All mice were placed in an environment with controlled light cycles, temperature, and humidity. The cardiac hypertrophy model was constructed by transverse aortic constriction (TAC) surgery as previously described (15). Briefly, male mice with C57BL/6 background (10-week-old; weight: 25–27 g) were anesthetized via i.p. injection of pentobarbital sodium (90 mg/kg, P3761, Sigma-Aldrich). After exposing the transverse aorta, the transverse aorta was ligated transversely with 7–0 silk and a 26-gauge needle. Identical operation without ligation the transverse aorta was performed in the sham operation group.

Seven days after TAC or sham surgery, mice were randomly given vehicle or luteolin treatment. Mice were administered orally with luteolin (40 mg/kg) or vehicle [1% carboxymethyl cellulose sodium (CMC-Na)] daily for 11 consecutive weeks. After 4, 8 and 12 weeks of TAC or sham surgery, heart function was respectively measured by echocardiography as previously described (16). At the end of the experiment, the mice were anesthetized using the above method and then subjected into euthanasia via cervical dislocation, and the heart, lung, and tibia were collected for further examinations.

Echocardiography was used to evaluate mice cardiac function as described previously (16). A small animal ultrasound imaging system (Mylab30CV, ESAOTE, S. P. A) was used to perform echocardiography. The left ventricle was evaluated on both long- and short-axis views of the parasternal sternum as described previously. The echocardiography operator is not informed about the grouping of mice.

Twelve weeks after TAC or sham surgery, mice hearts were collected. Hearts were macerated in 10% formalin and subsequently encapsulated in paraffin. Paraffin-embedded mice hearts were sectioned transversely (5 μm). Subsequently, hematoxylin-eosin and picrosirius red staining (for collagen volume fraction analysis) were executed. These analyses were performed using Image-Pro Plus 6.0. At least one hundred cardiomyocytes were examined in each section, and the collagen volume fraction was computed as picrosirius red staining area divided by total area.

Primary cardiomyocytes were obtained from Sprague-Dawley rat (1–2 days) hearts in accordance with previously described (17). Primary neonatal rat cardiomyocytes (NRCMs) were incubated in the DMEM/F12 medium with 10% fetal bovine serum, 1% penicillin/streptomycin, and 0.2 mM BrdU for 48 h. After culturing NRCMs in serum-free DMEM/F12 for 12 h, hypertrophy was induced by adding 50 μM phenylephrine (PE, P6126, Sigma) for 24 h. DMEM medium with 1% penicillin/streptomycin and 10% fetal bovine serum was used for the incubation of H9C2 cells and HEK 293 T cells.

The size of the cardiomyocyte surface was assessed by immunofluorescence staining for α-actinin after 24 h of incubation with PBS or PE as previously described in the established protocol (15, 16). Briefly, cardiomyocytes were successively soaked in 4% formaldehyde and 0.2% Triton-X 100 (T8787, Sigma-Aldrich). Then the cells were stained with α-actinin (1:100 dilutions, A7811, Sigma) and appropriate secondary antibody (1:200 dilutions, A11061, Invitrogen). The size of the cardiomyocyte surface was examined with Image-Pro Plus 6.0.

Rat Bnp (b-type natriuretic peptide) promoter (−2147, +132 bp) and Myh7 (myosin heavy chain 7) promoter (−2500, +89 bp) were expanded and cloned into pGL3-promoter luciferase reporter vector to obtain promoter reporter plasmids. The lentiviral plasmids encoding shRNA for Pparγ was constructed in the pLKO.1 vector. HEK 293 T cells were transfected with pLKO-shRNA or pLKO-sh Pparγ along with the packaging vectors pSPAX2 and pMD2G. After incubation of cells at 37°C for 40 h, lentiviral suspensions were harvested for infection with H9C2 cells followed by puromycin selection for one week for various analyses. Primers for plasmid and lentivirus construction are provided in Supplementary Table S1.

HEK 293 T cells were co-transfected with a plasmid expressing Firefly luciferase and a plasmid expressing Renilla luciferase. After 24 h, FDA-approved compounds (20 μM, L1300, Selleckchem), containing 2,570 compounds, were added separately to the culture medium. The cells were lysed using lysis solution, and the luciferase activity was examined by adding luciferase assay substrate (E1980, Promega). The results were shown as log2-fold change normalized to control.

The Cell Counting Kit (CCK-8, Beyotime) was applied to assay the cell viability of NRCMs. NRCMs were incubated in 96-well plates with 2 × 10⁴ cells per well and subsequently treated with different concentrations of luteolin. After the addition of CCK-8 solution, the NRCMs were further incubated. After 24 h of incubation, the absorbance of each well at 450 nm was detected.

After 24 h transfection with the appropriate plasmids, HEK293T cells were treated with biotin-linked luteolin (20 μM) or biotin for another 4 h and then lysed in pre-cooled immunoprecipitation (IP) buffer. Add the specified antibodies and protein G Bestarose 4FF beads (AA304307, Bestchrom) to the lysate supernatant and incubate for 4 h. The immunocomplexes were washed with pre-cooled IP buffer and subsequently collected for western blot analysis.

Proteins were obtained from cardiac tissue and cardiomyocytes. The BCA Protein Assay Kit (23225, Thermo) was applied to detect protein levels. Proteins were isolated by SDS-PAGE and electrotransferred to PVDF membranes, which were successively incubated with primary and secondary antibodies. Protein signals were detected using the ChemiDoc MP Imaging System (Bio-Rad). GAPDH served as the loading control. All antibodies are shown in Supplementary Table S2.

Cultured HEK 293 T cells were collected and lysed in SDS lysis buffer including protease inhibitor cocktail (04693132001, Roche). The lysates were centrifuged (12,000 rpm for 15 min), and the supernatants were analyzed by immunoprecipitation assays with indicated antibodies, followed by western blot analyses.

RNA was derived from mice heart tissue and NRCMs with Trizol regent (T9424, Sigma-Aldrich), and reverse transcription was conducted by the HiScript III RT SuperMix for qPCR (R323-01, Vazyme). Quantitative real-time PCR was conducted to detect gene expression level using SYBR Green (Q311-03, Vazyme), and Gapdh served as internal reference gene (18). The real-time PCR primers are shown in Supplementary Table S3.

RNA extracted from mice hearts and cultured NRCMs was used for library preparation. Single-end RNA-seq was carried out with MGISEQ-2000 (MGI, China). Based on the Ensembl mouse (mm10/GRCm38) genome, gene sequence alignment on clean reads was conducted by HISAT2 software (version 2.1.0). SAMtools software (version 1.4.1) was used to transform the mapped fragments into the Binary Alignment Map files. Then, the reads count value of the genes were computed using StringTie (version v1.3.3b). DESeq2 (v1.2.10) software was used for differentially expressed genes identification by processing read count information. Gene with adjusted P-values less than 0.05 and a fold change larger than 1.5 was recognized as differentially expressed gene. Gene Ontology analysis, Gene Set Enrichment Analysis (GSEA), and Kyoto Encyclopedia of Genes and Genomes analysis were conducted by R package clusterProfiler (version 4.2.2). Gene sets with adjusted P value of less than 0.05 were recognized as statistically significant. The R package gmodels (version 2.18.1) was used for principal component analysis.

SPSS 22.0 was applied to analyze all results and the results were presented as the mean ± SD. When the data were normally distributed, 2 group comparisons were conducted by 2-tailed Student t test, and multiple comparisons were conducted by one-way ANOVA. When data were skewed distributed, multiple comparisons were conducted by Kruskal-Wallis test. P < 0.05 was regarded as statistically significant.

Upregulation of Myh7 and Bnp transcriptional activity is an indication of the risk for developing cardiac hypertrophy (19, 20). To explore potential therapeutic drugs for cardiac hypertrophy and HF, our study screened the effects of FDA-approved chemical compounds library on the transcriptional activity of Myh7 and Bnp based on luciferase reporter assays (Figure 1A). The compounds were added to HEK 293 T cells transfected with Myh7 and Bnp promoter reporter plasmids, respectively, and luciferase activity was assayed after 24 h of incubation. Among 2,570 compounds, a total of five were screened as the candidates according to the criteria of downregulating Myh7 and Bnp promoter activity for more than 50% (Figure 1B). After excluding two candidate compounds with significant cardiotoxicity, three candidate compounds were further examined for their effects on PE-induced cardiomyocyte hypertrophy (Figure 1C). Notably, luteolin, a flavonoid from a variety of plants (10), produced the most pronounced protective effect compared to dimethyl sulfoxide (DMSO) as indicated by immunofluorescence staining (Figures 1C,D).

The cellular safety of luteolin was confirmed by its non-significant impact on cell viability of NRCMs (Figure 2A). We continued to treat PE-stimulated NRCMs with different dosages of luteolin to further evaluate its effect on the hypertrophy of NRCMs. Immunostaining results demonstrated that luteolin inhibited the increase in surface area of cardiomyocytes in a dose-dependent way (Figure 2B). Similarly, the mRNA levels of Anp (atrial natriuretic peptide), Bnp, and Myh7 were also significantly inhibited by luteolin (10 μM) (Figure 2C). To explore the effect of luteolin at the panoramic molecular level, RNA-sequencing analysis was conducted on PE-treated NRCMs in the presence or absence of luteolin. By principal component analysis, the transcriptome profiles were clearly divided into two clusters (Figure 2D). The volcano plot results indicated a huge number of differentially expressed genes between the two groups (Figure 2E). GSEA analysis based on gene ontology database showed that the genes modulated by luteolin were mainly concentrated in cardiac hypertrophy, fibrosis, and protein synthesis (Figure 2F). The heatmap showed that luteolin markedly repressed the expression levels of genes associated with the aforesaid pathways (Figure 2G). These results show that luteolin inhibits cardiomyocyte enlargement in vitro.

To evaluate whether luteolin ameliorated heart failure in mice, we randomly divided wild type mice (C57BL/6) into two groups for TAC or sham surgery. Each group were further randomly divided into two groups and orally administered with luteolin (40 mg/kg) or vehicle (1% CMC-Na) once daily at one week after surgery (Figure 3A). Cardiac function at 4, 8, and 12 weeks after TAC or sham surgery was detected by echocardiography, respectively (Supplementary Table S4). After 4 weeks of TAC, echocardiographic evaluation showed signs of cardiac hypertrophy and reduced cardiac function in TAC-treated mice compared to sham control, based on increased left ventricular end-diastolic diameter (LVEDd) and left ventricular end-systolic diameter (LVESd), reduced ejection fractions (EF) and fractional shortening (Figure 3C). Compared to the vehicle group, luteolin inhibited cardiac hypertrophy and deterioration of cardiac function in TAC mice (Figure 3C). After 12 weeks of TAC, mice were sacrificed for phenotypic and histological examination. Luteolin significantly blocked pressure overload-induced cardiac enlargement in TAC-treated mice (Figure 3D). Furthermore, echocardiographic evaluation, including EF, fractional shortening, stroke volume, cardiac output, LVEDd, LVESd, left ventricular end-diastolic volume, and left ventricular end-systolic volume, further verified the protective function of luteolin administration against cardiac remodeling and decreased cardiac function in TAC-treated mice after 12 weeks of TAC (Figures 3E,F). Notably, luteolin administration showed negligible influences in the sham settings (Figures 3B–F).

Sustained pathological overload induces maladaptation and cardiac remodeling, including cardiomyocyte hypertrophy, fibrosis, capillary rarefaction, cellular dysfunction, and other complex responses, ultimately leading to heart failure (21, 22). After 12 weeks of TAC, luteolin-treated TAC mice had significantly attenuated cross-sectional area of cardiomyocytes compared with TAC mice, as indicated by histological analysis with hematoxylin-eosin staining (Figures 4A,B). Furthermore, PSR staining of heart sections showed that luteolin treatment markedly inhibited cardiac fibrosis in TAC mice (Figures 4C,D). Consistent with these data, the expression levels of Anp, Bnp, Myh7, Col3a1 (collagen type III alpha 1), Col1a1 (collagen type I alpha 1), and Ctgf (connective tissue growth factor) were also notably downregulated in luteolin-treated TAC mice compared to the vehicle group (Figures 4E–G). These data clearly support that luteolin treatment attenuated the pathological cardiac remodeling induced by pressure overload.

It is known that in the case of pathological hypertrophy, the heart undergoes metabolism reprogramming characterized by an increased reliance on glucose metabolism and a reduced reliance on fatty acid oxidation. This metabolic profile reduces the capacity for ATP synthesis and ultimately promotes the progression of heart failure (23, 24). We detected the mRNA levels of genes associated with fatty acid and glucose metabolism in the mouse failing hearts. The mRNA levels of PPARγ coactivator-1α/1β, the critical regulators of fatty acid uptake and oxidation, were markedly reduced in TAC hearts, while luteolin attenuated this change (Figure 5A). Consistently, luteolin treatment reversed the reduction of mRNA levels of genes associated with fatty acid uptake (Figure 5B) and fatty acid oxidation (Figure 5C) in the heart from TAC-treated mice. The administration of luteolin treatment also reversed increased levels of genes associated with glucose metabolism (Figure 5D).

Considering the protective effects of luteolin on cardiac hypertrophy and HF, we tried to reveal the underlying molecular mechanisms of luteolin to ameliorates myocardial hypertrophy and heart failure. We analyzed RNA-sequencing data obtained from in vivo models. Principal component analysis revealed that the transcriptome profiles were clearly divided into two clusters (Figure 6A). GSEA analysis showed that the genes modulated by luteolin were mainly concentrated in cardiac hypertrophy, fibrosis, and protein synthesis (Figure 6B). The heatmap showed that luteolin markedly repressed the expression levels of genes associated with the aforesaid pathways (Figure 6C). To identify potential targets of luteolin, we performed a combined analysis of drug-target interacting investigations and RNA-sequencing (Figure 6D). First, Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis demonstrated that PPAR signaling pathway was the most notably enriched molecular event regulated by luteolin (Figure 6E). Moreover, we performed gene set enrichment analysis, which further confirmed the upregulated PPAR signaling in the luteolin-treated group (Figure 6F). Then, we queried the protein data bank (PDB) and obtain 10 candidates interacting with luteolin (Figure 6G, Supplementary Table S5).

Combining the results of KEGG analysis with PDB data, we hypothesized that PPARγ may be a crucial target for luteolin in the inhibition of cardiac hypertrophy and HF. In order to determine whether luteolin could bind to PPARγ protein, we synthesized the chemical probe biotin-labeled luteolin (biotin-luteolin) and performed the biotinylated protein interaction pull-down assay. The data confirmed the interaction between luteolin and PPARγ (Figure 6H). Furthermore, we validated that luteolin dose-dependently activated the transcriptional activity of PPARγ in cultured HEK 293 T cells transfected with the PPAR response element (PPRE) reporters and PPARγ (Figure 6I). Thus, we speculated that PPARγ might be the direct target required for luteolin to exert a protective effect in cardiac hypertrophy and HF.

PPARγ is a critical modulator against cardiac hypertrophy (25–27), and its agonists have been found to inhibit cardiomyocytes hypertrophy by improving metabolic homeostasis and inflammatory response (28). To further evaluate whether PPARγ activation is required for the protective effect of luteolin, we cotreated NRCMs by luteolin in combined with a PPARγ activation inhibitor, GW9662. Remarkably, GW9662 treatment largely eliminated the effect of luteolin in ameliorating PE-induced cardiomyocyte hypertrophy, as shown by immunofluorescence staining and immunoblotting analysis (Figures 7A–D). Furthermore, we constructed Pparγ knockdown H9C2 cells (Figure 7E). Consistently, Pparγ knockdown abrogated the protective effect of luteolin on PE-induced expression of the relevant cardiac hypertrophy markers (Figures 7F,G).

To explore the mechanism by which luteolin modulates PPARγ, we detected the mRNA and protein levels of PPARγ in heart samples from mice. Luteolin had no significant effect on PPARγ mRNA expression, whereas luteolin administration largely blocked TAC-induced decrease in PPARγ protein expression levels in mouse heart samples at 12 weeks after TAC surgery (Figures 8A,B). These results indicate that luteolin may activate PPARγ by regulating its protein stability. By treating NRCMs with cycloheximide (CHX), the half-life of PPARγ protein was remarkably extended in luteolin-treated NRCMs (Figure 8C). Furthermore, we found that the proteasome inhibitor MG132, rather than the lysosomal inhibitor chloroquine (Chlq), reversed CHX-induced destabilization of PPARγ, indicating that PPARγ was degraded mainly in a proteasome-dependent way (Figure 8D).

Subsequently, we investigated the impact of luteolin on PPARγ ubiquitination levels and found a dramatically reduced ubiquitination level of PPARγ by luteolin treatment (Figure 8E). A previous study reported that tripartite motif containing 55 (TRIM55), a muscle-specific ubiquitin ligase, mediates the ubiquitination-mediated degradation of PPARγ in myocardial tissue (29). We first confirmed the interaction with and enhancing ubiquitination function of TRIM55 on PPARγ (Figures 8F–H), and further examined the impact of luteolin on PPARγ and TRIM55 complex. The data revealed that luteolin weakened the interaction between TRIM55 and PPARγ and largely reversed TRIM55-induced enhancement of PPARγ ubiquitination level (Figures 8F–H). In summary, PPARγ is the direct pharmacological target of luteolin, and luteolin stabilizes PPARγ protein expression by inhibiting TRIM55-mediated ubiquitinational degradation (Figure 8I).