING at
   protein levels in macrophages. D,E) Representative images and
   statistical graph of cGAS and STING at protein levels in vitro. F)
   Schematic diagram of IPA. G–J) Representative images and statistical
   graph of PPT1 immunofluorescence intensity in M1 macrophages after
   treated with RU.521 or C176. Magnification: 200×, scale bar = 20 µm.
   K,L) Representative images and statistical graph of PPT1 protein levels
   in M1 macrophages after treated with RU.521 or C176. M) Graphical
   abstract. (n = 3–6, data are expressed as mean ± SEM, ^* p < 0.05, ^**
   p < 0.01 vs the Young or Control group; ^# p < 0.05, ^## p < 0.01 vs
   the Aged or LPS/IFN‐γ group).

3. Discussion

   Numerous studies have demonstrated that gut microbiota has been
   associated with cardiovascular disease, with gut dysbiosis implicated
   in the development of myocardial fibrosis.^[ [190]^31 ^] Previous
   research highlights the involvement of gut microbiota in inflammatory
   conditions and emphasizing its broad impact on inflammatory
   processes.^[ [191]^32 , [192]^33 , [193]^34 ^] In our study, we found
   that plasma IPA levels were significantly lower in older patients with
   heart failure compared to age‐matched individuals without heart
   disease. Furthermore, our findings highlight the therapeutic potential
   of IPA, a product of tryptophan metabolism by c_Clostridium, in
   ameliorating cardiac function in aging. Previous studies have shown
   that pretreatment with IPA attenuates LPS‐induced cardiac dysfunction
   and inflammation.^[ [194]^35 ^] Wang et al. reported that IPA protects
   against heart failure by preserving ejection fraction.^[ [195]^8 ^]
   Aging induces a wide range of cardiovascular alterations, including
   extracellular matrix remodeling, increased oxidative stress, and
   dysregulation of immune responses.^[ [196]^36 ^] However, the role of
   IPA in aging‐related heart disease has not been extensively reported.
   To address this gap, we found the protective effect of IPA on cardiac
   function in aged rats, focusing on its ability to reduce inflammation
   and collagen deposition in aging‐related myocardial fibrosis. Our
   findings position IPA as a clinically relevant therapeutic drug for
   cardiovascular health.

   Furthermore, we investigated PPT1, a lysosomal enzyme that has been
   extensively studied in other contexts, including neurodegenerative
   diseases and systemic autoimmunity, but remains underexplored in the
   cardiovascular system.^[ [197]^18 , [198]^37 ^] Using proteomic
   sequencing, this study is the first to identify the key role of PPT1 in
   the heart. Our research demonstrates that PPT1 expression is
   upregulated in aged and inflamed cardiac tissue and is mainly regulated
   by macrophages. scRNA‐seq analysis demonstrated that PPT1 was
   significantly upregulated in aged macrophages, with PPT1 high
   expressively macrophages exhibiting a pro‐inflammatory phenotype. These
   findings suggest that PPT1 may be a potential therapeutic target for
   treating cardiac inflammation. Long‐term administration of D‐gal in
   rodents can mimic the overproduction of reactive oxygen species caused
   by physiological aging, a common way to induce aging in animals in
   vivo.^[ [199]^38 , [200]^39 ^] By verifying transgenic mice with PPT1
   knockout in macrophages, we found that knocking out PPT1 in macrophages
   could significantly reduce D‐gal‐induced aging‐related myocardial
   fibrosis and inflammation in mice. Pathological analysis further
   confirmed that, compared to the D‐gal‐treated group, the D‐gal +
   cko‐PPT1 group exhibited a significant reduction in cardiac
   inflammation and collagen deposition. These findings were corroborated
   by experiments in aged rats, where AAV‐sh‐PPT1 also led to a
   significant reduction in cardiac fibrosis and inflammation. Inhibition
   of PPT1 mitigated cardiac fibrosis and alleviated age‐related
   inflammation, highlighting its potential as a therapeutic target. Aged
   bone marrow macrophages could drive systemic aging and age‐related
   dysfunction.^[ [201]^40 ^] To determine whether IPA alleviates
   myocardial fibrosis through macrophages or whether knocking down PPT1
   in macrophages can alleviate myocardial fibrosis, we adoptively
   transferred aged BMDMs into young rats. Our findings revealed that aged
   macrophages with PPT1 knockdown and aged macrophages treated with IPA
   significantly reduced inflammation and fibrosis compared to untreated
   aged macrophages. Additionally, the same conclusion was obtained when
   the conditioned medium of macrophages was applied to cardiac
   fibroblasts in vitro. Specifically, COL3A1, COL1A1, and α‐SMA were
   upregulated in Aged or oe‐PPT1 conditional medium groups, whereas their
   expression was significantly down‐regulated in aged or oe‐PPT1
   macrophage that incubate with si‐PPT1, DC661, and IPA.

   Autophagy is essential for maintaining cardiovascular homeostasis and
   function. Aging is associated with impaired autophagy, leading to the
   accumulation of metabolic waste and contributing to various
   pathological conditions. Notably, autophagy acts as a double‐edged
   sword, both playing a role in and being influenced by cardiac aging.^[
   [202]^41 , [203]^42 , [204]^43 ^] Previous studies have shown that PPT1
   regulates autophagy in the context of tumor immunity and skeletal
   muscle differentiation.^[ [205]^44 , [206]^45 ^] These results suggest
   that IPA regulates macrophage autophagy via PPT1 in aging model. To
   further elucidate the signaling pathway through which IPA regulates
   autophagy in aging macrophages via PPT1, we performed RNA‐seq analysis.
   Autophagy presents a dual role in cellular physiology. While
   appropriate levels of autophagy enhance overall cellular function,
   excessive and insufficient autophagy may disrupt normal physiological
   processes and contribute to disease progression.^[ [207]^46 ^] The
   results showed that autophagic function in aged BMDMs was impaired,
   with significant inhibition of the PI3K‐AKT‐mTOR pathway. Notably, IPA
   treatment upregulated this pathway. To validate these findings, we
   conducted in vitro experiments to determine whether IPA could restore
   PI3K‐AKT‐mTOR pathway activity. Our experimental results demonstrated
   that incubation with IPA enhanced the autophagic function of
   pro‐inflammatory macrophages within LPS/IFN‐γ or oe‐PPT1 groups. In
   contrast, our in vivo data showed that IPA reversed aging‐induced
   autophagy inhibition in BMDMs, as evidenced by autophagy flux analysis
   and the expression of autophagy marker proteins. Growing evidence
   suggests that inhibition of the cGAS‐STING signaling pathway can
   significantly suppress cardiac hypertrophy and inflammatory cell
   infiltration.^[ [208]^47 ^] The cGAS‐STING pathway indirectly promotes
   aging, primarily through cellular senescence. Our results suggest that
   the intestinal metabolite IPA reduces cGAS and STING levels in the
   hearts of aged rats and M1 macrophages. RU.521 or C176 are commonly
   used inhibitors of cGAS and STING, respectively.^[ [209]^48 , [210]^49
   ^] When M1 macrophages were incubated with RU.521 and C176, PPT1
   expression decreased, supporting the conclusion that IPA regulates PPT1
   expression via the cGAS‐STING pathway. In conclusion, we demonstrated
   in vivo and in vitro that IPA alleviates aging‐related myocardial
   fibrosis through PPT1. Additionally, we found that inhibiting PPT1
   expression indirectly mitigates aging‐related myocardial fibrosis by
   modulating the autophagic function of macrophages (Figure [211]10M).

   This study primarily focused on the anti‐inflammatory and antifibrotic
   functions of IPA but did not investigate the gut microbiota responsible
   for its production. Future studies will employ fecal microbiota
   transplantation (FMT) experiments to identify specific gut microbiota
   influencing IPA levels in older individuals and determine the effect of
   IPA on the gut microbiota in the elderly. Because natural aging in mice
   takes ≈16 months, we utilized D‐gal to induce aging. However, we plan
   to conduct additional experiments using 16‐month‐old cko‐PPT1 mice for
   further validation. Furthermore, in addition to its role in autophagy,
   PPT1 functions as a depalmitoylation enzyme, regulating palmitoylation
   modifications in cardiac disease. We would examine its involvement in
   the palmitoylation of cardiac tissue of cko‐PPT1 mice in the future.
   Although the cGAS‐STING pathway plays a pivotal role in autophagy, our
   study only explored the regulatory relationship between IPA and PPT1.
   Future research should investigate exact role of this pathway in
   cardiac aging and myocardial fibrosis.

4. Experimental Section

Clinical Study Subjects

   Fourteen healthy young people, 30 healthy elderly people, and 30
   elderly patients with obvious symptoms, signs, and diagnosis of heart
   failure were recruited from the physical examination center who were
   hospitalized in the Department of Cardiology of the First Affiliated
   Hospital of Harbin Medical University. All patients underwent routine
   physical examination, laboratory tests (blood routine, blood
   biochemistry), electrocardiogram and cardiac color Doppler ultrasound
   within 24 h after admission. 3 mL peripheral venous blood was collected
   using EDTA anticoagulated blood collection tubes, and the plasma was
   separated. The upper layer of plasma was divided and stored at −80 °C.
   This study was approved by the Ethics Committee of the First Affiliated
   Hospital of Harbin Medical University.

Animals Models

   All the animal experiments in this study were conducted by the Guide
   for the Care and Use of Laboratory Animals and approved by the
   Institutional Animal Care and Use Committee of the First Affiliated
   Hospital of Harbin Medical University (IACUC No. 2023084). Male SD rats
   were purchased from Beijing VitalRiver Laboratory Animal Technology Co,
   Ltd (Beijing, China). Defined 4–6 months old rats were young group and
   18–20 months old as aged group. The PPT1 conditional KO mice were
   generated by employing the CRISPR/Cas9 technology to disrupt macrophage
   PPT1 and purchased from Cyagen Biosciences (Suzhou) Inc. Animals were
   raised at the Experimental Animal Center of the First Affiliated
   Hospital of Harbin Medical University. The animals were individually
   housed under 12:12‐h light‐dark cycles and were offered water and food
   ad libitum. At the end of the feeding regimen, the rats were
   anesthetized with 3% pentobarbital sodium and sacrificed by deep
   anesthesia of pentobarbital sodium.

Echocardiographic Measurement

   Ehocardiography was performed on each rat after transplantation using
   echocardiography analysis system. After moderate anesthesia of
   isoflurane gas in each group of mice, the heart rate was stable. Then,
   the mice used the short‐axis and long‐axis 2D ultrasound imaging of the
   heart of the ultrasound system, and analyzed it with the ultrasound
   imaging analysis system to obtain their heart structure and functional
   parameters.

In Vivo PPT1 Knockdown

   Knockdown of the PPT1 gene was constructed by Shanghai Genechem Co.,
   Ltd using the recombinant serum type 9 adeno‐associated virus system
   (AAV9) and the empty vector as the negative control. The
   GV576‐F4/80p‐MCS‐SV40 PolyA vector contains at least three shRNA
   fragments that were capable of simultaneously knocking down a gene at
   multiple sites. These sequences were as follows:
   5′‐CGGCATGGACGAGCTGTACAAGTAAGTCTCGAGGGATCCGCTAGC‐3′,
   5′‐TTATGATCTAGAGTCGCGGCCGCCAGCTATGGTGGCGGGGCCC‐3′, and
   5′‐CCCTGAGCAAAGACCCCAAC‐3′. The virus was injected into 18–20 months
   old SD rats via the tail vein for 8 weeks.

Tryptophan Metabolomics

   First, the experimental samples were pretreated, the metabolites were
   extracted, and the raw data were obtained by using the Thermo UltiMate
   3000RS and the mass spectrometer Thermo Q Exactive. The raw data were
   transformed into a matrix for further processing using specialized data
   processing software, including mass‐to‐charge ratio, retention time,
   and peak area information of metabolites. Data processing and
   statistical analysis were performed on the dataset to obtain
   differential metabolites and perform data visualization analysis.
   Tryptophan and its metabolites contents were then detected by MetWare
   Biotechnology Co., Ltd (Wuhan, China).

Bacteria 16S rDNA Sequencing

   Stool samples were collected from young and aged rats, snap‐frozen in
   liquid nitrogen, and stored at −80 °C. The bacterial 16S rDNA genes in
   the samples were sequenced by a high‐throughput detection platform. The
   bacterial genome database was used for data collation and analysis. The
   data were detected by MetWare Biotechnology Co., Ltd (Wuhan, China).

Proteomics Sequencing

   The isotope labeling quantification technique was used to combine
   multiple isotope tags with the N‐terminal groups of peptides from
   multiple groups of samples, and then tandem mass spectrometry was
   performed. The peak area of the reporter ion was used to calculate the
   ratio of the same peptide between different samples, so as to realize
   the quantitative comparison of different samples. The transcriptome
   sequencing and analysis were conducted by OE Biotech Co., Ltd.
   (Shanghai, China).

Single‐Cell and Single‐Nucleus RNA Sequencing Data Processing and Analysis

   Single‐cell transcriptomic data ([212]GSE183852) of normal and failing
   human hearts were downloaded from the Gene Expression Omnibus (GEO)
   database ([213]https://www.ncbi.nlm.nih.gov/geo/), to characterize
   age‐associated, macrophage‐specific transcriptional signatures, myeloid
   cells from healthy donor hearts were selected based on the cell
   annotations provided in the original publication for downstream
   analysis. The raw gene expression matrices were first converted into a
   Seurat object using the Seurat package (v4.3.0) in R. Batch effects
   were corrected using the Harmony (v1.2.0) algorithm. Cell clustering
   was conducted with the “FindNeighbors” and “FindClusters” functions in
   Seurat, using a resolution parameter of 0.5. For visualization, the
   “RunTSNE” function was used to generate 2D map based on the
   t‐distributed stochastic neighbor embedding (t‐SNE) algorithm. Clusters
   expressing marker genes from two or more distinct cell types were
   considered doublets and excluded from further analysis. Marker genes
   for each cell population were identified using the “FindAllMarkers”
   function with default parameters.

Transcriptome and Bioinformatics Analysis

   Total RNA was extracted using the TRIzol reagent (Invitrogen, CA, USA)
   according to the manufacturer's protocol. RNA purity and quantification
   were evaluated using the NanoDrop 2000 spectrophotometer (Thermo
   Scientific, USA). RNA integrity was assessed using the Agilent 2100
   Bioanalyzer (Agilent Technologies, Santa Clara, CA, USA). Then the
   libraries were constructed using VAHTS Universal V6 RNA‐seq Library
   Prep Kit according to the manufacturer's instructions. The
   transcriptome sequencing and analysis were conducted by Shanghai
   Applied Protein Technology Co., Ltd. (Shanghai, China).

ELISA of IPA

   The levels of the IPA in heart tissues were determined using commercial
   ELISA kits (Jiangsu Meimian Industrial Co., Ltd) according to the
   manufacturer's instructions.

Histological Analysis

   Histological analysis of was determined with H&E or Masson method. The
   heart tissues were fixed in 4% paraformaldehyde for 2 days, embedded in
   paraffin, and sliced into 4µm sections. H&E staining kit (Solarbio,
   Beijing, China) and Masson's trichrome stain kit (Solarbio, Beijing,
   China) were used to measure cardiomyocyte morphology and cardiac
   fibrosis. The myocardial sections were observed by microscope (Carl
   Zeiss Co., Ltd, Germany).

Immunohistochemistry Assays (IHC)

   Heart sections were deparaffinized in xylene series and rehydrated
   through ethanol series, followed by treatment of sections with 3%
   H[2]O[2] for 10 min. The sections were heated with antigen retrieval
   buffers to repair the antigen. Then they were blocked with 5% bovine
   serum albumin at 37 °C for 1 h. Sections were subjected to antigen
   retrieval in citrate buffer before adding CD68 primary antibody (Santa
   Cruz, sc‐20060, USA), MCP‐1 primary antibody (Affinity Biosciences,
   DF7577, China), TNF‐α primary antibody (Proteintech, 66291‐1‐Ig, USA),
   COL1A1 primary antibody (Proteintech, 66761‐1‐Ig, USA), and α‐SMA
   (Abcam, ab124964, Great Britain) overnight at 4 °C. The following day,
   the sections were incubated with horseradish peroxidase
   (HRP)‐conjugated anti‐mouse antibody or anti‐rabbit antibody and
   stained with diaminobenzidine (DAB). Images were taken with the
   microscope (Carl Zeiss Co. Ltd., Germany).

Primary Culture of Rat Bone Marrow Macrophages

   Rats were sacrificed and femur and tibia were separated under sterile
   conditions and then placed into cell culture dishes containing 75%
   alcohol. The bone ends of the femur and tibia were cut off, and then
   bone marrow cells were washed from one of the broken ends of the bone
   into 10 mL sterile centrifuge tubes using DMEM cell medium and repeated
   three times. At 1000 r min^−1, the samples were centrifuged for 8 min
   and the supernatant was discarded. 5 mL of erythrocyte lysate was
   added, pipetted repeatedly, and then allowed to rest for 3 min at 1000
   r min^−1, centrifuged for 8 min, and the supernatant was discarded.
   Cells were resuspended by adding 5 mL DMEM cell medium and filtered
   through nylon filters. At 1000 r min^−1, the samples were centrifuged
   for 8 min, and the supernatant was discarded and repeated three times.
   Cells were resuspended using a cell culture medium containing DMEM (10%
   FBS), resuspended in DMEM (10% FBS) containing 20 ng mL^−1 M‐CSF, and
   seeded into cell culture dishes. The cells were incubated at 37 °C in a
   5% CO[2] incubator using DMEM medium containing 10 % fetal bovine
   serum.

Cell Culture

   Primary rat cardiac fibroblasts were cultured as previously
   described.^[ [214]^50 ^] Cardiac fibroblasts (CFs) were acutely
   isolated from neonatal SD rats (1–3‐day‐old). CFs cells were cultured
   under standard conditions (37 °C, 5% CO[2]). The cells were divided
   into different groups and subjected to experimental procedures at 80 %
   confluence. BMDMs were isolated from Young or Aged rats, cells were
   treated with 1 µmol L^−1 IPA or 0.1 µmol L^−1 DC661 for 24 h.

Cell Transfection

   The pcDNA3.1‐PPT1 plasmid (oe‐PPT1) and pcDNA3.1‐negative control
   (Vector) were provided by Generalbiol (Chuzhou, China). Small
   interfering RNA of PPT1 (si‐PPT1) and negative control (NC) were
   provided by Seven (Beijing, China). Knockdown of the PPT1 gene was
   constructed by Shanghai Genechem Co., Ltd. by shRNA. When the growth
   density of BMDMs in six‐well plates reached 80% confluence,
   transfection was performed using jetPRIME® transfection reagent
   (Polyplus, France). After 6 h, the medium was replaced with fresh
   medium, IPA was added to continue the culture for 24 h, and the BMDMs
   were collected and tested.

Western Blot Analysis

   Cell or tissue samples were lysed in a lysis buffer with protease
   inhibitors. Subsequently, protein samples were separated by 10%
   SDS‐PAGE and transferred onto nitrocellulose membranes. The membrane
   was blocked with 3% no‐fat milk in PBST (PBS with 0.1% Tween 20) at 37
   °C for 1 h and incubated with the antibodies overnight at 4 °C, and
   then with HRP goat anti‐mouse IgG (H + L) or HRP goat anti‐rabbit IgG
   (H + L) secondary antibody for 1 h at 37 °C. Blot visualization was
   performed according to the manufacturer's instructions by using a Fully
   Automatic Chemiluminescence Image Analysis System. The antibody
   resources are as follows:

   ANP primary antibody (Proteintech, 27426‐1‐Ig, USA), β‐tubulin primary
   antibody (Proteintech, 10094‐1‐Ig, USA), STING primary antibody
   (Proteintech, 19851‐1‐Ig, USA), AKT primary antibody (Proteintech,
   60203‐2‐Ig, USA), PPT1 primary antibody (Proteintech, 29653‐1‐Ig, USA),
   COL1A1 primary antibody (Proteintech, 66761‐1‐Ig, USA), TNF‐α primary
   antibody (Proteintech, 60291‐1‐Ig, USA), cGAS primary antibody
   (ABclonal, A8335, China), PI3K primary antibody (Affinity Biosciences,
   AF6241, China), p‐PI3K primary antibody (CST, 4228S, USA), p‐AKT
   primary antibody (Affinity Biosciences, AF0016, China), MCP‐1 primary
   antibody (Affinity Biosciences, DF7577, China), IL‐6 primary antibody
   (Bioss, bs‐0782, China), p‐mTOR primary antibody (CST, 5536T, USA),
   mTOR primary antibody (CST, 2983T, USA), CD68 primary antibody (Santa
   Cruz, sc‐20060, USA), P16 primary antibody (Santa Cruz, sc‐1661, USA),
   CD86 primary antibody (Santa Cruz, sc‐28347, USA), α‐SMA primary
   antibody (Abcam, ab124964, Great Britain), LC3B primary antibody
   (Abcam, ab192890, Great Britain), P62 primary antibody (Abcam,
   ab109012, Great Britain), Beclin1 primary antibody (Abcam, ab19662,
   Great Britain), COL3A1 primary antibody (Abcam, ab184993, Great
   Britain), iNOS primary antibody (Abcam, ab178945, Great Britain), P21
   primary antibody (Novus‐Bio, NBP2‐29463, USA), GAPDH primary antibody
   (ZSGB‐Bio, TA‐08, China),

Real‐Time Quantitative Reverse Transcription PCR (RT‐qPCR)

   The samples were lysed using the RNAkey Reagent (Seven, Beijing,
   China), and the concentration and integrity of RNA samples were
   detected with the Nanodrop 2000 spectrophotometer (Thermo Scientific,
   USA). The volume of total RNA required for reverse transcription was
   calculated, and the first‐strand cDNA was synthesized according to the
   instructions of the FastKing RT Kit (With gDNase). The cDNA was
   synthesized with 1 µg of total RNA following a Reverse transcription
   kit (Yeasen, 11141ES60, China). RT‐qPCR was processed by the SYBR Green
   Mix (Yeasen, 11184ES08, China) and detected the mRNA expression by the
   SLAN‐96S system (SLAN, China). The data were analyzed using the 2^−ΔΔCT
   method to quantify relative gene expression. The primers are as
   follows:

   Rattus norvegicus:

   TNF‐α Forward 5′‐ATCGGTCCCAACAAGGAGGA‐3′,

   TNF‐α Reverse: 3′‐CGCTTGGTGGTTTGCTACG‐5′.

   PPT1 Forward 5′‐GTACTGGCATGACCCCATCA‐3′,

   PPT1 Reverse: 3′‐CAGAGTCTACAGGGTCCACG‐5′.

   GAPDH Forward 5′‐CAACTCCCTCAAGATTGTCAGCAA‐3′,

   GAPDH Reverse: 3′‐GGCATGGACTGTGGTCATGA‐5′.

   IL‐6 Forward 5′‐CCAGTTGCCTTCTTGGGACT‐3′,

   IL‐6 Reverse: 3′‐CTGGTCTGTTGTGGGTGGTA‐5′.

   COL1A1 Forward 5′‐GAGACAGGCGAACAAGGTGA‐3′,

   COL1A1 Reverse: 3′‐GGGAGACCGTTGAGTCCATC‐5′.

   α‐SMA Forward 5′‐ACCATCGGGAATGAACGCTT‐3′,

   α‐SMA Reverse: 3′‐CTGTCAGCAATGCCTGGGTA‐5′.

   P21 Forward 5′‐GCACTCTGAAGATGTGCCTATG‐3′,

   P21 Reverse: 3′‐CCCATAACTCCTTGCTAACTCC‐5′.

   COL3A1 Forward 5′‐CAGTCATGGGACTGGCATTTA‐3′,

   COL3A1 Reverse: 3′‐CAGTCATGGGACTGGCATTTA‐5′.

   cGAS Forward 5′‐GAAGCCCTGCTGTAACCCTT‐3′,

   cGAS Reverse: 3′‐ACAAGATAAAACGGCTCTCGTC‐5′.

   STING Forward 5′‐CCCAGCAAAACACAGACCG‐3′,

   STING Reverse: 3′‐AAACAAGGTCTGCAAGGGGGT‐5′.

   Mus musculus:

   PPT1 Forward 5′‐GTCTTTGGACTCCCCCGATG‐3′,

   PPT1 Reverse: 3′–CAGGCGTTCTTGCACAAGTT5′.

   GAPDH Forward 5′‐TGTGTCCGTCGTGGATCTGA‐3′,

   GAPDH Reverse: 3′‐TTGCTGTTGAAGTCGCAGGAG‐5′.

   IL‐6 Forward 5′‐TTCCTCTGGTCTTCTGGAGT‐3′,

   IL‐6 Reverse: 3′‐TCTGTGACTCCAGCTTATCTCTTG‐5′.

   COL1A1 Forward 5′‐AGACCTGTGTGTTCCCTACT‐3′,

   COL1A1 Reverse: 3′‐GAATCCATCGGTCATGCTCTC‐5′.

   COL3A1 Forward 5′‐ACGTAAGCACTGGTGGACAG‐3′,

   COL3A1 Reverse: 3′‐CAGGAGGGCCATAGCTGAAC‐5′.

Immunofluorescence Assay

   Wash samples two times with PBS. Fix samples with 4% paraformaldehyde
   in PBS for 15 min at room temperature. Permeabilize samples with 0.1%
   triton x‐100 in PBS for 10 min. Incubate samples in 10% normal goat
   serum in PBS for 30 min at room temperature. Samples were stained with
   primary antibody for 16 h. Then, the secondary antibody Alexa Fluor 488
   or 647‐labeled Goat Anti‐Rabbit or Mouse IgG(H+L), was added for 1 h.
   DAPI was added for 5 min. Immunofluorescence was detected by microscope
   (Carl Zeiss, Germany).

Statistical Analysis

   Statistical analyses were performed using GraphPad Prism 9.0 software
   (GraphPad Software, Inc., La Jolla, CA, USA). Continuous variables are
   presented as the mean ± standard error of the mean (SEM), while
   categorical variables are expressed as numbers and percentages.
   Two‐group comparisons were performed using the Student's unpaired
   t‐test. For comparisons among multiple groups, a one‐way analysis of
   variance (ANOVA) was used. Differences were considered statistically
   significant at ^* p < 0.05, ^** p < 0.01.

Conflict of Interest

   The authors declare no conflict of interest.

Author Contributions

   J.L., H.Y.W., and H.Y.Z. contributed equally to this work. J.L.,
   Z.X.D., Y.L., and Y.Q.S. designing research studies. J.L., H.Y.W.,
   J.H.L., and Q.Y.C. conducting experiments. H.Y.Z. and J.X. acquiring
   data. H.Y.W., H.Y.Z., J.H.L., and J.X. analyzing data. H.Q.L., D.Y.H.,
   J.J.L., L.L., K.Y.Y., X.P.W., S.Q.S., and F.K.L. providing reagents.
   J.L. and Z.X.D. writing the manuscript.

Supporting information

   Supporting Information
   [215]ADVS-12-e01070-s001.docx^ (11.4MB, docx)

Acknowledgements