Introduction
Myocardial infarction (MI) causes cell death and myocardial necrosis, resulting in irreversible infarct scarring and permanent loss of cardiac function, which are directly related to a worsened prognosis and reduced quality of life. Although reperfusion therapy during the hyperacute phase effectively prevents these outcomes, the overall prognosis post-MI remains poor. However, additional treatment options are limited.
Lymphatic vessels are present in the heart. Their physiological roles and pathogenetic relevance are gradually being elucidated, and are expected to represent potential therapeutic targets1,2. We focused on inducing lymphangiogenesis following MI, exploring the development of therapeutic lymphangiogenesis as a novel therapeutic approach for ischemic heart disease. For instance, research has shown that administering vascular endothelial growth factor (VEGF)-C promotes cardiac lymphangiogenesis, resulting in resolution of tissue edema and inflammation, reduction of infarct foci, and improved cardioprotection3. Furthermore, adrenomedullin (ADM) can induce reparative cardiac lymphangiogenesis and improve myocardial edema after MI4.
In recent years, regenerative medicine strategies have been developed to target MI. For instance, stem cell transplantation provides cardioprotection by replacing lost myocardial tissue after MI and exerting anti-apoptotic, anti-inflammatory, and/or angiogenic effects5, 6–7. Cardiac fibroblasts have also been implicated in repair mechanisms following MI8. Cardiac fibroblasts have multiple sub-clusters, representing heterogeneous cell populations with distinct properties8. We have demonstrated that transplantation of a specific population of human fetal cardiac fibroblasts (fCFs), expressing vascular cell adhesion molecule 1 (VCAM1), induces cardiac lymphangiogenesis after MI to reduce cardiac infarct size9. However, immunogenicity and ethical issues remain regarding the use of fCFs in allogeneic transplantation. In addition, it is unclear whether aCFs, which can be used as autologous transplants in clinical practice, possess lymphangiogenesis-promoting properties similar to those of fCFs.
In this study, we identified a specific aCF population that could promote lymphangiogenesis and explored efficient induction and sorting methods. We also tested whether this cell population could be applied to the treatment of MI.
Results
Heterogeneity of human aCFs
To assess the heterogeneity of human aCFs, we first cultured primary fibroblasts derived from human heart ventricles. Primary human aCFs exhibited a flat, spindle-shaped morphology typical of fibroblasts (Fig. 1a). aCFs barely expressed cardiomyocyte or endothelial cell main markers, including cardiac troponin T (TNNT2, cTnT), α-actinin, and VE-cadherin (CD144), ruling out contamination by cardiomyocytes or endothelial cells in this population (Fig. 1b). Conversely, aCFs were strongly positive for numerous fibroblast- or myofibroblast-specific markers, including vimentin, alpha-smooth muscle actin (αSMA), discoidin domain receptor tyrosine kinase 2 (DDR2), and fibronectin. However, unlike these markers, CD90 (THY1) expression pattern showed a clear bimodal distribution (Fig. 1b), in which the polarity was quite different from that of fCFs, where the majority of cells were positive for CD90, as previously reported9.
Fig. 1 [Images not available. See PDF.]
Characterization of CFs. (a) Micrographs of plated aCF (scale bars: 500 μm (5X) and 100 μm (20X)). (b) Cell profiling showing the expression of several markers of cardiomyocytes (i.e., cTnT and α-actinin), endothelial cells (i.e., CD144) and fibroblasts (i.e., Vimentin, αSMA, DDR2, fibronectin, and CD90). (c) CD90 expression distinguishing two subpopulations of fibroblasts. (d) Increase in VCAM1 + relative cell population by TNF-α (mean ± SD, one-way ANOVA with Tukey–Kramer post-hoc analysis, P < 0.01 (**) vs. 0 ng/mL of TNF-α (Ctrl), P < 0.05 (*) versus Ctrl). (e) Effect of TNF-α and IL-4 (ng/mL) on the distribution of VCAM1 expression in aCF (mean ± SD, one-way ANOVA with Tukey–Kramer post-hoc analysis, P < 0.01 (**) vs. 0 ng/mL of TNF-α and IL-4). (f) Cell characteristics of each subpopulation in aCF treated with CKs (aCF + CKs). The proportion of each CD90/VCAM-1 subpopulation was as follows: CD90(−)/VCAM-1(−); 21.59%, CD90(−)/VCAM-1(+); 68.60%, CD90(+)/VCAM-1(−); 0.69%, and CD90(+)/VCAM-1(+): 9.12%. (g) Micrographs of cultures of aCF, aCF + CKs, or aCF treated with TGFβ1 (aCF + TGFβ) (αSMA (green), βActin (red), and Hoechst 33,258 (blue); scale bars: 100 μm). (h) RT-qPCR analysis of ECM rigidity gene expression in aCF, aCF + CKs, or aCF + TGFβ. i, Localization changes of GFP-labeled NF-kB and STAT6 in aCF upon addition of CKs. (j) Luciferase reporter assays. (k) Results of proliferation assay (WST-1 assay) using Conditioned Medium (CM) of aCF, aCF + CKs and aVCF. Control indicates DMEM (no-serum) medium. (Mean ± SD, one-way analysis of variance with Tukey’s multiple comparisons test, P < 0.0001 (****), P < 0.001 (***), P < 0.1 (**)). (l) Results of migration assay when culture supernatants of aCF, aCF + CKs and aVCF were applied to HDLECs.Results of DAPI staining of nuclei of migrated HDLECs (imaged at 10 × magnification). (m) Results of counting and quantifying the number of DAPIs using microscopic images obtained from migration assay.Control indicates DMEM (no-serum) medium. (Mean ± SD, one-way analysis of variance with Tukey’s multiple comparisons test, P < 0.0001 (****), P < 0.001 (***), P < 0.5 (*)).
We have previously demonstrated that the relative expression of VCAM1 defines two distinct subpopulations of fCFs9. Moreover, VCAM1-expressing fCFs play a role in cardiac lymphangiogenesis, and the injection of CD90+ VCAM1+ fCFs restores cardiac contractility during subacute heart failure following MI in rats by mobilizing LECs into the infarcted region. Therefore, we evaluated the expression of CD90+ VCAM1+ in aCFs and found that this population was almost absent (1.1 ± 0.77%, n = 3) (Fig. 1c), in contrast with the observations in fCFs9.
Mimicking cellular characteristics of aCFs to fCFs
A previous study reported that TNF-α induced VCAM1 upregulation in aCFs by activating the nuclear factor-kappa B (NF-κB) pathway, with these VCAM1-upregulated myofibroblasts enhancing monocyte adhesion10. Thus, we cultivated aCFs with various concentrations of TNF-α and observed that VCAM1 expression was induced by TNF-α at concentrations ≥ 1.0 ng/mL (Fig. 1d). However, only 2% of aCFs expressed VCAM1 after induction by TNF-α. Therefore, we investigated another VCAM1 expression driver: IL-4.
IL-4 induces STAT6 binding to the VCAM1 promoter in endothelial cells11. It has also been reported that co-expression of NF-κB and IL-4-induced STAT6 significantly enhances the expression of IL-4-inducible reporter genes12. Based on this, we treated aCFs with various concentrations of TNF-α and IL-4 (cytokines; CKs) in combination and evaluated the proportion of VCAM1-positive cells by flow cytometry. Compared to untreated aCFs (“aCFs”), the percentage of VCAM1-positive cells increased significantly in an IL-4 dose-dependent manner in the presence of TNF-α. Importantly, IL-4 alone induced only a very low proportion of VCAM1+ cells, as shown in Supplementary Fig. 1,while no significant differences were observed among TNF-α concentrations ranging from 1 to 50 ng/mL, the highest proportion of VCAM1+ cells were detected with the combination of 50.0 ng/mL TNF-α and 2.0 ng/mL IL-4 (Fig. 1e). Furthermore, treatment with this combination of CKs increased the proportion of VCAM1-positive cells across all fibroblast populations, including both CD90⁻ and CD90+ subsets, thereby promoting the emergence of CD90+ VCAM1+ activated cardiac fibroblasts (aVCFs) ( Fig. 1f). In addition, as shown in Supplementary Fig. 1, IL-4 alone failed to significantly induce VCAM1 expression, with the proportion of VCAM1+ cells remaining below 1% across all tested concentrations. These findings suggest that IL-4 and TNF-α must be used in combination to effectively induce the VCAM1+ phenotype in cardiac fibroblasts.” In subsequent experiments, the CD90+ VCAM1+ cell fraction of aCF + CKs isolated by cell sorting was referred to as aVCFs. In other words, whereas the aCF + CKs group includes a heterogeneous mixture of fibroblasts—only a subset of which are CD90+ VCAM1+—the aVCFs population represents a purified fraction of these activated fibroblasts. Additionally, the ECM rigidity, protein, and gene expression patterns of aVCF differed from those of myofibroblasts induced by TGF-β1 (Fig. 1g and h). Notably, although FSP1 expression was higher in aCFs treated with TNF-α and IL-4 than in TGF-β-induced myofibroblasts, other classical myofibroblast markers—such as POSTN, FAP, and ACTA2—were more strongly upregulated by TGF-β1. These findings suggest that TNF-α/IL-4-treated aCFs constitute a functionally distinct fibroblast subtype, characterized by enhanced inflammatory responsiveness and potentially greater lymphangiogenic capacity, rather than a classical myofibroblast phenotype.
Using a lentiviral vector to generate GFP-labeled human aCF cell lines for NF-κB and STAT6, we evaluated the signals enhanced by CKs and found that CKs promoted nuclear translocation of both NF-kB and STAT6 (Fig. 1i). Similar results were obtained in a luciferase reporter assay (Fig. 1j), suggesting that CKs induced a differentiation pathway different from that of myofibroblasts.
To assess whether aCFs have the potential to improve lymphangiogenesis, as observed with CD90+ VCAM1+ fCFs (fVCFs)9, we performed lymphatic endothelial proliferation and migration assays (Fig. 1k–m). The results showed that aVCF produced the highest lymphatic endothelial proliferative and migratory effects (P < 0.001 vs. the control).
Lymphangiogenic potential of fCF-like aCFs and their ability to restore cardiac function
To evaluate whether aVCF could improve cardiac function after heart failure by inducing restorative lymphangiogenesis, we administered the following types of fibroblasts to rat models of ischemic heart failure: aCFs, aCFs + CKs, and aVCFs. The proportions of VCAM1+ and CD90+ cells (%) in each population were confirmed using flow cytometry (see Supplementary Table 1).
A schematic diagram of the experimental procedure is shown in Fig. 2a. M-mode echocardiography of rats in the aCF-treated group showed no therapeutic effect on LVEF (44.1 ± 2.8%) or LVFS (17.8 ± 1.4%) at 18 weeks post-transplant (Fig. 2b). Conversely, rats treated with aCFs + CKs showed improvements in LVEF and LVFS up to 10 weeks after transplantation (Fig. 2b). However, these values decreased to 49.6 ± 4.0% and 20.9 ± 2.1%, respectively, by the end of the experiment (Fig. 2b). In contrast, the aVCF-treated group showed continuous improvement in LVEF and LVFS throughout the entire duration of the experiment, with LVEF reaching 61.0 ± 2.2% and LVFS 27.2 ± 1.4% at 18 weeks post-transplant (Fig. 2b).
Fig. 2 [Images not available. See PDF.]
Animal experiments. (a) Schedule of experiments with the administration of each fibroblast to the rat heart. (b) Effects of lymphangiogenesis by administration of each fibroblast; Results of LVEF and LVES up to 18 weeks. Statistical analysis was performed using the Kruskal–Wallis test followed by Dunn’s multiple comparisons test. Statistical significance was indicated in the graph when *P < 0.05 for comparisons between Vehicle Cntl (or aCF) and aVCF, and + P< 0.05 for comparisons between Vehicle Cntl (or aCF) and aCF + CKs. c, Graph showing variation in LVEF and LVFS. Statistical analysis was performed using the Kruskal–Wallis test followed by Dunn’s multiple comparisons test. Statistical significance was indicated in the graph when *P < 0.05 for comparisons between Vehicle Cntl (or aCF) and aVCF, and + P < 0.05 for comparisons between Vehicle Cntl (or aCF) and aCF + CKs. (d) Histological evaluation of the heart (evaluation of fibrosis by Sirius Red and visualisation of the infarct area by cTnT staining. For the analysis of the fibrosis area, equal variance was confirmed using the Shapiro–Wilk normality test. Therefore, one-way ANOVA with Tukey’s multiple comparisons test was applied (**P < 0.001, *P < 0.01). For the infarcted area, equal variance was not assumed based on the Shapiro–Wilk test, and statistical analysis was conducted using the Kruskal–Wallis test followed by Dunn’s multiple comparisons test (*P < 0.01). (e) Immunofluorescence staining of cTnT (green), Prox-1 (red), and DAPI (blue). Magnification images were captured at × 20. (f) The fluorescence intensity of Prox-1 (red) was quantified using ImageJ. (Mean ± SD, one-way analysis of variance with Tukey’s multiple comparisons test, P < 0.001 (***), P < 0.0001 (****)).
We also measured the variation in LVEF and LVFS (i.e., Delta LVEF and Delta LVFS) in each experimental group. The aCF-treated animals showed a decrease in LVEF of − 6.9 ± 2.6% and LVFS of − 3.5 ± 1.3% by the end of the monitoring period (Fig. 2c). Additionally, the aCFs + CKs-treated animals showed a decrease in LVEF of − 1.8 ± 3.4% and LVFS of − 0.6 ± 1.8% (Fig. 2c). In contrast, animals in the aVCF-treated group exhibited an increase in LVEF of 9.2 ± 2.6% and an increase in LVFS of 5.6 ± 1.6% at 18 weeks post-transplant (Fig. 2c). The other values and statistical analyses for all echocardiographic measurements are presented in Supplementary Table 2. These results suggest that factors uniquely or predominantly expressed in aVCF, but not in aCF + CKs, may play a key role in the long-term improvement of cardiac function.
To identify the mechanisms underlying the effects of aVCF, we first assessed myocardial tissue remodeling triggered by each type of fibroblast. To characterize infarction and fibrosis in each group, we performed SR and cTnT staining. Consistent with the H&E staining, these results showed that aVCF-treated animals had significantly less fibrosis and infarct areas than animals from the other groups (Fig. 2d).
To elucidate the cardiac lymphangiogenic effects of various cardiac fibroblasts, we assessed the localization of Prox-1-positive cells. Immunofluorescence staining of cTnT, Prox-1, and DAPI revealed that the number of Prox-1+ nuclei was significantly increased in the aVCF group compared to the vehicle control, aCF, and aCF + CKs-treated groups (Fig. 2e and f). As Prox-1 is also expressed in cardiomyocytes, the observed increase may reflect both lymphatic and non-lymphatic cell populations. Furthermore, in situ hybridization (ISH) revealed that VEGFR3-positive cells were most abundant in the aVCF-treated group, followed by the aCF + CKs-treated group, the aCF-treated group, and the Vehicle Control group. Notably, the number of VEGFR3+ cells was significantly higher in the aVCF-treated group compared to the Vehicle Control group (Supplementary Fig. 2). Given that VEGFR-3 is also highly expressed in cardiac macrophages, the observed increase may reflect both lymphatic endothelial cells and macrophages.
These results suggest that factors specifically or more dominantly expressed in aVCFs—with a phenotype similar to VCAM1+ fCFs compared to aCFs treated with CKs—may contribute to the long-term improvement of myocardial contractile function and reduction of fibrotic areas after myocardial infarction, at least in part through the promotion and maintenance of lymphangiogenesis and the attenuation of local inflammation.
Identification of paracrine factors involved in the lymphangiogenic effects of aVCFs
To clarify the role of aVCFs in long-term lymphatic vessel regeneration or maintenance, fibrosis prevention, and cardiac function recovery—as suggested by the findings in Fig. 2, which used a myocardial infarction (MI) model—we performed a comparative gene expression analysis using RNA-seq on human cardiac fibroblast populations. Specifically, stringent cut-offs were applied to redefine the DEG lists at first. Only genes with a fold change greater than or equal to 5 (highly upregulated) or less than or equal to 1/5 (highly downregulated) in various comparisons were considered for further analysis. All selected genes had a false discovery rate of < 0.01. The restricted DEG lists are as follows: aCF + CKs vs. aVCF: 161 DEGs; aCF vs. aCF + CKs: 283 DEGs; aCF vs. aVCF: 560 DEGs.
Additionally, PCA was performed on the complete DEG lists, and the 150 most variant genes were selected for further analysis (Fig. 3a). Genes encoding for secreted proteins among this selection were identified by comparison with the corresponding gene set available on UniProt (filter: “Secreted,” “Reviewed,” “Human”; 2088 genes). Among the 150 most variant genes, 64 were annotated as encoding secreted proteins.
Fig. 3 [Images not available. See PDF.]
Identification of paracrine factors involved in the lymphangiogenic effect of aVCF using RNA-seq. (a) As a result of extracting differentially expressed genes (DEG) and principal component analysis (PCA), 150 genes were identified. (b) Only the cluster of upregulated genes is shown in a Venn diagram. (c) The distribution of gene expression in the three types of aCF was shown in a heat map, and short-term treatment factors (STF) and long-term treatment factors (LTF) were identified.
In vivo experiments in rats suggested dual activity of the injected cells, resulting in both short- and long-term therapeutic effects. While aCFs + CKs and aVCFs contributed to the restoration of heart function over a short period (< 12 weeks after cell administration), only aVCFs maintained this effect until the end of the monitoring period (18 weeks after cell administration), as previously reported9. Moreover, lymphangiogenesis was confirmed only in the animals treated with aVCF. This experimental observation led to the establishment of two profiles of candidate gene expression: short-term effect-mediating factors (i.e., expression similar in aCFs + CKs and aVCFs compared to aCFs) and long-term effect-mediating factors (i.e., expression specific to aVCFs).
The comparison of DEG lists (highly up- or downregulated) allowed us to describe the distribution of gene expression among the three types of CFs. The comparison with the list of genes encoding secreted proteins identified 32 genes as short-term treatment factors (STF) and 20 genes as long-term treatment factors (LTF) (Fig. 3b).
Expression patterns within these two sub-lists were visualized using a heatmap. Since no downregulated genes had meaningful annotations in this context, only clusters of upregulated genes were selected for further analysis (Fig. 3c). Finally, the candidate genes were extracted as follows: STF: CXCL1, IL32, CXCL8, CXCL3, IL6, HBEGF, IL1RL1, CCL20, CXCL2, CSF3 (G-CSF), IL7R, EBI3, CXCL5, CSF2 (GM-CSF), OTOGL, IL1B, MMP1, IL1A, CXCL6, C1QTNF1, CCL2, ADM, CBLN2, and COL6A6; LTF: EREG, SERPINA9, LAMB3, SERPINB2, IL1B, MMP1, and IL1A.
Next, the protein production levels of all candidate genes were evaluated among aCFs, aCFs + CKs, and aVCFs using ELISA and multiplex analysis (Supplementary Fig. 3). As a result, ADM, which was selected as a candidate factor for the direct promotion of lymphangiogenesis, was expressed approximately eight times more in aVCFs than in aCFs (Fig. 4a)4. Additionally, as shown in Fig. 1i and j, aVCFs upregulate VCAM1 expression through NF-κB nuclear translocation and STAT6 activation. Notably, we confirmed that ADM expression is dependent on the NF-κB/STAT6 signaling axis, as shown in Fig. 4b. To confirm the effect of ADM in aVCFs, we generated aVCFs with ADM gene knockdown using siRNA transfection (si-ADM). The knockdown efficiency of ADM by siRNA transfection is shown in Supplemental Fig. 4. The aVCF cells that had not undergone ADM knockdown were transfected with a negative control siRNA (si-negative control). When these cells were directly co-cultured with HDLECs, tube formation was significantly disrupted in the si-ADM group (Fig. 4c). Quantitative analysis of the branch length in the microscopic images acquired using ImageJ revealed a significant decrease in tube formation in the si-ADM group (Fig. 4d). In addition, when LECs were stimulated with the supernatant from a conditioned medium of cultured aVCFs or aVCFs with ADM gene knockdown, the proliferative ability of LECs was partially inhibited in the si-ADM group (Fig. 4e). Moreover, the migration ability of LECs was attenuated in the si-ADM group (Fig. 4f and g).
Fig. 4 [Images not available. See PDF.]
The effect of ADM on aVCF. (a) Adrenomedullin, a lymphangiogenic paracrine factor that shows significantly high production levels in aVCF(Mean ± SD, one-way analysis of variance with Tukey’s multiple comparisons test, P < 0.001 (***), P < 0.01 (**)). (b) The expression of ADM is regulated via the NF-κB and STAT6 signaling pathways. Statistical analysis was performed using an unpaired t-test. ***P < 0.001. (c) Tube formation when ADM was transfected with siRNA. d, The results of quantitative analysis of the results of tube formation. Statistical analysis was performed using an unpaired t-test. ***P < 0.001. (e) Results of proliferation assay when ADM was transfected with siRNA. Statistical analysis was performed using an unpaired t-test. *P < 0.05. (f) Results of migration assay when culture supernatants of mVCF si-negative control and mVCF si-ADM were applied to HDLECs (imaged at 20 × magnification). Results of DAPI staining of nuclei of migrated HDLECs. g, The results of quantitative analysis of the results of migration. Statistical analysis was performed using an unpaired t-test. *P < 0.05.
Finally, we tested whether the cardioprotective effects of aVCF against ischemia–reperfusion (IR) injury occurred, at least in part, through an ADM-dependent mechanism. In this study, our primary objective was to establish proof-of-concept (PoC) regarding the adrenomedullin-dependent beneficial effects of aVCFs in vivo. To accurately assess the intrinsic effect of aVCFs without the confounding factors of xenogeneic immune responses or the need for immunosuppression, we employed a syngeneic mouse model. To support this, the morphology of mCFs and the expression of CD90.2 (the mouse homolog of human CD90) and VCAM-1 were shown in Supplementary Fig. 5a and b. In addition, the expression levels of ADM in mCFs and mVCFs were quantified by ELISA, and the results are presented in Supplementary Fig. 5c. These findings suggest that similar to aVCFs, mVCFs are also capable of promoting lymphangiogenesis. VCFs (mVCF) established in mouse-derived cells, ADM-knockdown mVCFs (mVCF siADM), or vehicle control was administered intramyocardially at the same time as myocardial ischemia was released for 60 min, followed by the evaluation of subsequent changes in cardiac function and heart pathology (Fig. 5a). Figure 5b shows that the protective effect of mVCF against cardiac systolic dysfunction after IR injury was partially abolished by ADM knockdown. In addition, pathological validation of hearts harvested after four weeks of reperfusion showed that myocardial fibrosis induced by IR injury was suppressed by mVCF administration. Importantly, ADM knockdown partially abolished these suppressive effects (Fig. 5c and d). Cardiac lymphatic vessel formation induced by mVCF administration was attenuated by ADM knockdown (Fig. 5e and f). Furthermore, quantification of Lyve-1 and podoplanin (PDPN) double-positive cells in the same cardiac tissue sections revealed a significant increase in the number of double-positive cells in the mVCF-administered group, indicating enhanced lymphangiogenesis. Notably, this increase was significantly attenuated in the mVCF siADM group, supporting a critical role of ADM in mediating the pro-lymphangiogenic effect of mVCFs (Supplementary Fig. 6).
Fig. 5 [Images not available. See PDF.]
In vivo studies of confirmation to the therapeutic efficacy of ADM. (a) Schedule of experiments with the administration of each fibroblast to the mouse heart. (b) Effects of lymphangiogenesis by administration of each fibroblast; Results of EF and up to 4 weeks. Statistical comparisons were made between groups at each time point using two-way ANOVA with Tukey’s multiple comparisons test. ****P < 0.0001, ***P < 0.001: mVCF-ADM-KD vs. Vehicle Ctrl, + + P < 0.01, + P < 0.05: mVCF-ADM-KD vs. mVCF-NC. (c) Fibrosis of the heart after 4 weeks of cell administration. (d) Quantitative evaluation of fibrosis in the mouse heart. Statistical analysis was performed using one-way ANOVA with Tukey’s multiple comparisons test. ****P < 0.0001, ***P < 0.001, *P < 0.05. (e) Localisation of lymphangiogenic factors assessed by fluorescence staining of mouse heart sections (Magnification: × 20). Statistical analysis was performed using one-way ANOVA with Tukey’s multiple comparisons test. ****P < 0.0001, ***P < 0.001, *P < 0.05. (f) Quantitative evaluation of lymphangiogenic factors (Lyve-1 and VEGFR3) by fluorescence staining of mouse heart sections.
Furthermore, to investigate whether the enhanced lymphangiogenesis induced by VCFs is associated with an anti-inflammatory effect, we performed additional qPCR analyses using mouse heart tissue to evaluate the expression of inflammatory cytokines, TNF-α and IL-1β. VCF-treated hearts showed a marked reduction in the expression of both cytokines compared to controls (Supplementary Fig. 7a).
In addition, immunofluorescence staining of heart tissue sections revealed a significant decrease in the number of CD68+ and F4/80+ macrophages in the VCF-treated group. This reduction in macrophage infiltration was partially reversed in the mVCF siADM group, indicating that the anti-inflammatory effect of VCFs is at least partially dependent on ADM from VCFs (Supplementary Fig. 7b).
Discussion
Compared with fCFs, aCFs exhibited a different distribution of cell surface protein markers (Fig. 1a–c)9 where aCFs expressed low levels of VCAM1. Therefore, we focused on TNF-α and IL-4, which were reported to induce VCAM1 upregulation via the nuclear translocation of NF-κB and STAT6 expression9, 10–11. The addition of TNF-α and IL-4 as a cocktail to the culture medium significantly and efficiently upregulated VCAM1 expression on aCFs (Fig. 1d and e). Additionally, the differentiation tendency differed from that of myofibroblasts induced by transforming growth factor beta 1 (TGF-β1) in terms of ECM rigidity (Fig. 1f)13.
TGF-β1 is a major inflammatory cytokine secreted by mobilized neutrophils after cardiac injury, which activates the TGF-β/Smad pathway in cardiac fibroblasts14. Smad proteins translocate to the nucleus, upregulating the expression of ECM proteins and αSMA, thereby changing the contractility of microfilaments in cardiac fibroblasts15,16. These activated myofibroblasts eventually proliferate and accumulate ECM in the infarcted area, forming stiff fibrotic tissue17. Our results suggest that the upregulation of VCAM1 by CKs is dependent on the NF-κB/STAT6 pathway, as reported previously 9, 10–11, and is functionally different from the fibrogenic response mediated by the TGF-β/Smad pathway (Fig. 1g–j). Although not directly examined in this study, it is possible that TGF-β1-treated myofibroblasts do not rely on the NF-κB/STAT6 pathway. This remains a limitation of the present work and will be addressed in future studies. Furthermore, it will be important to assess whether the profile, including FSP1, which differs from that induced by CKs, becomes more similar to that of fCFs.
aVCFs, similar to previously reported fCFs9, exhibited a restorative effect on cardiac function in a subacute heart failure rat model (Fig. 2a–c). Pathological evaluations suggested that fibrosis was related to the effect of aVCFs on the restoration of cardiac function (Fig. 2d). Cardiac fibrosis, caused by collagen deposition by activated cardiac fibroblasts, is derived from myocardial edema and inflammatory mediators released by infiltrated immune cells13, 14, 15–16,18,19. Cardiac lymphangiogenesis has been reported to be effective not only in recovering myocardial edema, inflammation, and cardiac function caused by impaired cardiac lymphatic flow but also in treating already-formed fibrotic areas as an outcome of flow disturbance18. We postulated that the previously documented effects of diverse fibroblast subtypes on cardiac function and fibrosis may be attributable to variations in their lymphangiogenic capacity. To investigate this, we assessed the spatial distribution of lymphatic endothelial cells within the infarcted region using histological staining techniques.
In the heart tissues of heart failure model rats, lymphatic endothelial cells were more abundant in the infarct region of the aVCF-treated group (Fig. 2e and Supplementary Fig. 2), resembling the previously reported effects of fVCFs. These findings suggest that aVCFs have acquired fVCF-like properties9, particularly in promoting lymphatic vessel regeneration. This suggests that aVCFs acquire fVCF-like characteristics in terms of lymphangiogenesis function.
Considering the differences in the influence of each fibroblast type on the recovery of cardiac function, we narrowed down the list of candidate factors involved in cardiac lymphangiogenesis to ADM (Figs. 3 and 4a). ADM is a cardioprotective peptide that significantly improves the cardiovascular function in patients with acute MI20,21. In addition, it has been reported to promote cardiac lymphangiogenesis and regulate cardiac lymphatic connexin 43, thus improving cardiac function after MI4.
Surprisingly, our ELISA analysis revealed that the ADM production by fCFs was markedly lower than that of aVCFs, contrary to our initial expectation (Supplementary Fig. 8a). To further investigate the differences in their paracrine profiles, we also quantified the mRNA expression levels of ADM, VEGF-C, and HGF (Supplementary Fig. 8b). Consistent with the ELISA findings, ADM mRNA expression was significantly higher in aVCFs. In contrast, VEGF-C expression was highest in fCFs, while HGF expression was comparably elevated in both fCFs and aVCFs, relative to aCFs.
These findings suggest that although aVCFs and fCFs differ in ADM production, they may share certain regenerative or pro-angiogenic characteristics through other paracrine factors, such as VEGF-C and HGF, which may contribute indirectly to therapeutic lymphangiogenesis and tissue repair.
This study has a few limitations as well. Although bioinformatics analyses and subsequent experiments suggest that ADM is a key effector molecule of aVCFs, it is also significantly upregulated in aCF + CKs compared to aCFs. The observed difference in cardiac functional recovery between aCF + CKs and aVCFs in MI models may be due to a threshold effect of ADM expression or the involvement of additional factors, which remain to be elucidated in future studies.
Furthermore, through bioinformatics analysis, ELISA, and multiplex assays, we identified several proteins that were produced at significantly higher levels in aVCFs compared to aCFs or aCF + CKs (Supplementary Fig. 3). These included inflammatory CKs and chemokines, CD8+ T cell-inhibiting factors, and apoptosis-inhibiting CKs, in addition to ADM, which are directly and indirectly involved in lymphangiogenesis. Several inflammatory CKs and chemokines have been reported to promote inflammation-induced lymphangiogenesis, such as through VEGF-C production by lymphatic endothelial cells22, 23, 24–25.
For example, G-CSF is widely known as a factor that induces macrophages to differentiate into M2-type macrophages26, 27–28. M2-type macrophages produce VEGF-C and act as sentinel cells for lymphatic vessels29,30. Additionally, VEGF-C promotes further polarization and transdifferentiation of M1 macrophages into lymphatic endothelial cells31, and GM-CSF induces the differentiation of M1 macrophages by increasing the expression of C–C chemokine receptor type 2 (CCR2), which is activated by its ligand, C–C motif chemokine ligand 2 (CCL2).32.
In addition, laminin subunit beta-3 (LAMB3), a gene upregulated in pancreatic ductal adenocarcinoma, promotes tumor invasion via activating the phosphatidylinositol 3-kinase (PI3K)/Akt axis33, 34–35. This signaling pathway is involved in regulating the proliferative capacity of cancer cells and their anti-apoptotic, metastatic, and invasive potentials36, 37–38. Similarly, C1q/TNF-related protein 1 has been reported to protect against acute myocardial injury after IR and to inhibit myocyte apoptosis and inflammatory responses by promoting cyclic AMP-dependent pathways39. It may have similar effects on cardiomyocytes in the infarct border zone and contribute to the repair and maintenance of structural defects in the myocardium at the infarct site.
Additionally, VCAM1 is a plasma membrane marker of aVCFs, identified outside of the factors analyzed by ELISA and multiplex analysis. However, we previously discovered that mouse cardiac fibroblast-derived VCAM1 protein can stimulate the proliferation of mouse cardiomyocytes derived from embryonic stem cells when co-cultured40. Furthermore, soluble VCAM-1 secreted from cardiac progenitor cells has shown cardioprotective effects through α4β1 integrin-mediated activation of Akt, ERK, and P38MAPK in cardiomyocytes41. Although these interactions are primarily observed in cardiac development during embryogenesis42, 43–44, we have observed that the injection of human fCFs expressing VCAM-1 into rats with ischemic heart failure increased the thickness of the left ventricular anterior wall9. From these perspectives, aVCFs may induce multiple responses beyond the mechanisms described in this paper and contribute to the recovery of cardiac function after MI. We plan to investigate these molecular mechanisms in depth, with Prox1 as the central signal, as previously reported45. These findings suggest that aVCFs may restore cardiac function after MI by inducing multiple biological processes other than lymphangiogenesis (Table 1, Supplementary Fig. 3).
Table 1. List of target proteins measured by ELISA or ProcartaPlex (MultiPlex).
ProcartaPlex(MultiPlex) | ELISA |
---|---|
IL-1 beta | Epiregulin |
MMP-1 | Laminin subunit beta-3 |
IL-1 alpha | SERPINB2 |
CXCL1(GROα) | IL-32 |
CXCL8 | CXCL3 |
IL-6 | IL-7R alpha |
HB-EGF | OTOGL |
IL-33R | C1qTNF1 |
CCL20 | Adrenomedullin |
CXCL2 | Cerebellin2 |
G-CSF | Collagen alpha-6(Ⅵ) chain |
IL-27 | SERPINA11 |
CXCL5 | |
GM-CSF | |
CXCL6 | |
CCL2 |
The results of this study suggest that the mechanism of aVCFs is driven by direct and indirect lymphangiogenesis. Moreover, aVCFs may be effective in treating human ischemic heart failure, as demonstrated by its impact on cardiac function recovery after MI in a rat model. Recent studies have shown that patients with heart failure over the age of 45, with NYHA functional class II or III and with reduced ejection fraction (HFrEF) due to left ventricular systolic dysfunction, who have been on routine oral medications, exhibit significantly more myocardial edema—a therapeutic target for lymphangiogenesis—compared to patients with other diseases or healthy individuals46. This suggests that the standard of care recommended in treatment guidelines, such as the current renin–angiotensin–aldosterone system (RAAS) inhibitors, has limited efficacy in addressing myocardial edema. It has been reported that myocardial edema, which results from RAAS activation, can be drained by neoplastic lymph vessels3,4,18. Therefore, aVCF, which facilitate recovery of cardiac function through lymphangiogenesis, is expected to be a novel cellular medicine for these diseases. In addition, ADM, a candidate factor for aVCF, has been reported to reduce myocardial inflammation and edema, improving acute myocarditis47.Consistent with these reports, our data also demonstrated that VCF treatment suppressed cardiac inflammation following ischemia–reperfusion injury. Specifically, the expression levels of pro-inflammatory cytokines Tnf-α and Il-1β were significantly reduced in VCF-treated mouse hearts (Supplementary Fig. 7a). Moreover, VCF administration markedly reduced macrophage infiltration, as evidenced by a decreased number of CD68+ and F4/80+ cells in heart sections (Supplementary Fig. 7b).
Importantly, while our histological analyses showed increased Prox-1+ and VEGFR-3+ cells in the aVCF-treated group, these markers are not specific to lymphatic endothelial cells. Prox-1 has been reported to be expressed in adult cardiomyocytes and to play physiological roles in cardiac tissue48. Likewise, VEGFR-3 is highly expressed by cardiac macrophages, particularly under inflammatory conditions such as myocardial infarction49, 50–51. These VEGFR-3+ macrophages are known to exhibit features of alternatively activated macrophages (M2-like), which may contribute to tissue repair and cardioprotection rather than lymphangiogenesis.
Given these findings, the observed increases in Prox-1+ and VEGFR-3+ cell populations in the nude rat model may partially reflect the activation of cardiomyocytes or macrophage polarization, not solely lymphatic vessel formation. Due to the absence of co-staining with endothelial markers such as CD31 or LYVE-1, we acknowledge that this represents a limitation of our study. Future work should incorporate multi-marker immunostaining to distinguish lymphatic endothelial cells more specifically and evaluate the contribution of non-endothelial cells to the observed effects. Accordingly, the pro-lymphangiogenic potential of aVCFs must be interpreted with caution, and the possibility of alternative mechanisms—including immune modulation and direct cardiomyocyte protection—should be further explored.
Furthermore, in our syngeneic mouse model experiments, we used Lyve-1 and VEGFR-3 as markers to assess cardiac lymphangiogenesis. However, it is important to note that both markers are also expressed by cardiac macrophages under pathological conditions. In particular, Lyve-1+ and VEGFR-3+ macrophages have been identified as a subset of M2-like macrophages that may participate in immune regulation and tissue repair52,53. This marker overlap makes it difficult to conclusively distinguish lymphatic endothelial cells from macrophages using Lyve-1 and VEGFR-3 alone. Therefore, future studies should incorporate additional endothelial markers such as CD31 or Podoplanin in combination with Lyve-1 and VEGFR-3 to more accurately identify lymphatic vessels. We acknowledge this limitation and advise cautious interpretation of our lymphangiogenesis-related findings in the mouse model. Despite these limitations, our results also highlight the potential clinical utility of aVCFs. It should also be noted that VEGFR-3+ macrophages have been reported to not only participate in immune regulation and tissue repair, but in certain contexts to transdifferentiate into lymphatic endothelial cells and directly contribute to lymphangiogenesis following myocardial injury51. Therefore, even if part of the observed VEGFR-3 signal in our study reflects macrophage populations, these cells may still promote lymphatic vessel formation either indirectly via secretion of lymphangiogenic factors such as VEGF-C or directly through lineage conversion, which could contribute to the cardioprotective effects observed after aVCF transplantation.
These cells could potentially be isolated from cardiac tissue obtained via catheter biopsy from the right ventricular septum and used as an autologous cell-based therapy. Therefore to aid in understanding the central concept and mechanism of action, a graphical abstract summarizing the enrichment of aVCFs, their transplantation, and the subsequent restoration of cardiac function is presented in Supplementary Fig. 9.
In conclusion, this study identified a population of aCFs with lymphangiogenesis-promoting activity (aVCFs) similar to that of fCFs, and established a method for their induction. In addition, it was unveiled that the aVCFs promoted lymphangiogenesis, at least in part, by secreting ADM and had cardioprotective effects against MI.
Methods
This study was reported in accordance with ARRIVE guidelines. All methods were performed in accordance with the relevant guidelines and regulations. Materials and Methods are available in the manuscript or Supplemental File.
Expansion and cell characterization of human adult cardiac fibroblasts
Human adult cardiac fibroblasts (aCF) were purchased from Lonza (Basel, Switzerland) and cultured with HFDM-1(+) medium (Cell Science & Technology, Osaka, Japan) supplemented with 1% (v/v) Newborn Calf Serum (NBCS). The expanded cells were cultured in the medium supplemented with TNF-α and IL-4 at the final concentrations of 50 ng/mL and 2 ng/mL, respectively, to upregulate VCAM-1 expression in aCF. Hereinafter, aCF treated with TNF-α and IL-4 is referred to as aCF + CKs. The medium was replenished on the second day, and the treatment was continued for a total of approximately 3 days. To create CD90+ VCAM1+ cells, aCF + CKs were labeled with anti-CD90-PE antibody (Milltenyi Biotech, CAT. #130-114-860). Subsequently, using Anti-PE MicroBeads (Miiltenyi Biotec, CAT. #130-048-801), and CD90-positive cells were collected using autoMACSpro separator (Miiltenyi Biotec). The CD90+ VCAM1+ cell fraction isolated in this manner is hereinafter referred to as aVCFs. The method used to generate aVCFs has been protected under WO2020045547A1.
Cells were characterized by flow cytometry (FCM) for various known markers of fibroblasts, cardiomyocytes, and endothelial cells. Regarding FCM analysis of proteins localized in the cytoplasm, cells were fixed and permeabilized with 0.1% saponin (Nacalai Tesque, Kyoto, Japan) in PBS for 15 min after fixation before incubation with primary antibodies. The same antibodies as described in our previous study were used9. FCM analyses were performed with a MACSQuant Analyzer following the manufacturer’s instructions (Miltenyi Biotec, Bergisch Gladbach, Germany).
Expansion of human fetal cardiac fibroblasts
Human fetal cardiac fibroblasts (fCF) were purchased from Cell Applications, Inc. (San Diego, CA, USA, CAT. # CA30605f., LOT. #1916) and cultured with HFDM-1(+) medium (Cell Science & Technology, Osaka, Japan) supplemented with 1% (v/v) Newborn Calf Serum (NBCS).
One-step RT-qPCR for Fig. 1h
Gene expression analysis for Fig. 1h was performed using a one-step RT-qPCR protocol with commercial reagents, following the manufacturer’s instructions. Primers and probes are listed in Table 2.
Table 2. Primer and probe information of ECM rigidity genes.
Forward primer | Reverse primer | Probe | |
---|---|---|---|
FAP | 5′- gaagttgaagaccagattacagc-3′ | 5′-gaccagttccagatgcaagg -3′ | /56-FAM/cctccatag/ZEN/gaccagccccatatg/3IABkFQ/ |
FSP1 (ATL1) | 5′- agccacagtatttgcccttag-3′ | 5′- gtttcctccattgccagtct-3′ | /56-FAM/aggtgctga/ZEN/agatcatcctcctgga/3IABkFQ/ |
αSMA(ACTA2) | 5′- agagttacgagttgcctgatg -3′ | 5′- ctgttgtaggtggtttcatgga -3′ | /56-FAM/agaccctgt/ZEN/tccagccatccttc/3IABkFQ/ |
POSTN | 5′- cgctattctgacgcctcaa -3′ | 5′- gttgctctccaaacctctacg -3 | /56-FAM/ttgtcccaa/ZEN/gcctcattactcggtg/3IABkFQ/ |
GAPDH(internal control) | 5′- acatcgctcagacaccatg-3′ | 5′- tgtagttgaggtcaatgaaggg-3′ | /56-FAM/aaggtcgga/ZEN/gtcaacggatttggtc/3IABkFQ/ |
Detailed reaction conditions are provided in the Supplementary Information.
Two-step RT-qPCR
Gene expression analysis was performed using a two-step RT-qPCR method for both cultured cells and mouse heart tissue. For normalization, RPL13A was used as a reference gene for cultured cells, and GAPDH for heart samples. Primer and probe sequences are listed in Tables 3 and 4.
Table 3. Primer and probe information related to ADM, NF-κB, and STAT6 signaling.
Forward primer | Reverse primer | |
---|---|---|
ADM | ATGTACCTGGGTTCGCTCGC | CCACGACTCAGAGCCCACTT |
VEGF-C | CACGAGCTACCTCAGCAAGA | GCTGCCTGACACTGTGGTA |
HGF | CTTAAAGCCTTGCCAACAGC | GCCTAGCCATGCTCT TTC |
RPL13A | CCTGGAGGAGAAGAGGAAAGAGA | TTGAGGACCTCTGTGTATTTGTCAA |
Table 4. Primer and probe information for quantitative PCR analysis of inflammatory cytokines (TNF-α and IL-1β) in mouse heart tissue.
Forward primer | Reverse primer | |
---|---|---|
TNF-α | AGCCCCCAGTCTGTATCCTT | CTCCCTTTGCAGAACTCAGG |
IL-1β | TGACAGTGATGAGAATGACCTGTTC | TTGGAAGCAGCCCTTCATCT |
GAPDH | ATGGTGAAGGTCGGTGTG | ACCAGTGGATGCAGGGAT |
Detailed procedures are provided in the Supplementary Information.
Fluorescent reporter assay
aCFs were seeded in 24-well plates and infected with lentiviral vectors encoding GFP-P65 or GFP-STAT6. GFP fluorescence was captured the following day using a Nikon Eclipse Ts2 microscope after cytokine stimulation (TNF-α and IL-4).
For plasmid-based reporter assays, aCFs were seeded in 96-well plates and transfected with either pGL4.14-5xNFkB-GLuc or pGL4.14-4xStat6-GLuc, along with pTK-CLuc, using GenomONE-GX (Ishihara Sangyo). Culture medium was replaced post-transfection with or without cytokines. Luciferase activity in collected supernatants was measured using Pierce Gaussia and Cypridina Luciferase Assay Kits (Thermo) and a PerkinElmer NIVO plate reader.
All experiments were performed in duplicate or quadruplicate.
Detailed plasmid information and cell densities are described in the Supplementary Information47.
Proliferation assay
Cells were grown in 96-well tissue culture plates, stimulated with human aCF-derived conditioned medium (CM) or aCF + CKs-derived CM or aVCF -derived CM or DMEM, and tested using a premixed WST-1 Cell Proliferation Assay System (TAKARA Bio Inc.) and a multiwell scanning spectrophotometer (440 nm; reference, 600 nm). The absorbance at 440 nm (with reference at 600 nm) was measured and correlated with the number of viable cells.
Migration assay
A transwell migration assay was performed using 24-well plates with 3-μm pore inserts (Corning)54, 55–56. Serum-starved HDLECs were seeded onto the upper chamber, and conditioned media (CM) from aCFs, aCF + CKs, or aVCFs was added to the lower chamber. After 8 h, migrated cells were fixed, DAPI-stained, and quantified under a 20 × objective lens.
Detailed procedures, including incubation and washing conditions, are provided in the Supplementary Information.
Preparation of nude rat models of ischemic heart failure, cell administration and echocardiographic observation
Animal experiments were performed as previously reported9 using F322/N Jcl-rnu/rnu male nude rats (8 weeks old, 128.7–166.6 g; purchased from CLEA Japan, Inc.). Myocardial infarction was induced by 30 min of left anterior descending artery occlusion followed by reperfusion. One week later, rats with LVEF ≤ 55% were randomly assigned to the following groups: aCF (n = 7), aCF + CKs (n = 8), aVCF (n = 9), vehicle (n = 4), and sham (n = 6).
Cell suspensions (2 × 106 cells in 50 µL of DMEM with 10% NBCS) or control medium were injected into two sites in the infarcted left ventricular wall. Echocardiography was performed every two weeks for 18 weeks. Anesthesia was induced and maintained using inhaled isoflurane (0.5–2%) delivered via a rodent anesthesia system. At the endpoint (week 18), animals were euthanized by blood withdrawal under deep anesthesia.
Detailed anesthesia, fixation, and euthanasia procedures are described in the Supplementary Information.
Mouse model of cardiac ischemia–reperfusion injury
Animal experiments were performed as previously reported9. Eight-weeks-old male mice were purchased from Charles River Laboratories.
Cardiac ischemia–reperfusion (IR) injury was induced in mice by 60 min of left coronary artery ligation followed by reperfusion, as previously described. Immediately after reperfusion, 4 × 105 VCFs with or without ADM knockdown (siADM) in 10 μL PBS were intramyocardially injected at six sites within the left ventricular myocardium. Control animals received PBS alone. Mice were anesthetized with an intraperitoneal injection of a cocktail of medetomidine hydrochloride (0.3 mg/kg), midazolam (4 mg/kg), and butorphanol tartrate (5 mg/kg). All animal procedures were approved by the Animal Ethics Review Board of the Nagoya University School of Medicine. Euthanasia was performed by cervical dislocation under anesthesia.
Detailed surgical procedures, anesthesia protocols, and ethical approvals are provided in the Supplementary Information.
Immunohistochemistry
Paraffin-embedded heart sections were used for Sirius red (SR) and hematoxylin and eosin (H&E) staining. Fibrotic area was quantified using ImageJ. Immunofluorescence staining was performed on rat cardiac sections using antibodies against cardiac troponin T (cTnT; Abcam, ab8295), Prox-1 (Proteintech, 11067-2-AP), and DAPI. Mouse heart sections were stained for CD68 (Santa Cruz, sc-20060) and F4/80 (Santa Cruz, sc-377009) with DAPI nuclear counterstaining.
Detailed image acquisition and quantification procedures are described in the Supplementary Information.
In situ hybridization (ISH)
Double staining of cardiac troponin T (cTnT) and Lyve-1 mRNA was performed on formalin-fixed paraffin-embedded (FFPE) heart sections. Anti-cTnT antibody (Abcam, ab209813) and Alexa Fluor 488-conjugated secondary antibody (Abcam, ab150077) were used for immunofluorescence staining.
ISH for Lyve-1 and Vegfr3 was performed using the ViewRNA Tissue Fluorescence Assay kit (Thermo Fisher), and autofluorescence was suppressed using the ReadyProbes™ Quenching Kit (Invitrogen, R37630). Sections were imaged using a SpinSR10 super-resolution microscope (Evident, Tokyo, Japan).
Detailed staining and hybridization procedures are provided in the Supplementary Information.
RNA extraction, library preparation, and sequencing
Total RNA from cardiac fibroblasts was extracted using the RNeasy Plus Mini Kit (Qiagen) according to the manufacturer’s protocol. RNA quality was confirmed (RIN > 7), and sequencing libraries were prepared using the NEBNext Ultra RNA Library Prep Kit for Illumina. Libraries were sequenced using an Illumina HiSeq 2500 platform.
Detailed library preparation steps and sequencing parameters are provided in the Supplementary Information.
Mapping
Technical sequences were removed using Trimmomatic (v0.30). The reference genome (Ensembl database hg38) was indexed, and the reads were aligned using HISAT2 (v2.0.1). Alternative splicing was analyzed with ASprofile (v1.0.4). Novel transcripts were predicted with Cuffcompare (part of Cufflinks v2.2.1). Single nucleotide variants were identified using samtools (v0.1.18) with command mpileup and Bcftools (v0.1.19). Gene and isoform expression was estimated with HTSEQ (v0.6.1).
Transcriptome analysis
RNA-seq was performed by GENEWIZ, and subsequent analyses were conducted in-house. Differentially expressed genes (DEGs) were filtered using a fold change threshold of ≥ 5 or ≤ 1/5. PCA was conducted on all DEGs, and the top 150 most variable genes were selected for downstream analysis.
Secreted protein-coding genes were identified based on UniProt annotations, and upregulated clusters were prioritized based on heatmap patterns.
Detailed analytical methods and software tools are described in the Supplementary Information.
Enzyme-linked immunosorbent assay (ELISA)
Each cell type was seeded onto T-25 flasks. After incubation for 5 days in serum-free DMEM, supernatant was collected as conditioned medium (CM) and residual cells and debris were removed by centrifugation. The protein concentration in culture supernatant was measured by ELISA. ELISA (R&D systems, Elabscience, Ray Biotech, MyBioSource, Abbexa, abcam, Invitrogen, LSBio) and Human ProcartaPlex Mix&Match (Invitrogen) were used according to the manufacturers’ instructions. The target proteins are listed in Table 1.
Assessment of the relationship between NF-κB/STAT6 signaling and Adrenomedullin (ADM) expression
To investigate the involvement of NF-κB and STAT6 signaling in ADM expression, aCFs were pre-treated with the NF-κB inhibitor BAY 11–7082 at a final concentration of 5 μM, one hour prior to stimulation with TNF-α (50 ng/mL) and IL-4 (2 ng/mL). Thirty minutes after the BAY 11–7082 addition, the STAT6 inhibitor AS1517499 was added at a final concentration of 100 nM. After pre-treatment, the medium containing the inhibitors was removed and replaced with fresh medium containing TNF-α and IL-4 at the indicated concentrations. The same treatment was repeated after 48 h, resulting in a total stimulation period of 72 h.
For the control group without inhibitors, TNF-α and IL-4 were added at the same time points as the inhibitor-treated group and incubated under the same conditions for 72 h.
After the stimulation period, cells were washed with calcium- and magnesium-free D-PBS and lysed using RLT buffer. RNA extraction and RT-qPCR were performed as described above. Gene expression levels were normalized to Ribosomal Protein L13A (RPL13A).
Preparation of aVCF si-ADM and aVCF si-negative control
The aCFs were seeded at a density of 20,000 cells/cm2 in a flask and cultured overnight. The next day, transfection was performed using Lipofectamine (Invitrogen™, Cat. #13,778,150). The siRNA sequences used were Silencer™ Select Negative Control No. 1 siRNA, Invitrogen™, Cat.#. 4,390,843 and Silencer® Select Pre-Designed siRNA, ambion, P/N 4,392,420.
Tube formation assay and quantitative evaluation
To assess lymphatic tube formation, HDLECs were co-cultured with aVCF-siScramble or aVCF-siADM at a 4:6 ratio in Endothelial Cell Basal Medium MV2 (PromoCell) containing 0.5% FBS. After 14 days of direct co-culture, immunofluorescence staining was performed using Anti-VE-Cadherin (Abcam, ab33168), Anti-Vimentin (Abcam, ab20346), Alexa Fluor 488-conjugated anti-rabbit IgG (Abcam, ab150077), Alexa Fluor 647-conjugated anti-mouse IgG (Abcam, ab150115), and Hoechst 33,258 (Dojindo, 343–17,961).
Detailed staining protocols, image acquisition, and quantitative analysis are provided in the Supplementary Information.
Expansion and cell characterization of mouse cardiac fibroblasts
Mouse cardiac fibroblasts (mCF) were isolated by primary culture from the hearts of male C57BL/6 J mice at 10 weeks of age. All animal procedures were approved by the Animal Care and Use Committee of the Innovation Center of NanoMedicine (Kawasaki, Kanagawa, Japan). Following euthanasia, the hearts were harvested, perfused to remove blood, and washed thoroughly. The hearts were then transferred into GentleMACS C Tubes (Miltenyi Biotec, Bergisch Gladbach, Germany) and dissociated using the Multi Tissue Dissociation Kit 2 (Miltenyi Biotec, Bergisch Gladbach, Germany; CAT. #130-110-203) according to the manufacturer’s protocol to obtain a cell pellet. The cell pellet was seeded into culture dishes containing D-MEM (High Glucose) supplemented with 10% fetal bovine serum (FBS) and 1% penicillin/streptomycin (P/S). On the following day, debris was removed by replacing the medium. The cells were expanded to passage 1 (P1), and then cultured in medium supplemented with TNF-α (PeproTech, Rocky Hill, NJ, USA; CAT. #315-01A) and IL-4 (PeproTech, Rocky Hill, NJ, USA; CAT. #214-14) at final concentrations of 50 ng/mL and 2 ng/mL, respectively, to upregulate VCAM-1 expression in mCF. The medium was replenished with fresh TNF-α and IL-4 at the same final concentrations 48 h after the initial supplementation, and the treatment was continued for approximately 3 days.
Following the upregulation treatment, the cells were labeled with anti-CD90.2-PE antibody (Miltenyi Biotec, Bergisch Gladbach, Germany; CAT. #130-102-489) and anti-CD106 (VCAM-1)-APC antibody (Miltenyi Biotec, Bergisch Gladbach, Germany; CAT. #130-104-713). Subsequently, using Anti-PE MicroBeads (Miltenyi Biotec, Bergisch Gladbach, Germany; CAT. #130-048-801), CD90.2-positive cells were isolated using the autoMACS Pro Separator (Miltenyi Biotec, Bergisch Gladbach, Germany) with the Depl05 program, and defined as mVCF.
Statistical analysis
Data are presented as mean ± SEM. Statistical tests appropriate for distribution and group comparisons were applied using GraphPad Prism. A P-value < 0.05 was considered statistically significant.
Detailed test selection and conditions are described in the Supplementary Information.
Acknowledgements
We would like to express our gratitude to the staff at LSI Medience Corporation for their invaluable contributions in establishing the rat ischemia-reperfusion model and conducting the cell administration experiments. Furthermore, the authors wish to acknowledge Division for Medical Research Engineering, Nagoya University Graduate School of Medicine for usage of SpinSR10 (Evident, Tokyo, Japan).
Author contributions
Y.M., conceived scientific idea, designed and performed experiments, analyzed data, wrote and edited manuscript.; Y.S., conceived scientific idea, designed experiments, analyzed data, wrote and edited manuscript, supervised project.; H.L. performed experiments, analyzed data.; B.S. performed experiments, analyzed data, edited manuscript.; M.M. performed experiments, analyzed data, edited manuscript.; T.H. performed experiments analyzed data.; T.M., conceived scientific idea, supervised project.; T.I. conceived scientific idea, designed and performed experiments, analyzed data, wrote and edited manuscript, supervised project.
Funding
The Ministry of Education, Culture, Sports, Science and Technology of Japan, 25K11361.
Data availability
The data, analytic methods, and materials employed in this study are available from the corresponding author upon reasonable request. The datasets for Gene expression data generated and/or analyzed during the current study are available in the ArrayExpress repository (E-MTAB-14788; https://www.ebi.ac.uk/biostudies/ArrayExpress/studies/E-MTAB-14788?key=187241b5-2f5e-47c1-91eb-7134f097451a).
Declarations
Competing interests
The authors declare no competing interests.
Abbreviations
CKsCytokines
CFCardiac fibroblast
DAPI6-Diamidino-2-phenylindole
ELISAEnzyme-linked immunosorbent assay
EFEjection fraction
PBSPhosphate-buffered saline
qPCRQuantitative polymerase chain reaction
VCAM1Vascular cell adhesion molecule 1
Supplementary Information
The online version contains supplementary material available at https://doi.org/10.1038/s41598-025-17224-6.
Publisher’s note
Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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Abstract
Myocardial infarction (MI) remains the leading cause of death worldwide. We previously found that a specific population of human fetal cardiac fibroblasts (fCFs), which express vascular cell adhesion molecule 1 (VCAM1), have cardioprotective effects after MI, inducing reparative cardiac lymphangiogenesis. This study investigated whether adult cardiac fibroblasts (aCFs), which are more feasible for autologous transplantation, differ in surface marker expression and lymphangiogenic potential compared to fCFs. Furthermore, we examined whether aCFs could be exogenously manipulated to acquire fCF-like lymphangiogenic potential and serve as a cell therapy for MI and MI-associated heart failure. In vivo MI models (rat and mouse) and in vitro coculture assays with lymphatic endothelial cells were conducted. We found that TNF-α and IL-4 stimulation induced aCFs to express VCAM1 via NF-κB and STAT6 signaling, yielding a subpopulation termed adult VCAM1+ cardiac fibroblasts (aVCFs). These aVCFs, distinct from myofibroblasts, expressed CD90 and improved cardiac function post-MI. Adrenomedullin (ADM) was identified as a key paracrine effector, and its knockdown attenuated the pro-lymphangiogenic and cardioprotective effects of aVCFs. Our findings demonstrate that aVCFs promote cardiac lymphangiogenesis and protect cardiac function following MI, highlighting their potential as an autologous cell therapy.
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Details
1 Department of Cardiology, Nagoya University Graduate School of Medicine, 65 Tsurumai, Showa-ku, 466-8550, Nagoya, Japan (ROR: https://ror.org/04chrp450) (GRID: grid.27476.30) (ISNI: 0000 0001 0943 978X); Advanced Regenerative Cell-Based Healthcare Education for Resources, Nagoya University Graduate School of Medicine, Nagoya, Japan (ROR: https://ror.org/04chrp450) (GRID: grid.27476.30) (ISNI: 0000 0001 0943 978X); Research & Development Department, LYMPHOGENiX Ltd, Covent Garden, London, England (ROR: https://ror.org/05wnh3t63) (GRID: grid.421947.d) (ISNI: 0000 0004 1782 6335)
2 Department of Cardiology, Nagoya University Graduate School of Medicine, 65 Tsurumai, Showa-ku, 466-8550, Nagoya, Japan (ROR: https://ror.org/04chrp450) (GRID: grid.27476.30) (ISNI: 0000 0001 0943 978X)
3 Advanced Regenerative Cell-Based Healthcare Education for Resources, Nagoya University Graduate School of Medicine, Nagoya, Japan (ROR: https://ror.org/04chrp450) (GRID: grid.27476.30) (ISNI: 0000 0001 0943 978X)
4 Department of Otorhinolaryngology, Faculty of Medicine, Juntendo University, Tokyo, Japan (ROR: https://ror.org/01692sz90) (GRID: grid.258269.2) (ISNI: 0000 0004 1762 2738)
5 Advanced Regenerative Cell-Based Healthcare Education for Resources, Nagoya University Graduate School of Medicine, Nagoya, Japan (ROR: https://ror.org/04chrp450) (GRID: grid.27476.30) (ISNI: 0000 0001 0943 978X); Research & Development Department, LYMPHOGENiX Ltd, Covent Garden, London, England (ROR: https://ror.org/05wnh3t63) (GRID: grid.421947.d) (ISNI: 0000 0004 1782 6335); Institute for Advanced Biosciences, Keio University, Tsuruoka, Yamagata, Japan (ROR: https://ror.org/02kn6nx58) (GRID: grid.26091.3c) (ISNI: 0000 0004 1936 9959)