MMP9 inhibition increases autophagic flux in chronic heart failure

Shyam S Nandi; Kenichi Katsurada; Neeru M Sharma; Daniel R Anderson; Sushil K Mahata; Kaushik P Patel

doi:10.1152/ajpheart.00032.2020

. 2020 Oct 16;319(6):H1414–H1437. doi: 10.1152/ajpheart.00032.2020

MMP9 inhibition increases autophagic flux in chronic heart failure

Shyam S Nandi ^1,^✉, Kenichi Katsurada ¹, Neeru M Sharma ¹, Daniel R Anderson ², Sushil K Mahata ^3,⁴, Kaushik P Patel ¹

PMCID: PMC7792705 PMID: 33064567

Abstract

Increased matrix metalloprotease 9 (MMP9) after myocardial infarction (MI) exacerbates ischemia-induced chronic heart failure (CHF). Autophagy is cardioprotective during CHF; however, whether increased MMP9 suppresses autophagic activity in CHF is unknown. This study aimed to determine whether increased MMP9 suppressed autophagic flux and MMP9 inhibition increased autophagic flux in the heart of rats with post-MI CHF. Sprague-Dawley rats underwent either sham surgery or coronary artery ligation 6–8 wk before being treated with MMP9 inhibitor for 7 days, followed by cardiac autophagic flux measurement with lysosomal inhibitor bafilomycin A1. Furthermore, autophagic flux was measured in vitro by treating H9c2 cardiomyocytes with two independent pharmacological MMP9 inhibitors, salvianolic acid B (SalB) and MMP9 inhibitor-I, and CRISPR/cas9-mediated MMP9 genetic ablation. CHF rats showed cardiac infarct, significantly increased left ventricular end-diastolic pressure (LVEDP), and increased MMP9 activity and fibrosis in the peri-infarct areas of left ventricular myocardium. Measurement of the autophagic markers LC3B-II and p62 with lysosomal inhibition showed decreased autophagic flux in the peri-infarct myocardium. Treatment with SalB for 7 days in CHF rats decreased MMP9 activity and cardiac fibrosis but increased autophagic flux in the peri-infarct myocardium. As an in vitro corollary study, measurement of autophagic flux in H9c2 cardiomyocytes and fibroblasts showed that pharmacological inhibition or genetic ablation of MMP9 upregulates autophagic flux. These data are consistent with our observations that MMP9 inhibition upregulates autophagic flux in the heart of rats with CHF. In conclusion, the results in this study suggest that the beneficial outcome of MMP9 inhibition in pathological cardiac remodeling is in part mediated by improved autophagic flux.

NEW & NOTEWORTHY This study elucidates that the improved cardiac extracellular matrix (ECM) remodeling and cardioprotective effect of matrix metalloprotease 9 (MMP9) inhibition in chronic heart failure (CHF) are via increased autophagic flux. Autophagy is cardioprotective; however, the mechanism of autophagy suppression in CHF is unknown. We for the first time demonstrated here that increased MMP9 suppressed cardiac autophagy and ablation of MMP9 increased cardiac autophagic flux in CHF rats. Restoring the physiological level of autophagy in the failing heart is a challenge, and our study addressed this challenge. The novelty and highlights of this report are as follows: 1) MMP9 regulates cardiomyocyte and fibroblast autophagy, 2) MMP9 inhibition protects CHF after myocardial infarction (MI) via increased cardiac autophagic flux, 3) MMP9 inhibition increased cardiac autophagy via activation of AMP-activated protein kinase (AMPK)α, Beclin-1, Atg7 pathway and suppressed mechanistic target of rapamycin (mTOR) pathway.

Keywords: autophagic flux, chronic heart failure, matrix metalloprotease 9, myocardial infarction

INTRODUCTION

Matrix metalloproteinase (MMP)9, a Zn²⁺-dependent endopeptidase enzyme gelatinase, is increased during post-myocardial infarction (MI) pathological cardiac remodeling (93). MMP9 is also known as gelatinase B, one of the most extensively studied MMPs in cardiovascular research (55, 78). MMP9 has the capacity to degrade both elastin and collagen, as these substrates are major components of cardiac extracellular matrix (ECM) protein. The term “post-MI-induced chronic heart failure (CHF)” in this study is referred to as a chronic and progressive pathological condition of ischemic left ventricular myocardium with infarction that causes the heart to pump inefficiently. Among the MMP family members, MMP9 plays a major role in myocardial infarction, heart failure, diabetes, hypertension, and atherosclerosis (16, 33, 41). Previous studies have shown that MMP9 inhibition can improve cardiac remodeling, fibrosis, infarct size, infarct expansion rate, and survival after post-MI induced heart failure or during ischemia-reperfusion injury, which leads to improved cardiac function (10, 28, 32, 33, 66, 67, 82, 94, 124). However, the cardioprotective component or specific mechanism of MMP9 inhibition remains to be identified. MMP9 expression has been reported to correlate directly with diastolic dysfunction and is reported to be a biomarker for cardiac remodeling in patients with heart failure (4, 25, 110). In addition, targeted deletion of MMP9 protects against ischemia-reperfusion injury (94), suggesting that MMP9 possibly plays a fundamental role in cardiac damage. Furthermore, pharmacological MMP9 inhibitors or monoclonal antibodies targeting MMP9 (andecaliximab, GS-5745) are currently being used in Phase I and II clinical trials (18, 99, 105).

Under normal physiological conditions, the gene and protein expression levels of MMP9 are low; however, MMP9 expression and activity are robustly increased under pathological conditions of the heart observed during multiple cardiovascular diseases (31, 66, 70, 85). Several known agents that elevate endogenous cardiac MMP9 level in post-MI myocardium are hypoxia, reactive oxygen species (ROS), growth factors, chemokines, ECM-activated proteins, and proinflammatory cytokines (111, 113). In addition, MMP9 is recruited to the infarcted myocardium via cardiac nonresident cell infiltration (neutrophil, macrophage, and leukocyte infiltration) during post-MI remodeling (55).

The collective changes in the myocardial arrangement and function after MI are termed pathological left ventricular remodeling (67, 76). The extracellular role of MMP9 is extensively studied; however, its intracellular or regulatory role in the intracellular signaling pathways remains unknown. We and others previously reported that the ablation of MMP9 expression and activity improved cardiomyocyte contractility in cardiomyopathy (28, 55, 66, 67, 85, 94). However, the mechanism for the MMP9 ablation-mediated cardioprotection is not entirely clear.

Autophagy is an evolutionarily conserved intracellular catabolic process that degrades or recycles defective cellular components and cytosolic protein aggregates and plays a fundamental role in cardiovascular physiology (24, 34, 37, 46, 49, 69, 70, 76, 86, 95, 109, 130, 132). Interestingly, autophagy is reported as a protective mechanism during myocardial ischemia (24, 53, 60, 101). Autophagic flux is decreased during ischemic damage (9, 21) or adaptively insufficient to clear the cellular debris of ischemic damage (51, 68, 101, 122). Furthermore, the uncontrolled activation of the pathway that leads to autophagy inhibition causes a reduction of myocardial tolerance to ischemia, which supports the contribution of the cardioprotective role of autophagy during post-MI cardiac remodeling (102). On the other hand, excessively high or more severe and prolonged autophagic activity leads to increased cell death and cardiomyocyte apoptosis, which leads to decline in myocardial function (48, 92). However, the role of MMP9 in the regulation of cardiac autophagic flux remains to be elucidated. In this study, we conducted a series of in vivo and in vitro experiments designed to determine whether the cardioprotective effects of MMP9 ablation are in part contributing to increased cardiac autophagic flux. We used pharmacological and genetic tools to manipulate MMP9 inhibition. Western blotting, zymography, and microscopy were used to validate that MMP9 activity was inhibited in the heart/cardiomyocytes and in fibroblasts with a concomitant increase in autophagic flux.

MATERIALS AND METHODS

Animals.

We used healthy male Sprague-Dawley rats (age 6–8 wk) weighing 220–240 g from the Jackson Laboratory that were housed in the animal facility of the University of Nebraska Medical Center. Rats were kept in hygienic cages with maintained room temperature of 22–24°C, humidity of 30–40%, and 12-h:12-h dark-light cycle, and food and water were available ad libitum. All experimental protocols were approved by the Institutional Animal Care and Use Committee of the University of Nebraska Medical Center, and all methods were conducted in accordance with the relevant guidelines and regulations of our institution, the American Physiological Society, and the National Institutes of Health Guide for the Care and Use of Laboratory Animals.

Hemodynamic measurements.

At the end of the experiments hemodynamic parameters were measured with a microtipped Millar catheter (SPR-838; Millar Instruments, Houston, TX) (38, 74). The catheter was inserted via the right carotid artery into the left ventricle (LV) as described previously (74, 80, 128). The mean arterial pressure (MAP) and heart rate (HR) were simultaneously recorded by this catheter before the LV was entered in a PowerLab data acquisition system (8SP; ADInstruments).

Chronic heart failure model.

CHF was induced by ligation of the left anterior descending coronary artery, which has been described previously and is routinely used in our laboratory (80, 106, 107, 128, 129). The rats were randomly assigned to two groups, a sham-operated (Sham) control group and a CHF group. Rats were subjected to ligation surgery with the use of a rodent ventilator with 2–3% isoflurane at a rate of 60 breaths/min during the surgical procedure. The left anterior descending coronary artery was permanently ligated with a silk suture, and infarction was confirmed by the pale color of the ventricular myocardium. The sham-operated control group underwent all the steps of the surgical procedure except for the actual ligation step. Rats were housed in a clean facility after postsurgery recovery for 6–8 wk. This extended period mimics the clinical condition of chronic heart failure in patients with heart failure. At the end of the study, cardiac hemodynamic parameters were assessed during nonsurvival surgery performed on anesthetized rats with a single injection of urethane (0.75 g/kg ip)-chloralose (70 mg/kg ip) mixture, followed by euthanasia with an overdose of urethane and chloralose.

In this study, by assessing left ventricular dysfunction with anatomic and hemodynamic criteria we randomly selected animals with confirmed myocardial ischemia, cardiac damage, and dysfunction. Infarct size was measured from exposed left ventricle imaging (Fig. 1B) using the ImageJ tool. Those rats with a significant infarct size (>35% of total left ventricle wall) and elevated left ventricular end-diastolic pressure (LVEDP, >15 mmHg) with significant reductions in maximum ±dP/dt (±dP/dt_max) were considered to be in the CHF condition. The peri-infarct tissues (Fig. 1B) are considered as the area at risk and were used in the present study. These characteristic features of the CHF model are summarized in Tables 1 and 2.

Fig. 1. — Post-myocardial infarction (MI) in rat induces chronic heart failure (CHF). A: schematic showing ischemic myocardium and 3 district zones of cardiac remodeling. B: representative exposed left ventricle showing 3 distinct zones of remodeling, infarct zone, peri-infarct zone, and remote zone in the post-MI heart. Scale bar, 1 cm. Sham, sham operated. C: traces showing left ventricular pressure in Sham and CHF rats. D: traces showing ±dP/dt in Sham and CHF rats. E, *top*: representative Western blot showing hypoxia-inducible factor (HIF)-1α level in the peri-infarct zone. *Bottom*: quantification of HIF-1α normalized to actin. F: correlation plot showing HIF-1α level with %infarct size in Sham and CHF rats. Values are means ± SE; n = 6 rats. RAU, relative arbitrary unit. *P < 0.05 vs. Sham. Student’s t test was used to generate P values.

Table 1.

Characteristics of Sham and CHF rats

	Sham	CHF
n	6	6
Body weight, g	338 ± 9	368 ± 11
Heart weight, g	1.0 ± 0.1	1.5 ± 0.1^*
HW/BW, ×1,000	3.1 ± 0.1	4.1 ± 0.3^*
Infarct size, %endocardial LV	0	41 ± 2^*
LVEDP, mmHg	2.2 ± 0.4	26.3 ± 4.2^*
+dP/dt, mmHg/s	6,644 ± 416	4,465 ± 161^*
−dP/dt, mmHg/s	−6,141 ± 415	−3,841 ± 86^*
LVEDD, mm	8.3 ± 0.3	10.6 ± 0.3^*
LVESD, mm	5.2 ± 0.3	9.0 ± 0.1^*
Ejection fraction, %	64 ± 1	34 ± 2^*
Fractional shortening, %	38 ± 1	18 ± 2^*
Basal MAP, mmHg	98 ± 2	91 ± 4
Basal HR, beats/min	330 ± 14	336 ± 12

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Values are means ± SE for n rats. BW, body weight; CHF, chronic heart failure; HR, heart rate; HW, heart weight; LV, left ventricle; LVEDD, left ventricular end-diastolic dimension; LVEDP, left ventricular end-diastolic pressure; LVESD, left ventricular end-systolic dimension; MAP, mean arterial pressure; Sham, sham operated.

P < 0.05 compared with Sham.

Table 2.

Characteristics of Sham and CHF rats treated with SalB or saline

	Sham+Saline	CHF+Saline	Sham+SalB	CHF+SalB
n	6	6	6	6
Body weight, g	344 ± 9	377 ± 14	368 ± 8	364 ± 13
Heart weight, g	1.0 ± 0.1	1.4 ± 0.1^*	1.1 ± 0.1	1.5 ± 0.1^**
HW/BW, ×1,000	3.1 ± 0.1	4.1 ± 0.2^*	3.1 ± 0.1	4.2 ± 0.2^**
Infarct size, %endocardial LV	0	40 ± 3^*	0	35 ± 2^**
LVEDP, mmHg	2.1 ± 0.4	25.3 ± 5.0^*	2.1 ± 0.3	24.1 ± 4.0^**
+dP/dt, mmHg/s	7,157 ± 237	4,453 ± 92^*	6,667 ± 391	4,349 ± 169^**
−dP/dt, mmHg/s	−6,258 ± 348	−4,003 ± 129^*	−5,975 ± 506	−4,202 ± 205^**
Basal MAP, mmHg	92 ± 2	94 ± 3	95 ± 3	92 ± 4
Basal HR, beats/min	332 ± 4	335 ± 9	335 ± 13	341 ± 12

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Values are means ± SE for n rats. BW, body weight; CHF, chronic heart failure; HR, heart rate; HW, heart weight; LV, left ventricle; LVEDP, left ventricular end-diastolic pressure; MAP, mean arterial pressure; SalB, salvianolic acid B; Sham, sham operated. Two-way ANOVA indicated a significant effect of CHF but not of SalB treatment. There was no interaction term, indicating that SalB did not affect Sham and CHF groups differently. There were no significant differences between the 2 CHF groups.

P < 0.05 compared with Sham;

^**

P < 0.05 compared with Sham+SalB.

MMP9 inhibition in vivo.

For MMP9 inhibition in vivo, rats were treated with salvianolic acid B (SalB; Cat. No. 40479, Astatech; 10 mg/kg ip) dissolved in isotonic saline for 7 days 6–8 wk after sham or post-MI surgery. Vehicle-treated animals received an intraperitoneal injection of isotonic saline.

Transmission electron microscopy.

The heart samples were processed for transmission electron microscopy (TEM) as described previously (62, 79, 97). Briefly, heart slices were immersion fixed with 2.5% glutaraldehyde and 2% paraformaldehyde in 0.15 M cacodylate buffer. The tissue slices were postfixed in 1% OsO₄ in 0.1 M cacodylate buffer for 1 h on ice, followed by en bloc staining (electron staining techniques) with 2–3% uranyl acetate for 1 h on ice. The tissue slices were dehydrated in graded series of ethanol (20–100%) on ice, followed by one wash with 100% ethanol and two washes with acetone (15 min each), and embedded with Durcupan. Ultrathin (∼50–60 nm) sections were cut on a Leica UCT ultramicrotome and picked up on formvar- and carbon-coated copper grids. Sections were stained with 2% uranyl acetate for 5 min and Sato’s lead stain for 1 min. Grids were viewed with a JEOL JEM1400-plus Transmission Electron Microscope (JEOL, Peabody, MA) and photographed with a Gatan OneView digital camera with 4k × 4k resolution (Gatan, Pleasanton, CA). NIH ImageJ was used to calculate the area by manually tracing around the autophagosomes, and the number of autophagosomes was also calculated as per a previous publication from our laboratory (62). Frequency distributions of autophagosome area were plotted in accordance with our previous publications (62, 79, 97, 120). These quantifications were done with blinded samples.

H9c2 cardiomyocyte culture.

The H9c2 rat cardiomyocytes were cultured in vitro according to the standard protocol as provided by the distributor, ATCC. The cell line was cultured and maintained with high-glucose DMEM (Cat. No. D5796, Sigma) supplemented with 10% FBS (Cat. No. F6178, Sigma) and 1% penicillin- streptomycin in a cell culture incubator (800-WJ; Thermoelectric Corporation) at 37°C with 5% CO₂. In brief, cardiomyocytes were cultured in T75 plates, seeded on six-well plates, and allowed to differentiate in the presence of 1 µM retinoic acid (RA) and 1% FBS (catalog F2442, Sigma) for 7 days. Differentiated cells were treated as per the applicable assay, and after completion of treatment cells were trypsinized to isolate samples for Western blot measurements or processed for immunocytochemical assays. The H9c2 cell line was maintained with regular DMEM and subcultured at a ratio of 1:3 at 70–80% confluence. Fresh medium was changed every 3 days, and the cardiomyocytes of passages 7–8 were used for differentiation and utilized in subsequent experiments. Cells were collected at 1,000 rpm centrifugation for 5 min with 37°C prewarmed trypsin at room temperature.

H9c2 cardiomyocyte treatment.

Once the cells were ∼80% confluent, differentiation was initiated by switching to DMEM culture medium supplemented with 1% FBS and daily administration of 1 μM retinoic acid (RA; Cat. No. R2625, Sigma). RA was dissolved quickly in DMSO (Cat. No. D2650, Sigma) to avoid air exposure and stored immediately in light-protected vials at −80°C. Multiple concentrations of RA were tested, and a 1 μM dose for 7 days was chosen as the most effective dose for H9c2 differentiation. The culture medium was changed every day with freshly prepared differentiation medium. After differentiation, cells were divided into three groups for treatment: 1) isotonic saline, 2) SalB (50 µM; Cat. No. 40479, Astatech), and 1) MMP-9 Inhibitor-I (5 nM; Cat. No. 1177749-58-4, Calbiochem) for 24 h. To block autophagosome formation, cells were treated with LY294002 (20 µM; Cat. No. L9908, Sigma) for 24 h. To induce cell death signaling, H9c2 cardiomyocytes were cotreated with 200 µM H₂O₂ (Cat. No. 31642, Sigma-Aldrich) for 24 h. For assessment of autophagic flux in vitro, cells were treated with 100 nM bafilomycin A1 (BafA1; Cat. No. 11038, Cayman) for the last 2 h of treatment before trypsinization and cell collection or cell fixation. Control groups of cells were treated with isotonic saline or DMSO alone, as applicable.

Mouse embryonic fibroblast culture.

Mouse embryonic fibroblasts (MEFs) were cultured in vitro according to the standard protocol. Fibroblasts were cultured and maintained in standard DMEM supplemented with 15% FBS (Cat. No. F6178, Sigma) and 1% penicillin-streptomycin in a cell culture incubator (800-WJ; Thermoelectric Corporation) at 37°C with 5% CO₂. The cell line was maintained with regular DMEM medium and subcultured in a ratio of 1:3 at 90% confluence. For autophagic flux measurement, fibroblasts were treated with 100 nM bafilomycin A1 (BafA1; Cat. No. 11038, Cayman) 2 h before trypsinization and cell collection or cell fixation. The control group of cells were treated with DMSO alone.

Western immunoblotting.

A standard Western immunoblotting protocol was performed to assess the level of protein expression (39, 71). Whole cell or tissue protein lysates was estimated by the Pierce BCA Protein Assay Kit (Cat. No. 23227, Pierce). An equal amount of protein lysate (30 μg/lane) was boiled in 4× denaturing Laemmli Sample Buffer (Cat. No. 161-0747, Bio-Rad Laboratories) and loaded on 10% or 15% SDS-PAGE gel as applicable for the molecular weight of the target protein. After electrophoresis, gels were transferred onto a polyvinylidene fluoride (PVDF) membrane. Transferred PVDF membranes were incubated with 5% nonfat dried milk (Cat. No. 1706404, Bio-Rad) in TBS for 1 h at room temperature. Primary antibodies were diluted to 1:1,000 in TBS and incubated overnight at 4°C. The primary antibodies used were LC3B (Cat. No. ab192890), SQSTM1/p62 (Cat. No. ab56416), Lamp1 (Cat. No. ab24170), hypoxia-inducible factor (HIF)-1α (Cat. No. ab1), and total oxidative phosphorylation (OXPHOS) rodent WB antibody cocktail (Cat. No. ab110413), which were obtained from Abcam (United States). MMP9 (Cat. No. sc-13520) and beta-actin (Cat. No. sc-47778, sc-47778-HRP) were purchased from Santa Cruz Biotechnology (United States). Phospho- (p-)AMP-activated protein kinase (AMPK)α^Thr172 (Cat. No. 2535S), AMPKα (Cat. No. 2532S), p-mechanistic target of rapamycin (mTOR)^Ser2448 (Cat. No. 2971S), mTOR (Cat. No. 2983S), Beclin-1 (Cat. No. 3495S), LC3A (Cat. No. 4599), Atg7 (Cat. No. 8558S), and caspase 3 (Cat. No. 9662) were obtained from Cell Signaling Technology (United States). All primary antibodies were incubated overnight at 4°C. All respective secondary antibodies with HRP conjugates were diluted at 1:4,000 in TBS and incubated at room temperature for 2 h. Restripping and reprobing of blots were performed with Western Blot Stripping Buffer (Cat. No. 46430, Thermo Scientific, Inc.) when/if applicable. The blots were developed with SuperSignal West Femto Stable peroxidase buffer (Cat. No. 1859023, Thermo Scientific) and the Molecular Imager ChemiDocXRS imaging system (Bio-Rad Laboratories). Multiple images were captured with Image Laboratory software version 6 (Bio-Rad Laboratories) at lowest to highest signal accumulation mode. The bands detected with optimal exposure of time were considered as within the linear range and selected for quantification. Quantifications are normalized from loading controls.

In-gel gelatin zymography.

In-gel gelatin zymography assay was performed with our previously published standard protocol (6). In brief, 10% SDS-PAGE gels with 0.1% gelatin substrate were prepared. Left ventricular heart tissues or cardiomyocytes were lysed in RIPA buffer (Cat. No. BP-115, Boston BioProducts) with no protease inhibitor. Sample quantification was performed with the Pierce BCA Protein Assay Kit (Cat. No. 23227, Pierce). An equal number of samples (30 µg/lane) was prepared with 2× sample buffer (62.5 mM Tris, 4% SDS, 25% glycerol, 0.01% bromophenol blue, pH 6.8). “Zymo” samples were mixed with 2× sample buffer, incubated at room temperature for 10 min, and then loaded into wells. The Zymo-gel electrophoresis was performed at 70 V with Tris-glycine-SDS running buffer for 4 h. After electrophoresis, the resolving gel was incubated in renaturation buffer [Triton X-100 2.5% (vol/vol) in distilled water] with gentle shaking for 45 min. Next, the gel was washed with distilled water and subjected to incubation with developing buffer (0.5 M Tris, 2 M NaCl, 50 mM CaCl₂, 0.2% Brij 35, pH 7.5) at 37°C with 50 rpm shaking for 48 h. Finally, the gel was stained with Coomassie brilliant blue R-250 dye [0.5% (wt/vol) in methanol; Cat. No. 20278, Thermo Scientific] for 30 min and washed with the destaining solution (30% methanol and 10% acetic acids) to remove the excess stain. The gels were scanned with the Molecular Imager ChemiDoc XRS imaging system on a white plate (Bio-Rad Laboratories). The images were captured with Image Laboratory software version 6 (Bio-Rad Laboratories). Quantifications are normalized from loading controls.

Immunocytochemistry.

Immunocytochemistry was performed on cultured rat H9c2 cardiomyocytes in T24 well plates. Cells were washed in phosphate-buffered saline (PBS, pH-7.4), after treatment, followed by being fixed in 4% paraformaldehyde solution (Cat. No. 158127, Sigma) for 30 min. Fixed cells were permeabilized in 0.02% (vol/vol) Triton X-100 solution (Cat. No. BP151-500, Fisher Bioreagents) in PBS for 30 min and blocked in 1% BSA solution for 1 h at room temperature. In the next step, cells were washed and incubated in diluted primary antibody solution (in TBS with 0.2% BSA) at 4°C overnight. The primary antibodies used were 1:200 dilution of LC3B (Cat. No. ab192890, Abcam), and fibronectin (Cat. No. ab23750, Abcam). The next day, cells were washed in PBS and incubated with 1:400 dilution (in TBS with 0.2% BSA) of anti-rabbit Alexa Fluor 488-conjugated secondary antibody (Cat. No. A11008, Life Technologies). Alexa Fluor 555 Phalloidin (Cat. No. A34055, Life Technologies) was used to counterstain the F actin filaments. Nuclei were counterstained with 1 µg/mL DAPI in PBS (Cat. No. A1001, AppliChem) and mounted with Fluoromount-G mounting medium (Cat. No. 0100-01, Southern Biotech). Images were captured by Olympus IX71 Imaging Systems (Olympus) and analyzed by ImageJ software (NIH).

Measurement of in vivo autophagic flux.

For assessment of autophagic flux in in vivo, sham-operated control and CHF rats were injected with BafA1 (6 μmol/kg body wt ip, dissolved in DMSO; Cat. No. 11038, Cayman) 2 h before euthanasia. Control groups were injected with DMSO alone (ip). Rats were euthanized 2 h after an injection of either BafA1 or DMSO, and hearts were collected. Left ventricular tissues were processed for whole protein lysate preparation in RIPA lysis buffer. The LC3B-II/actin level was measured by Western immunoblotting technique, which represents the underlying level of autophagic flux compared among groups with BafA1 versus no BafA1 (i.e., DMSO alone) treatment.

Measurement of peri-infarct fibrosis.

To quantify the peri-infarct fibrosis in CHF rats, we used Masson’s trichrome staining of collagen fibers. Area-matched left ventricular tissues from Sham and CHF rats were fixed in 10% formalin solution followed by dehydration using graded ethanol and embedded in a paraffin block. Paraffin-embedded blocks were sectioned into 5-µm transverse sections and processed for a standard Masson’s trichrome staining protocol (73). Bright-field colored images were captured with a bright-field microscope (Leica Microsystems, Buffalo Grove, IL) with Image Pro 7.0 software. Blue color represents collagen fiber staining, and red color represents the myocardium. Blue color intensity was measured by ImageJ software (Fiji version, NIH). The Tissue Core Facility of the University of Nebraska Medical Center was used for Masson’s trichrome staining.

MMP9 genetic ablation in H9c2 cardiomyocytes by CRISPR/cas9 technique.

MMP9 gene was deleted in H9c2 cardiomyocytes by transfecting rat MMP9 CRISPR/Cas9-knockout (KO) and MMP9-homology-directed repair (HDR) plasmids (Cat. No. sc-437355, Santa Cruz Biotechnology). In brief, cells were cotransfected with these KO plasmids [with green fluorescent protein (GFP) reporter] and HDR plasmids [with red fluorescent protein (RFP) reporter and puromycin selection marker] with Lipofectamine 3000 (Cat. No. L3000015, Life Technologies) and Opti-MEM (Cat. No. 31985070, Life Technologies). After 48 h of cotransfection, cells were selected with puromycin solution (10 mg/mL; Cat. No. P8833, Sigma) for 24 h. Puromycin treatment selects RFP H9c2 cells (both MMP9^+/null and MMP9^null/null). MMP9 knockdown in the RFP H9c2 cells was validated by Western immunoblot and in-gel gelatin zymography techniques. We represent these cells as knockdown MMP9^+/− H9c2 cardiomyocytes.

MMP9 genetic ablation in fibroblasts by CRISPR/cas9 technique.

MMP9 gene was deleted in mouse embryonic fibroblasts (MEFs) by transfecting mouse MMP-9 CRISPR/Cas9 KO and MMP9-HDR plasmids (Cat. No. sc-421679, Santa Cruz Biotechnology). In brief, cells were cotransfected with KO plasmids (with GFP reporter) and HDR plasmids (with RFP reporter and puromycin selection marker) with Lipofectamine 3000 (Cat. No. L3000015, Life Technologies) and Opti-MEM (Cat. No. 31985070, Life Technologies). After 48 h of cotransfection, cells were selected with puromycin solution (10 mg/mL; Cat. No. P8833, Sigma) for 24 h. Puromycin treatment selects RFP MEF cells (both MMP9^+/null and MMP9^null/null). MMP9 knockdown in the RFP MEFs was validated by in-gel gelatin zymography techniques. We represent these cells as knockdown MMP9^+/− MEFs.

Measurement of autophagic flux by tandem sensor RFP-GFP-LC3B assay.

To measure autophagy flux, the Premo Autophagy Tandem Sensor RFP-GFP-LC3B Kit (Cat. No. P36239) was used. This dual fluorescent reporter clone is designed to monitor autophagosomes (APs) as yellow (GFP+RFP) puncta and autolysosomes (ALs) as free red (RFP) puncta in vitro by transfection. H9c2 cardiomyocytes or fibroblasts were transduced with the LC3B BacMam 2.0 reagents and then treated with MMP-9 Inhibitor-I (5 nM; Cat. No. 1177749-58-4, Calbiochem) for 24 h. After 24 h of treatment, fluorescence imaging was performed. The numbers of ALs and APs were counted per transfected cell in each treatment group and quantified.

Statistical analysis.

Data are presented as means ± SE. Differences between groups were determined by one- or two-way ANOVA, followed by the Tukey’s multiple comparisons test for post hoc analysis if there was a significant interaction, with Prism7.0 Graph Pad software. Differences of means between two groups were analyzed with Student’s t test. Two-way ANOVA with Sidak’s multiple comparison test was used for electron micrograph analyses. P < 0.05 was considered statistically significant.

RESULTS

Characterization of myocardial infarction induced chronic heart failure in rats.

Table 1 illustrates the characteristic morphological and hemodynamic parameters for the sham-operated (Sham) control and CHF rats. The hearts of the CHF group had 41 ± 2% of the endocardial surface area infarcted, whereas the Sham rats had none. LVEDP was significantly elevated in the CHF group compared with the Sham group (26.3 ± 4.2 vs. 2.2 ± 0.4 mmHg) (Fig. 1C and Table 1). We measured HIF-1α protein stabilization in the hypoxic peri-infarct myocardium in CHF rats. Our data showed increased HIF-1α protein level within the infarct zone, and there is a strong correlation of HIF-1α protein level with the percent infarct size (Fig. 1, E and F). ±dP/dt_max was significantly decreased in the CHF group compared with the Sham group, indicating reduced contractility (Fig. 1D and Table 1). Taken together, percent infarct size, increase in LVEDP, and a decrease in ±dP/dt_max indicate that rats in the CHF group were experiencing significant cardiac dysfunction.

Increased MMP9 activity contributes to pathological cardiac remodeling in post-MI induced CHF.

Our results showed that MMP9 expression and activity are significantly increased in the peri-infarct areas of the LV at 6–8 wk after MI (Fig. 2). Moreover, the peri-infarct areas showed comparatively higher MMP9 activity compared with remote areas, suggesting that MMP9 predominantly exacerbated LV remodeling in the adjacent border areas of the infarct but less in remote areas when induced after MI. In a comparable matched area of LV tissue from Sham rats, the endogenous activity of MMP9 was detected as relatively low (Fig. 2, A and B). This suggests that MMP9 is activated during pathological LV remodeling, whereas in normal LV its activity is comparatively very low. Furthermore, the peri-infarct areas showed higher MMP2 activity compared with Sham LV, suggesting that MMP2 activation also contributes to pathological ECM remodeling in the peri-infarct area during post-MI induced CHF (Fig. 2C).

Suppressed cardiac autophagic flux in the peri-infarct area of post-MI induced CHF.

To test whether autophagy is involved in this cardioprotection, downstream of MMP9 inhibition, we investigated the effect of MMP9 inhibition on cardiac autophagic flux at 6–8 wk after MI. We measured cardiac autophagic flux in the peri-infarct areas (the most active remodeling zone in post-MI hearts) of the left ventricle, in the presence and absence of the lysosomal inhibitor bafilomycin A1 (BafA1; 2 h before euthanasia). We quantified the level of Lamp1, LC3B-II, and p62 protein levels in peri-infarct areas of the LV from Sham and CHF rats treated with BafA1 or DMSO as solvent control (Fig. 3). LC3B-II and p62 are markers of the autophagosome and accumulate in the Western blot when BafA1 blocks lysosomal degradation. The level of LC3B-II/actin level accumulation in the presence of BafA1 represents the underlying level of autophagic degradation compared among groups. Our results demonstrated that cardiac LC3B-II/actin level is accumulated with 2-h BafA1 treatment in sham-operated control rats, indicating a basal housekeeping level of autophagic degradation in Sham rats as anticipated (Fig. 3D). However, such LC3B-II accumulation was not observed in CHF rats, suggesting that the autophagic degradation is suppressed or adaptively insufficient in the LV of rats with CHF (Fig. 3D). This result suggests that autophagosomes are not at their terminal autolysosomal degradation process, perhaps because of suppressed autophagic degradation signal in CHF. In addition, our Lamp1 and p62 data (Fig. 3, B and C) indicate that autophagosome or autolysosome formation and their abundance are high in CHF peri-infarct areas, which further supports the notion that their terminal degradation is incomplete, which caused a high abundance. Furthermore, in remote areas of the LV our results showed a trend of reduced autophagic activity. However, in our subsequent experiments, we included the peri-infarct tissues for further investigation, as peri-infarct area is the most active area of remodeling in the post-MI heart. We used peri-infarct tissue slices dissected out precisely, without any part of the infarct tissue of necrotic myocardium.

Fig. 3. — Autophagic flux is suppressed in peri-infarct areas during post-myocardial infarction (MI) induced chronic heart failure (CHF). Autophagic flux was evaluated in the sham-operated (Sham) or peri-infarct areas with bafilomycin A1 (BafA1) treatment in each group of rats (Sham and CHF). BafA1 (6 μmol/kg body wt ip) was injected 2 h before euthanasia. A: representative Western blot images showing expression of Lamp1, p62, and LC3B-II in the peri-infarct areas of left ventricular myocardium 6–8 wk after MI. *B–D*: bar graphs showing quantification of Lamp1 (B), p62 (C), and LC3B-II (D) normalized to actin in the area-matched Sham or peri-infarct myocardium. The data were analyzed by 2-way ANOVA followed by Tukey’s multiple comparison test. Values are means ± SE; n = 6 rats. RAU, relative arbitrary unit. *P < 0.05 vs. Sham+BafA1; #P < 0.05 vs. Sham.

Transmission electron microscopy micrographs showed increased autophagosomes and mitochondrial injury in the peri-infarct area of post-MI induced CHF.

To visualize the autophagic vacuoles, we performed transmission electron microscopy (TEM). TEM revealed increased autophagic vacuole abundance and diminished degradation in the LV of the CHF group. TEM micrographs showed the presence of autophagosomes in Sham, Sham+BafA1, CHF, and CHF+BafA1 groups (Fig. 4, A–D). Smaller autophagosomes (up to 0.25 µm²) were preponderant in the Sham-operated group compared with the Sham+BafA1 CHF group (Fig. 4, A and B and E and F). The CHF group exhibited both the increased area (up to 4.5 µm²) and number (16 vs. 12 autophagosomes/58.83 µm² area) of autophagosomes compared with the Sham-operated group (Fig. 4, C and D and F and H). The CHF+BafA1-treated group showed an increased number of smaller autophagosomes (up to 0.5 µm²) and a decreased number of larger (from 0.5 to 1 µm²) autophagosomes (Fig. 4, D and G). Autophagosome numbers were comparable between CHF and CHF+BafA1 groups (Fig. 4H). TEM data demonstrated that at 6–8 wk after MI surgery the LV peri-infarct tissue was apparent, with a significantly increased abundance in the number of autophagic vacuoles, indicating an inhibited terminal degradation of autophagic vacuoles in the CHF group (Fig. 4C) that increased autophagosome abundance.

Fig. 4. — Transmission electron microscopy (TEM) micrographs show increased autophagic vacuoles and mitochondrial damage in post-myocardial infarction (MI) induced chronic heart failure (CHF). *A–D*: TEM micrographs showing autophagosomes and subcellular structures in rat heart. Autophagosomes were evaluated in the area-matched sham-operated (Sham) left ventricle (LV) or peri-infarct area with short-term bafilomycin A1 (BafA1) treatment in each group of rats (Sham and CHF). BafA1 (6 μmol/kg body wt ip) was injected intraperitoneally 2 h before euthanasia, and tissues were processed for ultrastructural evaluation. Representative micrographs are shown for Sham (A) Sham+BafA1-treated (B), CHF-induced (C), and CHF+BafA1-treated (D) rats (n = 3/group). Scale bars, 2 µm. Red arrows show autophagosomes; blue arrows point to Z-disk. Mc, mitochondria. *E–G*: morphometric measurements showing the frequency distribution of autophagosome area calculated from 3 rats in each group: sham-operated (n = 224 autophagosomes) vs. Sham+BafA1-treated (n = 163 autophagosomes) groups (E); sham-operated (n = 224 autophagosomes) vs. CHF-induced (n = 238 autophagosomes) groups (F); CHF-induced (n = 238 autophagosomes) vs. CHF+BafA1-treated (n = 234) groups (G). Individual data points are presented with a bin width of either 0.8 µm² (E and G) or 0.25 µm² (F). The data were analyzed by Kolmogorov–Smirnov test (KS test). H: no. of autophagosomes per 58.83-µm² area. Values are means ± SE. The data were analyzed by 2-way ANOVA followed by Sidak’s multiple comparison test. Interaction: not significant; procedure (sham vs. CHF): P < 0.001; treatment with BafA1: not significant. **P < 0.01. ns, Not significant.

Salvianolic acid B inhibits MMP9 protein expression and activity during post-MI induced CHF.

Since our data showed that cardiac MMP9 level and activity are increased in CHF rats, we searched for a selective MMP9 inhibitor to intervene. Although a pure and selective MMP9 inhibitor is practically unavailable currently, we searched for a pharmacological inhibitor that has a minimal off-target inhibitory effect on the other members of the MMP family. In this regard, we selected commercially available SalB (10 mg/kg) for in vivo MMP9 inhibition. Salvianolic acid is named for its presence in salvia roots (Salvia miltiorrhiza) and is used as a traditional medicine in China, Japan, and the United States as “dan shen” or “red sage” to treat cardiovascular disease, fibrotic disease, and cancer (7, 57, 131). SalB is reported to be a selective inhibitor of MMP9 in rat hearts during after MI and hypertensive cardiac remodeling in spontaneously hypertensive (SHR) rats (32, 33). Moreover, salvianolic acid has been reported to be cardioprotective and attenuates ANG II-induced fibrotic response in cardiac fibroblasts by inhibiting the NF-κB pathway (57, 116, 125). Importantly, SalB has no deleterious effects on health in rats treated for 15 days with a wide range of doses (32, 33). First, we validated the effect of SalB on cardiac MMP9 inhibition. We treated Sham and CHF rats with SalB for 7 days and isolated the hearts to measure the MMP9 expression and activity. Our Western blot and in-gel gelatin zymography validation showed that this dose and duration of SalB given via an intraperitoneal route of treatment effectively inhibited the cardiac level of MMP9 expression and activity (Fig. 5, A–D) and cardiac peri-infarct fibrosis (Fig. 5, E and F).

MMP9 inhibition by SalB increased cardiac autophagic flux in the peri-infarct area of post-MI induced CHF.

Our results validated that SalB inhibits MMP9 during CHF (Fig. 5, A–D) (32). Subsequently, we investigated the effect of MMP9 inhibition on cardiac autophagic flux in Sham and CHF rats. We quantified the level of Lamp1, p62, and LC3B-II protein level from the peri-infarct areas of LV in Sham and CHF rats treated with SalB and measured autophagic flux with BafA1 or DMSO (Fig. 6). Our results demonstrated that cardiac LC3B-II/actin level is accumulated with BafA1 treatment in the Sham+SalB-treated group, indicating the enhanced housekeeping level of autophagic degradation in the Sham rats treated with MMP9 inhibition (Fig. 6, A–D). Interestingly, the LC3B-II/actin level of accumulation is observed to be further increased in CHF rats treated with SalB (Fig. 6D), suggesting that the autophagic degradation is enhanced or adaptively increased in the CHF group peri-infarct areas with MMP9 inhibition. Furthermore, our data demonstrated that accumulations of Lamp1, p62, LC3B-II, and LC3A-II levels are significantly higher in the peri-infarct areas of the CHF group treated with SalB+BafA1 (Fig. 6, A–H), which further supports that MMP9 inhibition upregulates p62-mediated autophagic cargo selection and their clearance via increased autophagosome/autolysosome terminal degradation.

Fig. 6. — Matrix metalloprotease (MMP)9 inhibition by salvianolic acid B (SalB) increased autophagic flux in peri-infarct areas during post-myocardial infarction (MI) induced chronic heart failure (CHF). Autophagic flux was evaluated in the peri-infarct areas with bafilomycin A1 (BafA1) treatment in 2 groups of rats: sham operated (Sham)+SalB vs. CHF+SalB. SalB (10 mg/kg) was injected intraperitoneally for 7 days in 6–8 wk post-MI rats. BafA1 (6 μmol/kg body wt ip) was injected 2 h before euthanasia. A: representative Western blot images showing expression of LC3B-II and p62 in matched areas from Sham and CHF rats. *B–D*: bar graphs showing quantification of Lamp1 (B), p62 (C), and LC3B-II (D) normalized to actin in Sham and CHF rats. Values are means ± SE; n = 6 rats. E and G: representative single-gel Western blot images showing expression of LC3B and LC3A in 6 groups of samples. F and H: bar graphs showing quantification of LC3B-II (F) and LC3A-II (H) normalized to actin in sham-operated or peri-infarct areas of the myocardium, Values are means ± SE; n = 3 rats. The data were analyzed by 2-way ANOVA followed by Tukey’s multiple comparison test. RAU, relative arbitrary unit. *P < 0.05 vs. Sham+SalB+BafA1; **P < 0.05 vs. Sham; #P < 0.05 vs. CHF.

Pharmacological MMP9 inhibition in H9c2 cardiomyocytes increased autophagic degradation.

Since our in vivo data demonstrated that MMP9 inhibition upregulates cardiac autophagic flux in post-MI CHF rats, we attempted to corroborate the findings in cardiomyocytes using H9c2 cardiomyocytes. We used differentiated H9c2 cardiomyocytes with the protocol described in materials and methods, which mimics the features of adult cardiomyocytes. To investigate the effect of MMP9 inhibition on cardiomyocytes for autophagic change in vitro, we treated them with SalB (50 µM) or MMP-9 Inhibitor-I (another selective MMP9 pharmacological inhibitor at 5 nM dose) for 24 h, with lysosomal degradation being blocked by addition of BafA1 (100 nM) for the last 2 h of treatment to measure the autophagic flux. First, we validated the pharmacological inhibition of MMP9 with SalB or MMP-9 Inhibitor-I by in-gel gelatin zymography. The results confirmed the inhibition of MMP9 in H9c2 cardiomyocytes with these treatments (Fig. 7, A and B). Next, we measured the autophagic vacuole accumulation with BafA1 treatment, using LC3B immunocytochemistry assay in cardiomyocytes with MMP9 inhibition. Principally, we measured the LC3B immunofluorescence intensity in cardiomyocytes with BafA1 treatment and compared them with their corresponding treated groups with no BafA1 treatment (Fig. 7C). An increased LC3B intensity with BafA1 represents an increased level of autophagosomes. Our data showed that SalB or MMP-9 Inhibitor-I treatment in cardiomyocytes significantly increased the LC3B intensity (Fig. 7, C and D). Thus, our in vitro data confirmed that MMP9 inhibition in H9c2 cardiomyocytes increases autophagic flux. Furthermore, we performed an autophagy sensor RFP-GFP-LC3B fluorescence imaging assay on cardiomyocytes to reconfirm the autophagy flux, where autophagosomes (APs) are visible as yellow puncta and autolysosomes (ALs) as free red puncta. Our results showed that MMP9 inhibition in cardiomyocytes increased yellow puncta numbers (APs) and increased free red puncta numbers (ALs), validating increased autophagic flux (Fig. 7, E and F). Thus, our in vitro data confirmed that MMP9 inhibition in cardiomyocytes increases autophagic flux.

Fig. 7. — Pharmacological matrix metalloprotease (MMP)9 inhibition in H9c2 cardiomyocytes increased autophagic flux. A: representative in-gel gelatin zymography showing MMP9 activity in H9c2 cardiomyocytes treated with salvianolic acid B (SalB, 50 µM) or MMP-9 Inhibitor-I (MMP-9i, 5 nM) for 24 h. B: bar graph showing the quantification of MMP9 activity. Values are means ± SE; n = 4 rats. C: evaluation of autophagosome formation and degradation by immunocytochemistry technique in H9c2 cardiomyocytes with DMSO or bafilomycin A1 (BafA1) treatment (100 nM, 2 h before fixation). Representative immunocytochemistry images showing H9c2 cardiomyocytes treated with SalB or MMP-9i in presence or absence of BafA1; LC3B (indicates autophagosomes), F actin (red), and DAPI (blue) stained the nucleus. D: bar graph showing quantification of LC3B green color intensity in each group. Values are means ± SE; n = 5 rats. The LC3B intensity with BafA1 represents autophagosome quantity accumulated because of lysosomal inhibition of autophagic degradation. E: representative red fluorescent protein (RFP)-green fluorescent protein (GFP)-LC3B fluorescence imaging of H9c2 cardiomyocytes. Yellow puncta represent autophagosomes (APs), and free red puncta represent autolysosomes (ALs). F: bar graph showing the number of APs and ALs in each group, Values are means ± SE; n = 10 rats. The data were analyzed by 1-way ANOVA (B) and 2-way ANOVA (D and F) followed by Tukey’s multiple comparison test. RAU, relative arbitrary unit. *P < 0.05 vs. Control; #P < 0.05 vs. Control+BafA1, Scale bars, 25 µm.

Genetic ablation of MMP9 in cardiomyocytes and fibroblasts upregulates autophagic flux.

As an alternate approach to pharmacological inhibition of MMP9 in the heart, we implemented genetic manipulation of MMP9 ablation with the CRISPR/cas9 technique in H9c2 cardiomyocytes and fibroblasts in vitro. We performed CRISPR/cas9 and MMP9-HDR plasmid cotransfection in vitro and selected for MMP9 knockdown cell population, with puromycin selection having an expression of RFP-fusion protein (Fig. 8A and Fig. 9C). Our Western blot and in-gel gelatin zymography results demonstrated that MMP9 expression and activity were significantly decreased in the MMP9^+/− H9c2 cardiomyocytes and MMP9^+/− fibroblasts (Fig. 8, B–E, and Fig. 9, D and E). We measured the autophagic flux in control and MMP9^+/− cardiomyocytes with LC3B immunocytochemistry. Our data demonstrated that MMP9 genetic ablation significantly accumulated LC3B puncta intensity in MMP9^+/− cardiomyocytes with lysosomal inhibitor BafA1 treatment compared with control cells with BafA1 treatment (Fig. 8, F and G), suggesting increased autophagic flux. Interestingly, similar increase in autophagy flux was observed with the Western blot (WB) technique in cardiomyocytes and fibroblasts with MMP9 genetic ablation, where we measured the Lamp1, p62, and LC3B-II protein levels by WB in control and MMP9^+/− cells with BafA1 treatment and compared their levels of accumulation. Our autophagic flux data demonstrated that MMP9 genetic ablation significantly accumulated Lamp1, p62, and LC3B-II protein levels in the MMP9^+/− fibroblasts and in cardiomyocytes with lysosomal inhibitor BafA1 treatment compared with control cells with BafA1 treatment (Fig. 9, F–I, and Fig. 10, A–D), demonstrating that MMP9 ablation in cardiomyocytes and fibroblasts increased autophagic flux. Moreover, RFP-GFP-LC3B data reconfirm that MMP9 inhibition increased autophagic flux in fibroblasts (Fig. 9, J and K).

Fig. 8. — Genetic ablation of matrix metalloprotease (MMP)9 in H9c2 cardiomyocytes increased autophagosome formation. A: representative cellular images showing rat MMP9^+/− H9c2-red fluorescent protein (RFP) cardiomyocytes. Rat MMP9 CRISPR/Cas9-knockout (KO) construct and MMP9-homology-directed repair (HDR) construct (Cat. No. sc-437355, Santa Cruz Biotechnology) were cotransfected in H9c2 cells and selected with puromycin. B: representative Western blot images showing reduced MMP9 expression in MMP9^+/− H9c2 cardiomyocytes. C: bar graph showing quantification of the MMP9 level. D: representative zymography image showing reduced MMP9 activity in MMP9^+/− H9c2 cardiomyocytes. E: bar graph showing quantification of the active MMP9 level. F: representative immunocytochemistry images showing control and MMP9^+/− H9c2 cardiomyocytes treated with DMSO or bafilomycin A1 (BafA1, 100 nM) for 2 h. LC3B (indicates autophagosomes), F actin, and DAPI stained the nucleus. G: bar graph showing quantification of LC3B green color intensity. The increased LC3B intensity with BafA1 indicates autophagosomes accumulated because of lysosomal inhibition. Values are means ± SE; n = 6 rats. The data were analyzed by Student’s t-test (C and E) and 2-way ANOVA followed by Tukey’s multiple comparison test (G). RAU, relative arbitrary unit. *P < 0.05 vs. Control; #P < 0.05 vs. Control+BafA1. Scale bars, 25 µm.

Fig. 9. — Matrix metalloprotease (MMP)9^+/− fibroblasts showed increased autophagic flux. A: representative phase-contrast image showing wild-type mouse embryonic fibroblasts (MEFs). B: differentiated MEFs are validated for fibronectin immunoreactivity (Cat. No. ab23750, Abcam), with immunocytochemistry (ICC) technique. C: representative cellular image showing mouse MMP9^+/− red fluorescent protein (RFP) embryonic fibroblasts. Mouse MMP9 CRISPR/Cas9-knockout (KO) construct and MMP9-homology-directed repair (HDR) construct (Cat. No. sc-421679, Santa Cruz Biotechnology) were cotransfected in H9c2 cells and selected with puromycin. D: representative in-gel gelatin zymography showing MMP9 activity in control and MMP9^+/− MEFs. E: bar graph showing quantification of MMP9 activity. Values are means ± SE; n = 3 rats. F: representative Western blot images showing Lamp1, p62, and LC3B-II protein level accumulation in control and MMP9^+/− RFP embryonic fibroblasts treated with DMSO or bafilomycin A1 (BafA1, 100 nM) for 2 h before harvest. *G–I*: quantification of Lamp1 (G), p62 (H), and LC3B-II (I) normalized to actin. Values are means ± SE; n = 3 rats. J and K: representative RFP-green fluorescent protein (GFP)-LC3B fluorescence imaging of control fibroblasts and fibroblasts treated with MMP9 inhibitor for 24 h. Yellow puncta represent autophagosomes (APs), and free red puncta represent autolysosomes (ALs); n = 5 rats. RAU, relative arbitrary unit. The data were analyzed by Student’s t test (D) and 2-way ANOVA followed by Tukey’s multiple comparison test (*G–I* and K). *P < 0.05 vs. Control+BafA1; #P < 0.05 vs. Control. Scale bars, 100 µm (A), 200 µm (B and C), 50 µm (J).

Fig. 10. — Genetic matrix metalloprotease (MMP)9 ablation in H9c2 cardiomyocytes increased autophagic flux and autophagy signaling. A: representative Western blot images showing expression of Lamp1, p62, and LC3B-II in MMP9^+/− H9c2 cardiomyocytes. Autophagic flux was evaluated in control and MMP9^+/− H9c2 cardiomyocytes with bafilomycin A1 (BafA1, 100 nM for 2 h) or DMSO treatment in each group. *B–D*: bar graphs showing quantification of Lamp1 (B), LC3B-II (C), and p62 (D) band intensity normalized to actin in the control and MMP9^+/− H9c2 cardiomyocyte groups. Values are means ± SE of analyses; n = 3 rats. Two-way ANOVA was used to generate P values. E and F: representative Western blot images showing expression of phospho- (p-)AMP-activated protein kinase (AMPK)α^Thr172 and p-mechanistic target of rapamycin (mToR)^ser2448 activity in control and MMP9^+/− H9c2 cardiomyocytes and bar graphs showing quantification of p-AMPKα/total (t)-AMPKα and p-mTOR/t-mTOR; actin is shown as a loading control. n = 6 rats. G: representative Western blot images showing expression of Beclin-1 and Atg7 in control and MMP9^+/− H9c2 cardiomyocytes. H and I: bar graphs showing quantification of Beclin-1 and Atg7 expression normalized to actin. Values are means ± SE; n = 6 rats. RAU, relative arbitrary unit. The data were analyzed by 2-way ANOVA followed by Tukey’s multiple comparison test (*B–D*) and by Student’s t test (*E–I*). *P < 0.05 vs. Control+BafA1; #P < 0.05 vs. Control.

Furthermore, to understand the mechanism of MMP9 inhibition and autophagosome formation signaling in cardiomyocytes, we measured protein expression level and phosphorylation activity of key autophagy regulators. We measured autophagy upstream regulators AMPKα (inducer) and mTOR (suppressor) phosphorylation activity (40, 64) and autophagy downstream regulator Beclin-1 and Atg7 expression levels, which are involved in the cargo membrane shuttling during autophagosome formation (61, 64, 81). Our data demonstrated that MMP9 ablation in cardiomyocytes increased the activity of AMPKα and decreased the activity of mTOR (Fig. 10, E and F), suggesting that MMP9 ablation affects the upstream regulators AMPKα and mTOR to increase the autophagic flux in cardiomyocytes. Furthermore, MMP9 ablation showed increased Beclin-1 and Atg7 level in the cardiomyocytes (Fig. 10, G–I), suggesting that reduced MMP9 activity increased autophagic signaling.

SalB-induced autophagy upregulation is essential for its cytoprotective effect.

To examine whether autophagy upregulation is essential for executing the cytoprotective effect of SalB, we performed cleaved caspase 3 WB cell death signaling assay with or without autophagy inhibitor in cultured H9c2 cells. In brief, SalB-treated cardiomyocytes were cotreated with or without LY294002 (an autophagy inhibitor) and hydrogen peroxide (H₂O₂) for 24 h (123). Our results demonstrated that addition of autophagy inhibitor LY294002 exacerbated H₂O₂-induced cell death signaling via increased cleaved caspase 3 protein level in cardiomyocytes. However, in the absence of autophagy inhibitor this effect was abolished, which suggests that autophagy inhibition blunted SalB-mediated cytoprotective effects (Fig. 11, A and B). Thus, it confirms our finding that autophagy upregulation is essential for executing the protective effect of SalB.

Fig. 11. — Autophagy upregulation is essential for cytoprotective effect of salvianolic acid B (SalB). A: representative Western blots (WB) showing caspase 3 and cleaved caspase 3 protein expression in H9c2 cardiomyocytes treated with or without autophagy inhibitor LY294002 (LY), H₂O₂, and SalB for 24 h. B: bar graph showing quantification of cleaved caspase 3 WB band intensity normalized to actin bands. C: SalB treatment improved lysosomal degradation of mitochondrial proteins in chronic heart failure (CHF); representative Western blots showing expression of total oxidative phosphorylation (OXPHOS) WB (Cat. No. ab110413, Abcam) for mitochondrial proteins in the peri-infarct whole tissue lysates of sham-operated (Sham) and CHF rats. C, complex. D: bar graph showing selective quantification of complex III intensity of total OXPHOS WB normalized to actin bands. Values are means ± SE of analyses; n = 3 rats. The data were analyzed by 2-way ANOVA followed by Tukey’s multiple comparison test. RAU, relative arbitrary unit. *P < 0.05 vs. H₂O₂+SalB; #P < 0.05 vs. Sham; §P < 0.05 vs. CHF; ¥P < 0.05 vs. CHF+SalB.

SalB increased lysosomal degradation of mitochondrial proteins in CHF heart.

We performed total OXPHOS WB to explore whether mitochondrial proteins are the main cargo of autophagy in CHF and if their degradation in lysosome is induced by MMP9 inhibition. We monitored the expression of mitochondrial total OXPHOS WB protein complex level in peri-infarct whole tissue protein extracts from Sham and CHF rats treated with or without SalB and bafilomycin A1. Our results indicated that SalB treatment alleviates defective cardiac OXPHOS mitochondrial protein complex accumulation in CHF. However, SalB restored the normal cardiac total OXPHOS protein levels in CHF hearts by increasing their terminal degradation in lysosomes, which is shown by a bafilomycin A1 chase experiment for mitochondrial proteins (Fig. 11, C and D). Our data suggest that damaged mitochondrial bulk autophagy or mitophagy might possibly be the main form of autophagy, which is induced by SalB treatment or MMP9 inhibition at the chronic phase of cardiac remodeling. Therefore, our overall data confirmed that the pharmacological or genetic ablation of MMP9 increased autophagic flux in the heart and cardiomyocytes and is most possibly associated with improved cardiac fibrosis and ECM remodeling in post-MI induced CHF (Fig. 13). Although SalB treatment for 7 days at 6 wk after MI reduced ECM fibrosis (Fig. 5, E and F), it did not show a significant change in the hemodynamics parameters (Table 2). In conclusion, for proof of principle we have shown that pharmacological inhibition of MMP9 yields an enhanced rate of autophagic degradation in the post-MI heart that is associated with reducing fibrosis and extracellular matrix remodeling.

Fig. 13. — Schematics for the mechanism of autophagy-mediated cardioprotection in post-myocardial infarction (MI) cardiac remodeling via matrix metalloprotease (MMP)9 inhibition. A: schematic showing that activation of MMP9 contributes to suppressed autophagic activity in chronic heart failure (CHF). Factors like hypoxia, increased hypoxia-inducible factor (HIF)-1α, reactive oxygen species (ROS), and cytokines caused MMP9 activation and reduced autophagic activity in CHF. However, inhibition of MMP9 increased autophagic activity in CHF. Proposed schematic flowchart showing that increased autophagic flux is one of the mechanisms by which MMP9 inhibition executes cardioprotection in CHF. ECM, extracellular matrix. B: schematic showing that MMP9 inhibition-increased autophagic cargo degradation in CHF contributes to reduced cardiac fibrosis and ECM remodeling via activation of AMP-activated protein kinase (AMPK)α, ATG7, and Beclin-1 pathway. mTOR, mechanistic target of rapamycin.

DISCUSSION

The present study shows that MMP9 is increased in the peri-infarct ECM remodeling in rats with CHF. Concomitantly, there is reduced autophagic flux in this ECM remodeling zone. Using short-term (2 h) lysosomal inhibitor BafA1 treatment, we showed adaptively insufficient autophagic degradation and impaired autophagic cargo clearance in the peri-infarct myocardium of rats with CHF. Interestingly, the pharmacological MMP9 inhibition increased cardiac autophagic degradation in the peri-infarct area, which indicates that MMP9 inhibition increased autophagic flux in the hearts of rats with CHF. Notably, MMP9 pharmacological inhibition reduced the peri-infarct fibrosis with a concomitant increase in cardiac autophagic flux. In vitro studies using rat H9c2 cardiomyocytes showed that MMP9 inhibition in cardiomyocytes reduced MMP9 activity but increased the cardiomyocyte-specific autophagic flux. Furthermore, in an alternate approach, MMP9 genetic deletion by CRISPR/cas9 technique in in vitro cardiomyocytes or in fibroblasts showed a similar result of increased autophagic flux. Taken together, these data suggest a significant increase in autophagic flux following MMP9 inhibition, possibly mitigating myocardial fibrosis, that leads to the preservation of cellular viability, reduction of infarct size, and attenuation of adverse left ventricular pathological ECM remodeling in CHF (Fig. 13).

We observed that there was an increase in HIF-1α and positive correlation of HIF-1α protein level with infarct size, which indicates that chronic hypoxia induces peri-infarct expansion in CHF (Fig. 1, E and F). Concomitantly, we observed a significant increase of MMP9 and MMP2 during the peri-infarct ECM remodeling in rats with CHF (Fig. 2). Our results are consistent and confirmatory of previous studies in which increase in HIF-1α, MMP9, and MMP2 expression and activity are documented in chronic heart failure (25, 89, 103, 106, 121). Our data support the notion that hypoxia-induced increase in HIF-1α and MMP9 level due to low oxygen increases cardiac oxidative stress, which is associated with the impaired autophagic clearance process in the peri-infarct myocardium during CHF (91). An acute or a chronic correlation between HIF-1α and MMP9 for autophagic impairment in a cardiac ischemia-reperfusion disease model is not explored, although autophagy can be considered a HIF-1-dependent adaptive metabolic response to hypoxia (126), where HIF-1α exacerbates MMP9-dependent mechanisms of ECM remodeling (11, 115).

The damaged myocardial tissue area in post-MI induced CHF was demarcated into three distinct zones termed infarct, peri-infarct, and remote zone, based on the extent of cardiac ECM remodeling (Fig. 1, A and B) (82). Previous literature suggests that the endogenous activation of MMP9 during the acute post-MI period is cardioprotective since it rapidly stimulates inflammation and ECM turnover. This is necessary to clear the necrotic myocytes and tissue debris for myocardial healing. Previous literature further supports the notion of a major cardioprotective role for MMP9 inhibition in mitigation of maladaptive left ventricular ECM remodeling in various rodent models of heart failure (31). In addition, MMP9 inhibition is known to reduce cardiac injury induced by damage-associated molecular patterns (DAMPs) released during ischemic insult for protection of the damaged myocardium in chronic cardiac pathologies (1, 31–33, 47, 67, 82, 94, 124). Moreover, multiple reports on targeted or global MMP9 inhibition provide the evidence for the net cardioprotective outcome including ischemic heart disease, cardiac hypertrophy, and heart failure (10, 25, 31, 66, 85). Despite this knowledge, it is unknown whether autophagy activation is in part responsible for the cardioprotective mechanism of MMP9 inhibition.

In this study, for the first time, we present evidence to demonstrate that MMP9 inhibition increased cardiac autophagic flux, thus suggesting that increased autophagy flux is one of the possible mechanisms by which MMP9 inhibition protects the heart during CHF. To determine this, we measured the autophagic flux in the heart of rats treated with salvianolic acid B (SalB), a selective MMP9 inhibitor previously reported for its MMP9 selective inhibition and cardioprotection during hypertension and post-MI induced pathological cardiac remodeling (32, 33). Our results demonstrated that MMP9 inhibition by SalB for 7 days inhibited MMP9 activity and reduced cardiac fibrosis in the LV peri-infarct area during CHF with a concomitant increase in autophagic flux, suggesting that increased autophagic flux with MMP9 inhibition contributes to reduce fibrosis in the LV remodeling during CHF (Fig. 5). However, this did not translate into a significant improvement in cardiac function parameters monitored in our study. This lack of improvement may be likely explained by two possibilities: First, 7 days (relatively short) may be an insufficient period of SalB treatment to see improvements after a chronic established infarct period of 6 wk in a global marker like cardiac function. Second, it is truly challenging to monitor such changes in cardiac function in a limited number of animals, as each animal develops a variable size of %infarct after coronary artery ligation corresponding with a variable decrease in cardiac function, leading to data with large variability in a limited number of animals. Therefore, further studies using a larger cohort of animals with a longer duration of SalB treatment is warranted to address this issue.

At this phase of the disease, we observed that there was significant and dramatic damage to the mitochondria in the cardiomyocytes from rats with CHF, which is consistent with previous reports (20, 60, 117, 119). Our TEM micrographs demonstrated an increase in damaged mitochondria and increased abundance of large autophagic vacuoles within close vicinity of damaged mitochondria in the hearts of rats with CHF (Fig. 4, A–D). An increased abundance of autophagosomes in CHF suggests a reduced autophagosome degradation in the peri-infarct areas of the heart in CHF, thus increasing their abundance under EM. Remarkably, short-term lysosomal inhibition by BafA1 reduced the TEM autophagic vacuole size and number in rats with CHF, which is surprising and perhaps is noticeable under TEM because of an acute lysosomal inhibition, which remains to be explored further (Fig. 4, E–H). However, the increased protein level of Lamp1 and p62 and decreased LC3B-II/actin within the CHF peri-infarct areas support the notion of an increased abundance of autophagosomes and their suppressed clearance or degradation in CHF peri-infarct areas (Fig. 3). This leads us to conclude that the reduced autophagosome degradation in the peri-infarct areas of the heart in rats with CHF increases their abundance, perhaps due to having a defect in formation, trafficking, and inability to move toward the lysosome (17, 30).

Recent research suggests that mitochondria are new horizons in cardioprotection, and mitochondrial abnormality is associated with heart diseases that compromise cardiovascular physiology (15, 83, 100). Considering that >30% of the intracellular volume of a cardiomyocyte is mitochondria (84), it is unknown whether ischemia-induced damaged mitochondria in cardiomyocytes are potential cargo for autophagosome formation in CHF. Damaged mitochondria produce excessive ROS, and their effective clearance can be mitochondria protective during hypoxia via autophagic/mitophagic vesicular fusion with lysosomes (20, 60, 117, 119). It can be accepted that under the conditions of chronic cardiac ischemic injury the process of macroautophagosomes or damaged mitochondrial autophagosome formation is activated in response to oxidative stress and energy crisis; however, their terminal clearance in lysosome is suppressed (26, 27, 64, 65, 122). We anticipated that the autophagic cargos are generated because of mitochondrial reactive species mechanisms during myocardial ischemia with or maybe without reperfusion injury (13, 23, 50) (Fig. 11. C and D). In addition, autophagosomes may be formed from the debris of pathological ECM remodeling induced by ischemia, inflammation, and collagen metabolism after MI (5, 19, 52). Moreover, reports have demonstrated a distinct role of autophagy in ischemia-reperfusion injury (58, 64, 87, 112). Furthermore, previous comprehensive studies produce several lines of evidence that autophagy is beneficial or may be insufficiently adaptive in other contexts (2, 68, 114). Notably, the enhanced or suppressed rate of autophagic flux can be accurately determined if using a lysosomal inhibitor chase experiment as per the autophagy monitoring guideline, and only an apparent level of autophagosome marker measurement might be insufficient to delineate the actual autophagic degradation (42, 44, 45). Considering this guideline, our data showed that autophagic flux is suppressed and adaptively insufficient during chronic ischemic peri-infarct at 6–8 wk post-MI induced CHF in rats (21, 26), whereas MMP9 inhibition at this stage enhanced cardiac autophagic flux in the peri-infarct myocardium (Fig. 6). In this context, suppressed autophagic flux is also reported in patients with Danon disease cardiomyopathy, a glycogen storage disease due to lysosomal protein Lamp2 mutation characterized by abnormal accumulation of autophagosome or blocked autophagosome-lysosome fusion and their terminal degradation (3, 63, 75). Furthermore, autophagy suppression below physiological level is reported in the chronic stage of transverse aortic constriction (TAC)-induced pressure-overload heart failure in mice (108).

To corroborate our whole animal in vivo data in hearts, we used in vitro H9c2 cardiomyocyte and fibroblast cultures (Figs. 7–9). To validate the mechanism in vitro that MMP9 inhibition increased autophagy flux, we measured autophagic flux in H9c2 cardiomyocytes by two independent pharmacological MMP9 inhibitors (SalB and MMP-9 Inhibitor-I treatment) and with the use of RFP-GFP-LC3B autophagy flux sensor (Fig. 7). Our data showed that pharmacological MMP9 inhibition increased autophagosome accumulation in the presence of BafA1. The RFP-GFP-LC3B autophagy flux sensor data further showed that MMP9 inhibition in cardiomyocytes increased autophagosomes and autolysosomes, validating increased autophagic flux, which confirms that MMP9 inhibition increased autophagic flux in cardiomyocytes (Fig. 7, E and F). Further testing of this hypothesis demonstrated that inhibition of MMP9 by genetic ablation via CRISPR-cas9 knockdown experiments reproduced similar results of enhanced autophagic activation both in MMP9^+/− cardiomyocytes and in MMP9^+/− fibroblasts (Figs. 8 and 9). Using MMP9^+/− cardiomyocytes, we further demonstrated that the mechanism of autophagic upregulation by MMP9 inhibition is due to AMPKα and Beclin-1 overstimulation, along with increased ATG7-mediated membrane shuttling (2, 61, 64, 81) and a concomitant decrease in autophagy suppressor mTOR activity, a negative regulator of autophagy (40, 64) (Fig. 10, E–I). Therefore, these data confirm that reduced MMP9 activity in cardiomyocytes accelerates a high demand of autophagic cargo material to meet the requirements of complete autophagosome and/or bulk mitophagosome formation and their complete terminal degradation (Fig. 11, C and D). This appears to be the key mechanism of autophagic upregulation by MMP9 inhibition in the heart and cardiomyocytes.

Modulation of the autophagic pathway has become a potential therapeutic target in several human diseases and pathophysiological conditions, including neurodegenerative diseases, diabetes, heart failure, and cardiovascular complications (56, 96). The autophagy process is categorized into microautophagy, macroautophagy, and chaperone-mediated autophagy (43). We studied macroautophagy, which has received the most focused attention in the pathophysiology of cardiovascular diseases and is referred to as “autophagy” (45). During the autophagic process, cellular cargos are engulfed in a double-membrane vacuole called the autophagosome; later they fuse with lysosomes and are degraded by lysosomal enzymes (45). A basal level of autophagic degradation is essential for normal cardiomyocyte function (68, 118). An increase in the basal rate of autophagic recycling and clearance is adaptive during acute hypoxia, perhaps allowing cardiac cells to preserve a sufficient supply of recycled energy to adapt to an acute cardiac stress (92). Our in vitro data demonstrated that MMP9 inhibition increased autophagic flux in cardiomyocytes and in fibroblasts (Figs. 7–10). These data suggest that MMP9 inhibition-induced autophagy is a conserved cellular mechanism. We believe that an improved autophagic flux brings cardiomyocyte and fibroblast specific cellular homeostasis via accelerating degradation and recycling process in post-MI stressed myocardium. Therefore, our data support that increased autophagy reduced cardiomyocyte damage and apoptosis-related cell death signaling (Fig. 11, A and B) in post-MI myocardium, which improved post-MI cardiac remodeling. In parallel, increased autophagy in cardiac fibroblasts brings cellular homeostasis that perhaps diminishes fibroblast activation in response to post-MI, and thereby executes cardioprotection via reduced myocardial fibrosis and associated cardiac inflammation. Our data support a reduced autophagic flux in the heart of rats with CHF, suggesting a reduced removal of autophagic cargos generated from damaged organelles [e.g., damaged mitochondria, endoplasmic reticulum (ER)] and misfolded proteins during CHF (Figs. 3, 4, 11, C and D, and 12); however, the exact mechanism by which MMP9 inhibition increased autophagic degradation that helps to reduce the cardiac ECM fibrosis is not completely understood. We demonstrated that SalB-induced autophagy upregulation is essential for its cytoprotective effect (Fig. 11, A and B). Our study showed that MMP9 inhibition caused autophagy induction, and SalB has been reported for MMP9 inhibition in the heart (32, 33). Moreover, reports demonstrated that SalB is a novel autophagy inducer (35, 54).

Fig. 12. — Autophagic flux is suppressed in remote areas of post-myocardial infarction (MI) induced chronic heart failure (CHF). Autophagic flux was evaluated in sham-operated (Sham) or remote areas with bafilomycin A1 (BafA1) treatment in each group of rats (Sham and CHF). BafA1 (6 μmol/kg body wt ip) was injected for 2 h before euthanasia. A: representative Western blot images showing expression of Lamp1, p62, and LC3B-II in the remote areas of left ventricular myocardium 6–8 wk after MI. *B–D*: bar graphs showing quantification of Lamp1 (B), p62 (C), and LC3B-II (D) normalized to actin in the area-matched Sham or remote areas of CHF myocardium. The data were analyzed by 2-way ANOVA followed by Tukey’s multiple comparison test. Values are means ± SE; n = 6 rats. There were no significant differences for Lamp1, p62, and LC3B-II in Sham+BafA1 vs. CHF+BafA1 groups. *P < 0.05 vs. Sham.

Furthermore, our data suggest that the activation of AMPKα downstream to MMP9 inhibition might be accomplishing anti-inflammatory and antifibrosis activity during myocardial ischemia (Fig. 13), as reported previously (8, 12, 59, 88, 127). In conclusion, the implications of the present study demonstrate that one of the cardioprotective mechanisms of MMP9 inhibition is mediated via autophagic upregulation. This novel and insightful finding of autophagy activation can be considered and utilized to develop a potential therapeutic and translational approach to manage and mitigate ECM remodeling in chronic heart failure.

Limitations of the study.

Some limitations of this study that we acknowledge are the following: 1) A purely selective MMP9 inhibitor has been difficult to synthesize pharmacologically; this is due to a variety of reasons that have been discussed and widely reviewed previously (36). Some of these include similar catalytic site of multiple MMP members and their overlapping expression and activation pattern. A cardiac-specific MMP9 inhibitor also has been difficult to synthesize pharmacologically because of cardiac nonresident MMP9 or circulating MMP9 sources (via macrophage, neutrophil, lymphocyte infiltration) in the heart. 2) Further identification of MMP9 substrate proteins/molecules that trigger autophagy signaling is necessary to understand how MMP9 regulates autophagic signaling at cargo selection. 3) Identification of MMP9 preferential cleavable target molecules in the heart during the CHF setting that execute cardioprotection is further necessary to consider. 4) Since the cardiac MMP9 level is constituted of multiple sources in the post-MI heart, e.g., cellular, extracellular, and systemic cell compartment via circulatory infiltration, the nontargeted MMP9 inhibition may have an off-target negative outcome in CHF. Thus, the approaches of the therapeutic intervention for targeting the cardiac level of MMP9 must be carefully evaluated. 5) Autophagy induction by SalB has been reported so far (35, 54). SalB showed multiple mechanisms of cardiovascular protection including antioxidant, anticoagulant, antiapoptotic, and antitumor activities (14, 29). Considering SalB as a naturally occurring compound (exists in herbs, e.g., S. miltiorrhiza), we are assuming that its off-target effect may be minimal. However, we cannot entirely rule out these off-target effects in this study. 6) We used bafilomycin A1 to inhibit lysosome-autophagosome fusion based on autophagy monitoring guidelines for measuring flux (45). However, a purely selective lysosomal inhibitor has been difficult to synthesize pharmacologically, and lysosomal inhibitors are also reported to have off-target effects (90, 104). Consistent with our observations, Pan et al. (77) recently reported that bafilomycin A1 has an inhibitory effect on WB LC3-II level. In addition, Gottlieb et al. (22) reported and discussed that the reduction in WB autophagy marker LC3-II after lysosomal inhibition is associated with chronic flux blockade. 7) Our study provides a possible link that SalB increased autophagic flux via its effect on MMP9 inhibition (Fig. 13). We acknowledge that there are multiple limitations to experimentally monitoring the direct effects of autophagy in vivo. 8) The relationship between autophagy and cardiovascular diseases is highly complex, as we also know that increased and/or decreased autophagic flux are reported as context and stimulus dependent in the setting of cardiac acute or chronic pathological conditions (22, 51, 64, 108).

Future directions.

The present study demonstrated that MMP9 regulates cardiac autophagy flux during CHF. Our data suggest that autophagic flux increased by MMP9 inhibition in part protects the MI heart from adverse left ventricular remodeling. MMP9 selective inhibition at a lower dose is effective in the cardiac setting (18, 32, 33, 98, 99, 105). Therefore, studies evaluating the effects of a highly selective and cardiac-specific targeted MMP9 inhibition approach for CHF remodeling are potentially promising for therapeutic avenues for future research. Our study supports the idea that reactivation or accelerating autophagic degradation, in particular through the inhibition of MMP9 or other autophagy-boosting pharmacological agents, may represent a promising therapeutic option to treat subjects with CHF (72). Similar studies with a monoclonal antibody targeting selectively cardiac MMP9 (andecaliximab, GS-5745) and highly selective MMP9 inhibitor molecules are warranted in the future. MMP9 knockdown stimulated intracellular signaling of AMPK and inhibited mTOR signaling; the underlying correlation or causative mechanism needs future study. Furthermore, rescue experiments with AMPK inhibitors and/or autophagy-targeting drugs will dissect the mechanistic association of MMP9 and autophagy. In addition, MMP2 is activated in the peri-infarct myocardium after MI; therefore the contribution of MMP2 in autophagy warrants another future study. Furthermore, the level of autophagy and its function can be phase dependent in post-MI peri-infarct or remote areas. In addition, future studies will elucidate the level of autophagy/mitophagy and the effect of MMP9 inhibition at multiple time points of post-MI remodeling.

GRANTS

This work was supported by American Heart Association Career Development Award Grant 19CDA34490029 (to S. S. Nandi), National Institutes of Health Grants R01 DK-114663 and P01 HL-62222, and endowed McIntyre Professorship (to K. P. Patel).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

S.S.N. conceived and designed research; S.S.N., K.K., and S.K.M. performed experiments; S.S.N., K.K., and S.K.M. analyzed data; S.S.N. and S.K.M. interpreted results of experiments; S.S.N., K.K., and S.K.M. prepared figures; S.S.N. drafted manuscript; S.S.N., N.M.S., D.R.A., and K.P.P. edited and revised manuscript; S.S.N., K.K., N.M.S., D.R.A., S.K.M., and K.P.P. approved final version of manuscript.

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PERMALINK

MMP9 inhibition increases autophagic flux in chronic heart failure

Shyam S Nandi

Kenichi Katsurada

Neeru M Sharma

Daniel R Anderson

Sushil K Mahata

Kaushik P Patel

Series information

Abstract

INTRODUCTION

MATERIALS AND METHODS

Animals.

Hemodynamic measurements.

Chronic heart failure model.

Fig. 1.

Table 1.

Table 2.

MMP9 inhibition in vivo.

Transmission electron microscopy.

H9c2 cardiomyocyte culture.

H9c2 cardiomyocyte treatment.

Mouse embryonic fibroblast culture.

Western immunoblotting.

In-gel gelatin zymography.

Immunocytochemistry.

Measurement of in vivo autophagic flux.

Measurement of peri-infarct fibrosis.

MMP9 genetic ablation in H9c2 cardiomyocytes by CRISPR/cas9 technique.

MMP9 genetic ablation in fibroblasts by CRISPR/cas9 technique.

Measurement of autophagic flux by tandem sensor RFP-GFP-LC3B assay.

Statistical analysis.

RESULTS

Characterization of myocardial infarction induced chronic heart failure in rats.

Increased MMP9 activity contributes to pathological cardiac remodeling in post-MI induced CHF.

Fig. 2.

Suppressed cardiac autophagic flux in the peri-infarct area of post-MI induced CHF.

Fig. 3.

Transmission electron microscopy micrographs showed increased autophagosomes and mitochondrial injury in the peri-infarct area of post-MI induced CHF.

Fig. 4.

Salvianolic acid B inhibits MMP9 protein expression and activity during post-MI induced CHF.

Fig. 5.

MMP9 inhibition by SalB increased cardiac autophagic flux in the peri-infarct area of post-MI induced CHF.

Fig. 6.

Pharmacological MMP9 inhibition in H9c2 cardiomyocytes increased autophagic degradation.

Fig. 7.

Genetic ablation of MMP9 in cardiomyocytes and fibroblasts upregulates autophagic flux.

Fig. 8.

Fig. 9.

Fig. 10.

SalB-induced autophagy upregulation is essential for its cytoprotective effect.

Fig. 11.

SalB increased lysosomal degradation of mitochondrial proteins in CHF heart.

Fig. 13.

DISCUSSION

Fig. 12.

Limitations of the study.

Future directions.

GRANTS

DISCLOSURES

AUTHOR CONTRIBUTIONS

REFERENCES

ACTIONS

PERMALINK

RESOURCES

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