Inactivation of Notch1-TGF-β-Smads Signaling Pathway by Atorvastatin Improves Cardiac Function and Hemodynamic Performance in Acute Myocardial Infarction Rats

Acute myocardial infarction (AMI) triggers coronary artery occlusion and subsequent cellular hypoxia, which in turn leads to the release of intracellular substances that induce cellular damage or necrosis [1]. Then inflammatory response is initiated by myocardial cells, causing myocardial fibrosis that is a crucial pathological process for cardiac remodeling [1]. In the presence of excessive accumulation of extracellular collagen, fibrosis is triggered resulting in increased myocardial ventricular tension and ultimately cardiac insufficiency [1]. There are two types of myocardial fibrosis, including alternative fibrosis caused by the deposition of collagen between muscle cells and interstitial fibrosis occurring when collagen fills the space previously occupied by necrotic cardiomyocytes. Collagen generation is an important process in myocardial repair. Myocardial fibrosis and scar production increase ventricular tension and lead to cardiac insufficiency [2].

Several studies have reported that statins have effects on the cardiovascular system such as the lipid-lowering function [3‒7]. Statins also increase the stability of atherosclerotic plaques; reduce the proliferation, migration, and number of smooth muscle cells; decrease collagen content in atherosclerotic plaques; inhibit oxidative stress and inflammatory reaction; alleviate cardiac hypertrophy and left ventricular remodeling; and inhibit neuroendocrine activation [3‒7]. It remains unclear whether atorvastatin has anti-inflammatory and anti-fibrotic effects in AMI rats. In addition, it is not completely clear whether the anti-inflammatory and anti-fibrotic effects of atorvastatin are related to the transforming growth factor-β (TGF-β), Smad, and notch1 signal pathways.

Male Sprague-Dawley rats (6–7 weeks, 210 ± 30 g) were fed with the standard protocols provided by the Animal Laboratory Center at a temperature of 22°C–24°C and humidity of 45–55%. All procedures followed were approved by the Institutional Animal Care and Use Committee (Approval No.: IRB2022-DW-13). The current study was carried out in accordance with the Guide for the Care and Use of Laboratory Animals and followed the Helsinki Declaration about animal studies.

Male SD rats were anesthetized using inhalation of fluorinated hydrocarbon after a fasting period of 12 h. Tracheal intubation was performed for assisting breathing. The chest was opened between the 3rd and 4th lib in the left chests. The pericardium was removed, and the heart was exposed. The left anterior descending coronary arteries at the intersection of left atrium appendages and pulmonary artery cones were ligated using a 7-0 monofilament suture (Peters Surgical, Bobigny, France). Myocardial infarction was confirmed by dyskinesis of the ischemic region and epicardial cyanosis. The intercostal space was sutured, and the tracheal intubation was removed until the rats were awakened. To prevent infection, the rats were continuously injected with 400,000 units of penicillin for 3 days. The sham group rats underwent the same surgical procedures without the final ligation of the coronary artery. Rats underwent echocardiography on the day after surgery. Rats with left ventricular ejection fraction (LVEF) ≤50% were successfully established as AMI models. The modeling process is shown in Figure 1.

The animals were randomized into 4 groups: (i) sham control group (n = 12), (ii) AMI group (n = 15), (iii) AMI + atorvastatin group (n = 12), (iv) AMI + losartan potassium group (n = 13). Atorvastatin and losartan potassium were dissolved in sodium carboxymethyl cellulose. All rats were subjected to gavage of sodium carboxymethyl cellulose solution. Rats in the atorvastatin group were administrated with atorvastatin (10 mg/kg/day) by gavage. Rats in the losartan potassium group were treated with losartan potassium (5 mg/kg/day) by gavage. Rats were treated with the drugs or vehicle on the first day after the surgery, and the drug was administrated for 28 days. All rats were euthanized on day 28 without suffering.

Echocardiography was conducted 28 days after drug administration using Vevo 2100 (VisualSonics, Ontario, Canada). The rats were continuously anesthetized with the mixture of 1% oxygen and 2% isoflurane and then monitored by electrocardiogram equipment. The left ventricular anterior wall end-diastolic thickness (LVAWd) was measured. The left ventricular internal diameter was examined to detect LV systolic function. The left ventricular posterior wall thickness, left ventricular internal diameter, left ventricular ejection fraction (EF), fractional shortening (FS), left ventricular volume, aortic value peak velocity during the systolic and diastolic phases were performed using the parasternal long-axis view. These indices were measured in at least three cardiac cycles, followed by calculating the mean values.

About 28 days after administration, all rats were anesthetized using inhalation of fluorinated hydrocarbon. The skin of the rat was incised in the middle of the neck, and its right common carotid artery was exposed by blunt dissection. The distal end of the right common carotid arteries was ligated with a 4-0 silk thread. The proximal ends were clipped by the bulldog clamp for blood occlusion. An arterial catheter was inserted into the right heart common carotid artery. The catheter was then pushed to the left ventricle along the right common carotid artery. The data were recorded, and the position of the catheter was adjusted according to the pressure curve.

At least three rat heart tissues were collected from each group, and the isolated hearts were quickly separated on ice bags. The separated hearts were then rapidly cooled in liquid nitrogen and stored at −80°C for future use. Simultaneously, the remaining rat heart tissues in each group was immersed in 10% formalin solution for 1 month and then embedded in paraffin to prepare sections (4 μm). Sections were deparaffinized and stained using hematoxylin and eosin stain. Masson’s trichrome staining was used to detect collagen, muscle fibers, and nucleus. Notch1, TGF-β, and Smad2 were detected by immunohistochemical staining.

To extract proteins, 300 mg of frozen heart tissues were weighted and lysed with 1 mL of radioimmunoprecipitation assay buffer (Beyotime, Nantong, Jiangsu, China). The bicinchoninic acid protein assay kit was used to detect protein concentration. The denatured proteins were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis and transferred to nitrocellulose (NC) membranes (Millipore, Billerica, MA, USA). NC membranes were blocked with 5% milk powder for 2 h at room temperature. Next, the membranes were probed with the primary antibodies and incubated for 24 h at 4°C. The primary antibodies used included anti-notch1 antibody (CST4380; Cell Signaling Technology, Danvers, MA, USA), anti-Smad2 antibody (ab92486; Abcam, Cambridge, UK), anti-GAPDH antibody (ab181602), anti-collagen III antibody (ab7778), anti-collagen I antibody (ab34710), and anti-Gal-3 antibody (ab76466). After washing in phosphate-buffered saline, the NC membranes were incubated with secondary antibody labeled by horseradish peroxidase. Immunoreactive protein bands were visualized by enhanced chemiluminescence. GADPH was an internal reference. The protein expression was calculated in gray values with the Image J software. Heart tissues from at least three rats in each group were tested.

At 28 days after administration, 30 mg of heart tissues were subject to RNA extraction with the TRIzol reagent (Invitrogen, Waltham, MA, USA). RNA concentration was measured on a nucleic acid spectrophotometer (Thermo Fisher Scientific, Waltham, MA, USA). cDNA was reversely synthesized using 100 ng RNA using the Transcriptor First Stand cDNA Synthesis Kit according to the instructions. Then, DNA amplification was performed using the SYBR Green PCR Master Mix kit under the required reaction conditions (25°C for 10 min, 50°C for 60 min, 85°C for 5 min). The primer sequences are shown in Table 1. GAPDH served as an internal reference. The mRNA expression of notch1, TGF-β, smad2, and smad7 was calculated according to 2^−△△CT formula.

SPSS 17.0 software was used for statistical analysis. Numerical variables were expressed as mean ± standard deviation. The post hoc test was utilized in the group comparison. p < 0.05 represented statistical difference from comparison among groups analyzed by one-way analysis of variance.

The cardiac function of AMI rats was assessed by echocardiography, and the findings are shown in Figure 2. On day 28, a significant decrease was noticed in LVAWd, LVAWs, EF, FS, and aortic valve (AV) peak in the AMI group compared with sham group (all p < 0.05). In contrast, compared with the AMI group, the LVAWd, LVAWs, EF, FS, and AV peak value increased in AMI rats after administration with atorvastatin or losartan (p < 0.05). No significant differences were noticed between atorvastatin and losartan groups in terms of LVAWd, EF, FS, and AV peak (p > 0.05), except for LVAWs (p < 0.01). Left ventricular internal dimension at end-diastole (LVIDd), left ventricular internal dimension at end-systole (LVIDs), left ventricular end-diastolic diameter (LVEDd), and left ventricular end-systolic diameter (LVESd) showed significant increase in the AMI group compared with the sham group (all p < 0.05). In contrast, compared with the AMI group, LVIDd, LVIDs, LVEDd, and LVESd showed significant decrease in animals treated with losartan (all p < 0.05). Losartan reduced LVIDd, LVIDs, LVEDd, and LVESd more effectively than atorvastatin (p < 0.01). There was no difference in left ventricular posterior wall end-diastolic thickness and left ventricular posterior wall end-systolic thickness between the treatment groups and the AMI group (all p > 0.05).

The terminal hemodynamics on day 28 are summarized in Figure 3. CO was reduced in the AMI group versus the sham rats (p < 0.05). A trend toward preservation of CO was apparent in atorvastatin- and losartan-treated rats compared with that of the AMI group (p < 0.05). A significant decrease was noticed in stroke volume of the AMI group compared with the sham group (p < 0.05). Compared with the AMI group, significant increase was noticed in the stroke volume in the atorvastatin and losartan groups (p < 0.05). Significant reduction was observed in the dP/dt_max in the AMI rats compared with the sham rats (p < 0.05). In contrast, dP/dt_max in rats treated with atorvastatin and losartan was preserved compared with the AMI rats (p < 0.05). Compared with the sham group, significant increase was noticed in the dP/dt_min in the AMI rats, and significant decrease was seen in the dP/dt_max in atorvastatin and losartan group compared with the AMI rats (all p < 0.05). Significant differences were noticed in dP/dt Max between the atorvastatin and losartan groups (p < 0.05).

The induction of protein expression of notch1, smad2, collagen I, collagen III, and Gal-3 was quantified by Western blot analysis. On day 28, significant increase was observed in the expression of notch1, smad2, collagen I, collagen III, and Gal-3 in the AMI group compared with the sham group (all p < 0.05, Fig. 4). Compared with the AMI group, notch1, smad2, collagen I, collagen III, and Gal-3 were decreased by atorvastatin and losartan (all p < 0.05). There was a significant difference in Gal-3 expression between the atorvastatin and losartan groups (p < 0.05).

Figure 5 showed the mRNA expression of notch1, TGF-β, smad2, and smad7. There was a significant increase in the relative expression of notch1, TGF-β, and smad2 in the AMI group compared with the sham group (all p < 0.05). In contrast, compared with the AMI group, notch1, TGF-β, and smad2 were significant decreased by atorvastatin and losartan (p < 0.05). There was a significant difference in TGF-β between atorvastatin and losartan groups (p < 0.05). The relative expression of smad7 was significantly decreased in the AMI group compared to the sham group (all p < 0.05). In contrast, compared with the AMI group, smad7 showed significant increase in the groups treated with atorvastatin or losartan (p < 0.05).

Hematoxylin and eosin staining (Fig. 6a) showed that the myocardial cells in the sham group were arranged regularly, and the intercalated discs of the myocardium were well structured. Clear infarction zones and loss of myocardial cells were observed in AMI group. Fibrocytes and collagen fiber bundles were generated. Disordered myocardial cells and inflammatory cell infiltration were observed in the MI border zones. In atorvastatin group, disordered myocardial cells were observed in the infarction border zones, together with a small number of inflammatory cells. In losartan group, disordered and reduced numbers of myocardial cells were observed with a small infiltration of inflammatory cells.

Masson’s trichrome staining revealed the degree of myocardial fibrosis. Figure 6b showed normal myocardial tissues without obvious trace of collagen fibers in the sham group. The deposition of collagen fibers was marked from the endocardium to the epicardium in the infarction zone, and thinner ventricular walls were observed in the AMI group. In the atorvastatin group, collagen fibers were distributed in the myocardial tissues of the infarction zone, presenting a less extent than in the AMI group. Compared with the AMI group, the losartan group showed less collagen and collagen fibers in the infarction zone and collagen fiber infiltration at the infarct border.

Immunohistochemical staining indicated the expression of notch1, TGF-β, and smad2 in myocardial tissues (Fig. 7a). The relative expression of notch1, TGF-β, and smad2 was significantly upregulated in the AMI group compared with those of the sham group (all p < 0.05) (Fig. 7b). In contrast, notch1, TGF-β, and smad2 showed significant downregulation in the atorvastatin- and losartan-treated groups compared with the AMI group (p < 0.05). No significant differences in the expression of notch1, TGF-β, and smad2 were noticed between the atorvastatin and losartan groups (all p > 0.05).

Collagen, the main component of the extracellular matrix (ECM), forms a network around muscle cell bundles [8]. The two main types of collagen in the heart are collagen I and collagen III [9], which are crucial for maintaining heart structure and assisting muscle contraction [10]. In AMI rats, collagen I and III levels were higher compared to healthy rats. In groups treated with atorvastatin or losartan, collagen levels were lower, which indicated that these drugs could reduce myocardial fibrosis.

Cardiac fibroblasts regulate fibrosis by producing collagen, especially during myocardial fibrosis [11]. After AMI, necrotic cells in the infarction zone trigger fibroblast migration, leading to collagen production and interstitial fibrosis in surrounding tissues. The healing process involves clearing necrotic cells and depositing ECM. Atorvastatin has shown anti-fibrotic effects in AMI and metabolic syndrome [12, 13], inhibiting the proliferation of fibroblasts and myofibroblasts [14]. It also reduces collagen production in cardiac fibroblasts stimulated by angiotensin II or TGF-β1 in a dose-dependent manner [15]. A previous study suggested that the pleiotropic effects of pravastatin on the proliferation of fibroblast were linked to the inhibition of TGF-β expression [16]. Current evidence indicates that statins can directly suppress fibroblast proliferation, migration, myofibroblast differentiation, and ECM metabolism, contributing to the prevention of myocardial remodeling.

Cardiomyocytes and macrophages are capable of producing TGF-β [17]. This study found that AMI activated the myocardial TGF-β-Smad2 signaling pathway, with upregulation of TGF-β and Smad2 protein levels and downregulation of Smad7. TGF-β is crucial in myocardial remodeling and fibrosis, likely due to its roles in promoting fibrosis or cardiac hypertrophy [18, 19]. AMI has been shown to induce upregulation of TGF-β [20], which may be associated with reduced EF in AMI patients [21, 22]. In patients with diabetic mellitus, heart failure after AMI often reflected excessive activation of the pro-fibrotic TGF-β/Smad axis [23]. TGF-β and cytokines (e.g., IL-1β and TNF-α) would induce migration of cardiac fibroblasts [11, 24]. During AMI, TGF-β inhibited inflammatory phagocytes and promoted cardiac myofibroblast differentiation and matrix synthesis via the Smad pathway. This contributed to regulation of the transition from inflammation to the scarring phase of AMI [25].

TGF-β family members are crucial in suppressing inflammation and triggering fibrosis. Inactive TGF-β is stored in the myocardium and quickly activated after injury [26]. The biological activity of TGF-β shows rapid rise in the early stages of AMI [27]. However, high levels of inflammatory mediators reduce cellular response to TGF-β, delaying myofibroblast differentiation and matrix deposition until the removal of necrotic cells and debris [28]. In the presence of inflammation suppression, TGF-β signaling stimulates fibroblast differentiation, promotes the production of collagen and fibronectin, and enhances protease inhibitor synthesis [29].

Cardiac insufficiency, myocardial apoptosis, and myocardial fibrosis are related to the activation of Smads [30]. Inhibition of Smad shows beneficial effects on attenuating myocardial fibrosis and apoptosis [31]. Smad7 overexpression was reported to ameliorate myocardial inflammation and fibrosis [32]. Smad7 treatment can halt myocardial degradation, increase left ventricular mass, and prevent TGF-β/Smad3-mediated myocardial fibrosis [32]. Many synthetic drugs have been reported to mitigate AMI by inhibiting the TGF-β/Smad signal pathway. For example, simvastatin reduces left ventricular remodeling by inhibiting the TGF-β/Smad signal pathway in rats [33]. In our study, atorvastatin treatment downregulated the expression of TGF-β and Smad2 in AMI rats. In the presence of losartan potassium, the expression of TGF-β and Smad2 was downregulated, and the expression of Smad7 was upregulated, which suggested that both atorvastatin and losartan effectively inhibited TGF-β/Smad signal transduction pathway.

Notch pathway regulates embryonic development and differentiation of various cell types. It involved in the proliferation and inflammatory response of smooth muscle cells in coronary atherosclerosis and diabetic mellitus [34]. Notch mutations could affect embryonic development and were responsible for various diseases in adults. For example, Notch1 mutations were associated with cardiomyopathy [35, 36]. Our study indicated that atorvastatin or losartan can inhibit the Notch1 signal pathway, which may serve as a treatment target in AMI.

Notch1 facilitates the differentiation of M1 macrophage and the release of cytokines, causing inflammatory reactions and atherosclerosis. Inhibition of the Notch1 signal pathway promotes the production of anti-inflammatory M2 macrophage and the secretion of anti-inflammatory mediators [37]. Notch pathway is associated with atherosclerosis process. The Notch1 pathway is activated in endothelial cells of atherosclerotic plaques, triggering upregulation of inflammatory factors and adhesion molecules, ultimately leading to an inflammatory response and endothelial cell aging [38]. The pathophysiology of cardiac failure is associated with Notch signaling pathway in the ECM [39]. Recent data have showed that Notch signaling pathway in ventricular muscle cells is activated during cardiac modeling after AMI [40, 41]. To date, the effects of atorvastatin on Notch1 signal pathway in AMI rats have not been reported. In the current study, Notch1 signal pathway was activated in AMI rats, and atorvastatin inhibited Notch1 signal pathway. For the mechanisms, we speculated that statins inhibit the inflammatory responses by directly inhibiting Notch1 signaling, thereby reducing cardiac remodeling. In addition, statins may affect the expression of Smads through regulating the Notch signaling and further regulate the TGF-β/Smad signaling pathways in vascular endothelial cells [42].

In this study, the expression of galectin-3, serving as a marker of myocardial inflammation and cardiac fibrosis, was significantly upregulated in the presence of myocardial fibrosis in AMI rats. Cardiac fibrosis was more severe in rats with high galectin-3 expression developing congestive heart failure within 12–14 weeks [43]. This study found that atorvastatin and losartan significantly downregulated the expression of galectin-3 in myocardium compared with AMI rats. For the mechanism, as a lectin expressed mainly in atherosclerosis, galectin-3 coexisted with macrophage in atherosclerotic plaques. Its expression represents the severity of plaque inflammation. Besides, atorvastatin can significantly decrease the expression of galectin-3 and inhibit macrophage activity in plaques [44]. Indeed, statins have been clinically shown to reduce plasma galectin-3 content in plasma [45].

Echocardiogram examination revealed that the AMI group showed the decline of cardiac functions. The ligation of anterior descending arteries of the rats caused the blockage of blood and anterior myocardial infarction, thereby the anterior wall of the left ventricle was significantly narrowed, and the posterior wall of the left ventricle showed no significant changes. Left ventricular remodeling led to dilatation of the cardiac chambers and a significant reduction in cardiac function, followed by myocardial fibrosis within 3–5 days. The early inflammation was gradually reduced as the expression of anti-inflammatory factors (e.g., TGF-β and IL-10) and apoptosis of inflammatory cells showed significant increase [2]. The pathological process of AMI consists of three stages: the acute inflammatory reaction stage, the proliferation stage, and the late maturation stage [46, 47]. During the proliferative stage, injury healing initiated from the migration of multiple fibroblasts and vascular endothelial cells to the injured area. Activated cardiac fibroblasts are converted into myofibroblasts and produce extracellular collagen and scars [47]. Myocardial fibrosis is the pathological foundation of most cardiac diseases, and excessive ECM synthesis may change cardiac structure and functions, causing degradation of cardiac functions [48]. Our results showed that in the atorvastatin group, myocardial remodeling was ameliorated, ventricular walls were slightly thicker, cardiac chambers became slightly smaller, and cardiac functions were improved. Hemodynamic analysis further showed that the administration of statins improved cardiac functions.

There are indeed some limitations in our study. First, it is still a challenge for us to exclude the possibility that TGF-β and notch sign in AMI. In the future, more studies are required to further illustrate this. Second, the exact mechanism on how atorvastatin modulated the notch1-TGF-β-smads signal pathways is still unclear. On this basis, more experimental studies are required.

In summary, atorvastatin involved in inhibiting the myocardial fibrosis via the notch1-TGF-β-smads signal pathways. It could mitigate myocardial remodeling and improve cardiac function in an effective manner.

This study protocol was reviewed and approved by the Institutional Animal Care and Use Committee, Approval No. IRB2022-DW-13.

The authors have no conflicts of interest to declare.

This study was not supported by any sponsor or funder.

Qi Kang: conceptualization, methodology, writing – original draft, and writing – review and editing. Mei Kang and Taniya Fernando: formal analysis and writing – review and editing. Mengyun Yang: investigation and writing – review and editing.

All data generated or analyzed during this study are included in this article. Further inquiries can be directed to the corresponding author.