Abstract
Myocardial pathologies resulting from SARS-CoV-2 infections are consistently rising with mounting case rates and reinfections; however, the precise global burden is largely unknown and will have an unprecedented impact. Understanding the mechanisms of COVID-19-mediated cardiac injury is essential toward the development of cardioprotective agents that are urgently needed. Assessing novel therapeutic strategies to tackle COVID-19 necessitates an animal model that recapitulates human disease. Here, we sought to compare SARS-CoV-2-infected animals with patients with COVID-19 to identify common mechanisms of cardiac injury. Two-month-old hamsters were infected with either the ancestral (D614) or Delta variant (B.1.617.2) of SARS-CoV-2 for 2 days, 7 days, and/or 14 days. We measured viral RNA and cytokine expression at the earlier time points to capture the initial stages of infection in the lung and heart. We assessed myocardial angiotensin-converting enzyme 2 (ACE2), the entry receptor for the SARS-CoV-2 virus, and cardioprotective enzyme, as well as markers for inflammatory cell infiltration in the hamster hearts at days 7 and 14. In parallel, human hearts were stained for ACE2, viral nucleocapsid, and inflammatory cells. Indeed, we identify myocardial ACE2 downregulation and myeloid cell burden as common events in both hamsters and humans infected with SARS-CoV-2, and we propose targeting downstream ACE2 downregulation as a therapeutic avenue that warrants clinical investigation.
NEW & NOTEWORTHY Cardiac manifestations of COVID-19 in humans are mirrored in the SARS-CoV-2 hamster model, recapitulating myocardial damage, ACE2 downregulation, and a consistent pattern of immune cell infiltration independent of viral dose and variant. Therefore, the hamster model is a valid approach to study therapeutic strategies for COVID-19-related heart disease.
Keywords: ACE2, COVID-19, heart, inflammation
INTRODUCTION
Cardiovascular complications in severe COVID-19, namely, microvascular dysfunction, myocarditis, conduction abnormalities, and heart failure, represent an emerging global health crisis. Furthermore, cardiovascular injury resulting from SARS-CoV-2, the causative virus of COVID-19, is complicated by the postacute sequelae as also linked to an increased incidence of inflammatory heart disease and thrombotic disorders; thus, cardiac injury in COVID-19 is likely currently underestimated (1). Reports of myocardial damage and the fact that angiotensin-converting enzyme 2 (ACE2), the indispensable viral entry receptor, is expressed in the heart and other affected organs suggest a probable mechanism of direct infection (1, 2). Therefore, in light of this public health emergency, it is of paramount importance to delineate the pathological mechanisms of myocardial inflammation and injury in COVID-19 and to discover novel cardioprotective interventions.
Studying SARS-CoV-2-mediated myocardial injury requires animal models to test therapeutic interventions, a feat challenged by the limited ability of small animal models to adequately recapitulate this human disease. Generally, mouse models are studied extensively and are readily available; however, they have limited utility for COVID-19 studies since murine ACE2 does not effectively bind to the SARS-CoV-2 spike protein (3). Strategies to overcome this limitation have been developed, including modifying the viral spike protein to bind mouse ACE2 or developing humanized mice that express the human ACE2 protein (3). Despite these approaches, these mice can develop additional symptoms following SARS-CoV-2 infection, such as lethal encephalitis in humanized mice that are not captured in humans; yet, they do not develop cardiac symptoms. In addition, extensive manipulation of animals and the viral spike protein limits translational significance (3). Alternatively, the Syrian hamster model superficially resembles human COVID-19 disease, with low mortality, is readily available, and succumbs to infection by unmodified SARS-CoV-2; therefore, is a valuable tool for COVID-19 research (4). To date, however, it is unclear if hamsters exhibit similar cardiac manifestations as in human patients with COVID-19.
Here, we provide evidence that the Syrian hamster is a suitable animal model to study the cardiovascular manifestations of COVID-19, thus providing a means of studying therapeutic interventions to prevent or treat myocardial injury in patients with COVID-19.
MATERIALS AND METHODS
Cells and Viruses
Vero cells (ATCC CCL-81) were maintained in minimum essential medium (MEM) supplemented with 100 U/mL penicillin, 100 U/mL streptomycin, 0.25 µg/mL amphotericin B, and 10% fetal bovine serum. An ancestral (D614) SARS-CoV-2 strain (GISAID No. EPI_ISL_425177) and a SARS-CoV-2-Delta variant of concern strain (B.1.617.2) were used in these studies.
In Vivo Hamster Infections
Animal experiments were approved by the University of Alberta Animal Care and Use Committee (AUP00001847 and AUP00003869). All SARS-CoV-2 infection studies were conducted in a certified BSL3 containment facility at the University of Alberta. Briefly, 2-mo-old male Syrian hamsters were inoculated intranasally with the ancestral SARS-CoV-2 variant at a dose of 2.0 × 103 PFU in a total volume of 100 µL (50 µL per nare). Nasal swabs were performed on days 1, 3, and 6 after challenge and collected for histopathology on day 7 or day 14 postinfection. To determine cytokine responses following virus infection, another set of hamsters were inoculated at a viral dose of 1.0 × 106 PFU with SARS-CoV-2-Delta (B.1.617.2) and heart and lung tissues were collected on day 2 and day 7. Control hamsters were inoculated with MEM containing no virus. Animals were randomized to either the infected or control groups. The hamsters were monitored daily for signs of infection and morbidity (Fig. 1A). Animals who lost greater than 20% of their initial body weight were to be humanely euthanized and excluded from the study; however, no animals exceeded this threshold.
Figure 1.
SARS-CoV-2 infection induces lung injury and upregulates cytokine expression in hamsters. A: experimental timeline for Delta (collected at days 2 and 7 for RT-PCR) and ancestral (collected at days 7 and 14 for histopathology) SARS-CoV-2-infected hamsters. Vehicle-inoculated control hamsters were analyzed in parallel. B: anthropometric data of body weight and lung weight of ancestral SARS-CoV-2-infected hamsters. Body weight was measured and recorded daily and reflected as percent of weight change relative to day 0. Time points were compared using two-way repeated-measures ANOVA and Sidak’s multiple comparisons test. Lungs were harvested and weighed at the end of the 14-day study period. C: SARS-CoV-2 nasal swabs and plaque measurement D: gross histopathological assessment of hamster lungs using hematoxylin and eosin staining (H&E) and Masson’s trichrome staining at day 7 after ancestral SARS-CoV-2 infection E: staining for SARS-CoV-2 nucleocapsid protein in the lungs at day 7 after viral challenge F: SARS-CoV-2 viral RNA copies in the lung following SARS-CoV-2 Delta challenge G: expression of Il-1β, Tnf-α, Il-6, and Il-10 in hamster lungs at day 2 and day 7 following Delta SARS-CoV-2 infection. Cytokines are visualized as a relative expression compared with controls. Data are represented as means ± SE, and each point represents biological replicates (n = 4 hamsters/group). Unpaired Student’s t test was performed for comparisons of controls to SARS-CoV-2-treated animals. One-way ANOVA with Dunnett’s multiple comparisons test or Kruskal Wallis test with Dunn’s multiple comparisons were used to compare parametric or nonparametric data, respectively; *P < 0.05, **P < 0.01.
Virus Titrations
For virus culture, 2 × 105 cells were seeded into each well of a 12-well tissue culture plate 1 day before titration. Tenfold serial dilutions of the virus stock or nasal swab were plated in duplicate on Vero CCL-81 cells and cultured for 3 days at 37°C in MEM containing 0.5% carboxymethylcellulose (Sigma). Cells were fixed and stained with a solution containing 0.13% (wt/vol) crystal violet, 11% formaldehyde (vol/vol), and 5% ethanol (vol/vol) to visualize plaques.
Human Samples
Patients who succumbed to COVID-19 from the Lazio region were autopsied in Rome, Italy. In parallel, hearts were procured from age- and sex-matched donors following cardioplegic arrest according to the Human Organ Procurement and Exchange (HOPE) protocol. Control hearts were obtained from brain-dead donors (DBD) with no known history of cardiovascular disease. Transmural myocardial sections were formalin fixed and paraffin embedded for histological analysis. All protocols are approved by the Health Research Ethics Board of the University of Alberta.
Histological Staining
Formalin-fixed paraffin-embedded tissues were sectioned onto slides at 5-µm thickness. Hematoxylin and eosin (H&E) staining was performed according to a standard protocol. Briefly, slides were dewaxed and rehydrated by decreasing alcohol gradient. Nuclei were stained with Harris hematoxylin for 15 s, rinsed, and differentiated with 1% acid alcohol. Sections were rinsed in Scott’s tap water substitute (20 mM sodium bicarbonate, 166 mM magnesium sulfate) and then stained with eosin. Sections were dehydrated, cleared, and mounted. Masson’s trichrome staining was performed using a commercially available kit (Abcam ab150686). Briefly, deparaffinized and rehydrated slides were incubated in preheated Bouin’s fluid, cooled, and rinsed. The sections were next stained in Weigert’s iron hematoxylin working solution (equal parts of solutions A and B) and rinsed under tap water. The slides were next stained in Biebrich Scarlet/Acid Fuchsin solution and washed with distilled water. Slides were differentiated in phosphomolybdic/phosphotungstic acid solution until collagen was no longer red and transferred directly to aniline blue solution. Slides were rinsed and then differentiated in 1% acetic acid. Slides were quickly dehydrated, cleared in xylene, and mounted with a resinous mounting medium. Images were captured with a Leica DM4000 B LED microscope system.
Immunohistochemical Staining
Immunohistochemical (IHC) staining was performed on formalin-fixed paraffin-embedded tissues sectioned at 5-µm thickness and then dewaxed and rehydrated by ethanol gradient. Heat-induced epitope retrieval was achieved with preheated sodium citrate buffer, consisting of 10 mM sodium citrate and 0.05% Tween 20 (pH 6.0). Slides were blocked with 10% goat serum in 1% bovine serum albumin (BSA) and incubated with primary antibodies diluted in 1% BSA for ACE2 (1:50, R&D Systems AFF933), SARS-CoV-2 nucleocapsid (1:1,000, Bioss bs-41408R), CD15 (1:50, Abcam ab135377), CD68 [1:100, Thermo Fisher MA5-13324 (human); 1:100, AbD Serotec MCA1597 (hamster)], CD4 [1:100, Abcam ab133616 (human); 1:50, Millipore Sigma MABF415 (hamster)], and CD8 (1:50, Biolegend 200702). Antibody specificity was validated by staining control hamsters and human hearts, which do not have immune cell infiltration or virus. Antibody specificity for ACE2 was validated previously (5). Endogenous peroxidases were blocked with 10% H2O2, and slides was then incubated with horseradish peroxidase (HRP)-conjugated secondary antibodies (1:1,000, Cell Signaling Technologies) and visualized with freshly prepared 3,3′-diaminobenzidine substrate (Abcam). Slides were counterstained with regressive Harris hematoxylin, differentiated with 1% acid alcohol, then dehydrated by alcohol gradient, cleared, and mounted with organic mounting media. Staining was quantified as a percent area that the staining occupies over the total area of the image [staining area (%)] and as a count of positive staining regions per square millimeter (count/mm2). Imaging and quantification were performed blinded to the experimental groups.
RT-PCR
RNA was extracted from the lung and left ventricle of control and SARS-CoV-2-Delta-infected hamsters using TRIzol-chloroform. cDNA was reverse transcribed from 1 µg of RNA template using SuperScript II Reverse Transcriptase (Invitrogen). Real-time quantitative polymerase chain reaction (RT-PCR) with TaqMan premixed assays (ThermoFisher Scientific) and 25 ng of cDNA was used to quantify Ace2 (Cg04585346_m1), Tnf-α (Cg04607188_g1), Il-1β (Cg04576706_g1), Il-6 (Cg04486380_m1), Il-10 (Cg04628513_m1) with Hprt (Cg04448432_m1) as the housekeeping gene. Viral RNA copy number was quantified using the 2019-nCoV RUO kit (Integrated DNA Technologies) and a serial dilution of the 2019-nCoV_N positive control (Integrated DNA Technologies). Analyses of 2.5 ng of cDNA for the lung and 25 ng of cDNA for the heart were done.
Protein Extraction
Immunoblot was performed from protein samples extracted in TRIzol following removal of the aqueous phase for RNA extraction. DNA was precipitated from the organic phase with one third volume of ethanol and then centrifuged at 2,000g at 4°C for 5 min. Proteins were precipitated from the supernatant with ispropanol (1:1) and then centrifuged at 12,000g at 4°C. The pellet was washed with 0.3 M guanidine hydrochloride in 95% ethanol and centrifuged at 12,000 g, then repeated, and then washed in ethanol by the same procedure. The supernatant was decanted, and the protein pellet was dried under vacuum and then resolubilized in 500 µL of CelLytic buffer (Sigma-Aldrich) supplemented with protease inhibitors (Roche). Samples were sonicated on ice using a tipultrasonicator (Sonic Dismembrator Model 100, Fisher Scientific) set to level 2 in 4 × 20-s bursts with 30-s intervals.
Immunoblot
Extracted proteins were quantified with DC Protein Assay (BioRad), and 90 µg of protein was resolved by SDS-PAGE and then transferred to polyvinylidene fluoride membranes in transfer buffer, consisting of 25 mM Tris, 192 mM glycine, and 20% methanol (pH 8.3). Membranes were blocked in 5% nonfat milk and incubated with ACE2 (1:1,000, Abcam ab108252) primary antibody overnight and subsequently detected with HRP-conjugated secondary antibodies (1:4,000, Cell Signaling Technology) and Clarity ECL substrate (Bio-Rad). Lanes were normalized to MemCode total protein stain (Thermo Fisher Scientific). Band densitometry was quantified with Image Studio Software (LI-COR Biosciences).
Statistical Analysis
Statistics were performed with SPSS software, and graphs were created with GraphPad Prism. Data are represented as means ± SE. Unpaired Student’s t test was performed for comparisons of controls to patients with COVID-19. In comparisons exceeding two groups, cases that followed normal distribution were subject to one-way ANOVA with Dunnett’s multiple comparisons tests, and nonparametric data sets were subject to Kruskal—Wallis test with Dunn’s multiple comparisons.
RESULTS
Anthropometric assessment of SARS-CoV-2 infection in hamsters demonstrated weight loss compared with baseline over the course of the study and increased lung weight at the end of the 14-day period following ancestral SARS-CoV-2 infection (Fig. 1B). Nasal swabs of control and ancestral SARS-CoV-2-infected animals were measured on days 1, 3, and 6 to assay viral burden (Fig. 1C). Histologically, infected hamster lungs revealed multifocal parenchymal damage and thickening of the alveolar septa, indicating diffuse alveolar damage (DAD) (Fig. 1D). Trichrome staining corroborated alveolar thickening; however, staining did not demonstrate substantive parenchymal fibrosis (Fig. 1D). SARS-CoV-2 viral nucleocapsid staining was absent in control hamster lungs; however, SARS-CoV-2-treated hamsters displayed positive perivascular staining for viral proteins (Fig. 1E). To examine earlier time points, we next analyzed control and Delta SARS-CoV-2-infected hamsters at days 2 and 7, which revealed positive amplification for SARS-CoV-2 viral RNA (Fig. 1F) and increased expression of Il-1β, Tnf-α, and Il-10 in lung homogenates (Fig. 1G).
We next examined myocardial changes in the hamster heart consequent to SARS-CoV-2 infection. Infected hamsters have infrequent regions of increased mononuclear infiltrates and focal fibrosis that was absent in control animals (Fig. 2A). Furthermore, the viral nucleocapsid was detected in the heart in ancestral SARS-CoV-2-infected animals (Fig. 2B). This finding aligns with the positive amplification of SARS-CoV-2 viral RNA copies in the heart of SARS-CoV-2 Delta-infected animals, supporting myocardial infection as strain and dose independent (Fig. 2C). Following a basic assessment of myocardial damage and discovering SARS-CoV-2 viral proteins and RNA in the hamster heart, we aimed to investigate the consequence of infection on ACE2 levels. We first assessed the ancestral SARS-CoV-2 by histology, which demonstrated a reduction in ACE2 that persisted at both time intervals (days 7 and 14) (Fig. 2D). Interestingly, when we examined early time points, Ace2 expression was unchanged between control and Delta SARS-CoV-2-infected animals (Fig. 2E); however, ACE2 protein levels were significantly reduced compared with controls as in the ancestral SARS-CoV-2-infected animals (Fig. 2F). Because of the prominent myocardial inflammation present in certain COVID-19 cases (6), we next aimed to identify and quantify the immune cells of the heart following infection. Staining for mature neutrophils (CD15), macrophages (CD68), and T-cell antigens (CD4 and CD8) demonstrated a macrophage- and neutrophil-dominant pattern, with a mild increase in lymphocyte staining compared with that in control animals (Fig. 2, G and H). Congruently, expression of proinflammatory cytokines was mildly elevated in the heart, namely, Il-1β, and a trend toward an increase of Tnf-α (Fig. 2I).
Figure 2.
SARS-CoV-2 infection downregulates myocardial angiotensin-converting enzyme 2 (ACE2) and induces immune cell infiltration in hamsters. A: histological assessment of hamster hearts using hematoxylin and eosin staining (H&E) and Masson’s trichrome staining 7 days following ancestral SARS-CoV-2 challenge B: SARS-CoV-2 nucleocapsid staining in the hamster heart at 14 days after ancestral SARS-CoV-2 infection C: SARS-CoV-2 viral RNA copies in the heart following SARS-CoV-2 Delta challenge. D: representative images and pooled analysis of ACE2 staining in the hamster heart at day 7 (empty squares) and day 14 (filled squares) following inoculation with ancestral SARS-CoV-2. ACE2 staining area (%) and the number of areas with positive staining (count/mm2) are quantified. E: Ace2 mRNA expression by RT-PCR in the hamster heart at day 2 and day 7 after Delta SARS-CoV-2 infection. F: Western blot analysis and quantification of immunoreactivity (band densitometry for protein levels) of ACE2 following Delta SARS-CoV-2 inoculation. Immunoblots were visualized at a standard exposure (STD) or overexposed (OE) to visualize low protein levels of ACE2. Band densitometry was quantified and normalized to MemCode total protein stain (MEM). G: representative immune cell staining for neutrophils (CD15), macrophages (CD68), and T cells (CD4 and CD8) in the hearts of vehicle (control) and ancestral SARS-CoV-2-inoculated hamsters at day 14, and immune cell quantification (H). Day 7 (empty squares) and day 14 (filled squares) are pooled for analysis. I: expression of Tnf-α and Il-1β in hamster heart at day 2 and day 7 following Delta SARS-CoV-2 infection. Il-6 and Il-10 were below the limit of detection in the heart. Cytokines are visualized as a relative expression compared with controls. Data are represented as means ± SE, and each point represents biological replicates (n = 4–6 hamsters/group). Unpaired Student’s t test was performed for comparisons of controls to SARS-CoV-2-treated animals. One-way ANOVA with Dunnett’s multiple comparisons test or Kruskal Wallis test with Dunn’s multiple comparisons was used to compare parametric or nonparametric data, respectively; *P < 0.05, **P < 0.01.
We next compared the hamster phenotype with myocardial samples of patients who died from COVID-19 (Table 1). Basic histological staining revealed extensive mononuclear infiltrates and interstitial and vascular fibrosis (Fig. 3A). Viral nucleocapsid staining was predominantly perivascular in the myocardium (Fig. 3B). ACE2 was significantly downregulated in hearts of patient with COVID-19 compared with sex- and age-matched controls (Fig. 3C). Finally, the myocardium from patients with COVID-19 demonstrated a significant inflammatory cell burden biased toward neutrophils and macrophages consistent with a pathological diagnosis of myocarditis in 60% of the autopsied hearts (Fig. 3, D and E; Table 1).
Table 1.
Patient clinical characteristics
| Control | COVID-19 | |
|---|---|---|
| Demographics | ||
| n | 6 | 10 |
| Age, yr | 54.3 [47–67] | 69.2 [44–86] |
| Sex, male | 3 (50.0) | 7 (70.0) |
| Comorbidities, n (%) | ||
| Preexisting cardiac conditions | ||
| Dilated cardiomyopathy | 0 (0) | 1 (10.0) |
| Ischemic cardiomyopathy | 0 (0) | 2 (20.0) |
| Hypertension | 1 (16.7) | 2 (20.0) |
| Diabetes mellitus | 0 (0) | 1 (10.0) |
| Obesity | 1 (16.7) | 0 (0) |
| CKD | 0 (0) | 2 (20.0) |
| COPD | 0 (0) | 2 (20.0) |
| Acute injury, n (%) | ||
| Myocarditis/pericarditis | 0 (0) | 6 (60.0) |
| Arrhythmia | 0 (0) | 4 (40.0) |
| Coagulopathy | 0 (0) | 6 (60.0) |
| AKI | 0 (0) | 8 (80.0) |
| ARDS | 0 (0) | 10 (100) |
Continuous variables are reported by mean with the range in brackets: age. Categorical variables are reported by count with percentage in parenthesis: sex, comorbidities, and diagnoses.
Obesity is defined as a BMI ≥ 30 kg/m2. AKI, acute kidney injury; ARDS, acute respiratory distress syndrome; BMI, body mass index; CKD, chronic kidney disease; COPD, chronic obstructive pulmonary disease.
Figure 3.
Severe COVID-19 leads to myocardial angiotensin-converting enzyme 2 (ACE2) downregulation and immune cell infiltration in humans. A: routine histological assessment of human control and COVID-19 hearts with hematoxylin and eosin staining (H&E) and Masson’s trichrome staining. B and C: SARS-CoV-2 nucleocapsid and ACE2 staining in the human heart. ACE2 staining area (%) and the number of areas with positive staining (count/mm2) are quantified. D: immune cell staining for neutrophils (CD15), macrophages (CD68), and T cells (CD4 and CD8) in the hearts of control donors and patients with COVID-19, and immune cell quantification (E). Data are represented as means ± SE, and each point represents individual control donors (n = 3–4) or COVID-19 patients (n = 8–10). Unpaired Student’s t test was performed for comparisons of controls to patients with COVID-19; *P < 0.05, **P < 0.01.
DISCUSSION
Our data corroborate pulmonary histological findings in hamsters that mirror findings from human patients with COVID-19 (7), with a common mechanism of myocardial ACE2 downregulation that aligns with previous work from the SARS-CoV epidemic (8). Although humans had a greater abundance of immune cell infiltrates, this difference likely reflects the enhanced severity of humans who died of COVID-19 compared with animals that survived and recovered. Nevertheless, the pattern of immune cell burden coincided in humans and hamsters, suggesting SARS-CoV-2 infection facilitates a consistent macrophage- and neutrophil-dominant recruitment. These findings are unexpected, as cell-mediated immune responses driven by T lymphocytes are implicated in typical cases of viral myocarditis, such as coxsackievirus B-mediated myocardial inflammation (6). It remains unclear if SARS-CoV-2 predominately mediates direct cardiac injury because of ACE2 tropism or if myocarditis is consequent of indirect, cytokine-activated cardiotoxicity. However, we provide evidence of SARS-CoV-2 viral nucleocapsid and positive viral RNA amplification in the lungs and heart independent of viral strain and dose that appears preferentially localized to vessels. This corroborates the detectable myocardial SARS-CoV-2 viral load in the hearts of patients with COVID-19 (6) and work supporting a direct viral infection of the myocardium in a subset of postmortem autopsy hearts in patients with SARS, which demonstrated increased fibrosis, inflammation, and a reduction of myocardial ACE2 (8). Furthermore, the localization of viral nucleocapsid and ACE2 is supported by single-nucleus RNA sequencing studies that identify pericytes, vascular smooth muscle cells, and fibroblasts as harboring the greatest ACE2 expression in the human left ventricle (9).
Predominant myeloid recruitment aligns with tangential evidence emerging from the COVID-19 pandemic. Interstitial macrophages loaded with viral particles reside in the myocardium of a patient with COVID-19-mediated cardiogenic shock (10), and macrophages are the primary cell for SARS-CoV viral replication (11). Consistently, macrophage-derived cytokine interleukin 6 (IL-6) is part of a significant predictive signature of COVID-19 mortality (12). Although Il-6 was below the limit of detection in hamster lung and myocardial tissue, we observed the paradoxical upregulation of anti-inflammatory cytokine Il-10 that was found to predict increased disease severity (13).
As concomitant cardiovascular disease (CVD) in patients with COVID-19 is a risk factor for severe disease, SARS-CoV-2 itself mediates myocardial damage, and SARS-CoV-2 exploits ACE2 for cellular entry, it is essential to delineate the role of ACE2 in CVD and COVID-19 (14–16). ACE2 is protective in CVD to counteract the proinflammatory, hypertensive arm of the canonical renin-angiotensin system (RAS) mediated by angiotensin II (ANG II). Specifically, ACE2 deactivates ANG II to Ang-(1–7), a peptide that acts on the Mas receptor to promote vasodilation and anti-inflammatory effects (1, 14). A disintegrin and metalloprotease 17 (ADAM17) facilitates proteolytic ectodomain shedding of membrane-bound proteins, including ACE2; thus, deactivating and releasing it into the systemic circulation (17, 18). Indeed, ADAM17 is aberrantly activated in CVD and may lead to a deficiency in membrane ACE2, which is associated with worsened outcomes in hypertension, heart failure, and coronary artery disease (1).
SARS-CoV-2 also activates ADAM17 to foster ACE2 loss (19), which is suggested to be critically involved in COVID-19 pathogenesis as coronaviruses that cause the common cold do not activate ADAM17 (19). We previously proposed that increased ACE2 in healthy, aged males may confer susceptibility to SARS-CoV-2 infection (20); however, the downstream consequence of SARS-CoV-2 is a reduction of membrane ACE2 (measured as an increase in plasma ACE2) as a result of ADAM17 activity (1, 19). Consistently, progressive elevation in soluble plasma ACE2 in an intraindividual serial sampling of hospitalized patients with COVID-19 predicted cumulative mortality, similarly with increased surrogate markers of ADAM17 activity (1). This highlights the double-edged sword of ACE2 in COVID-19, as higher ACE2 may increase SARS-CoV-2 viral load in the initial stages of infection, where viral load is positively associated with disease severity (20); however, postacute ADAM17 activity and viral endocytosis promote ACE2 proteolytic shedding (1). This suggests ADAM17 inhibition is likely a promising therapeutic target in SARS-CoV-2 infection to circumvent loss of protective membrane-bound ACE2.
Taken together, our comparative study demonstrates that hamsters exhibit similar downstream effects of SARS-CoV-2 infection as human patients with COVID-19, recapitulating myocardial damage, ACE2 downregulation, and a consistent pattern of immune cell infiltration. Therefore, ACE2 is a double-edged sword in COVID-19, such that increased ACE2 may enhance infection susceptibility in the initial stages, yet maintaining tissue levels of cardioprotective ACE2 will likely ameliorate myocardial injury. Critical to resolving the COVID-19 pandemic aftermath is screening novel therapeutic strategies; thus, the hamster model provides a means to target both the immune cell burden and loss of membrane ACE2—two mechanisms that drive disease pathogenesis.
GRANTS
This study was supported by Canadian Institutes of Health Research Grant PJT-451105.
DISCLOSURES
Z. Kassiri is an editor of American Journal of Physiology-Heart and Circulatory Physiology and was not involved and did not have access to information regarding the peer-review process or final disposition of this article. An alternate editor oversaw the peer-review and decision-making process for this article. None of the other authors had any conflicts of interest, financial or otherwise, to disclose.
AUTHOR CONTRIBUTIONS
X.C-C., D.J.M., Z.K., F.D.N., D.H.E., and G.Y.O. conceived and designed research; A.V., R.S.N., M.G., D.C., L.M.B., and F.D.N. performed experiments; A.V., M.G., D.C., Z.K., and F.D.N. analyzed data; A.V., X.C-C., D.J.M., Z.K., D.H.E., and G.Y.O. interpreted results of experiments; A.V. and R.S.N. prepared figures; A.V. drafted manuscript; A.V., R.S.N., D.C., L.M.B., X.C-C., D.J.M., Z.K., F.D.N., D.H.E., and G.Y.O. edited and revised manuscript; A.V., R.S.N., M.G., D.C., L.M.B., X.C-C., D.J.M., Z.K., F.D.N., D.H.E., and G.Y.O. approved final version of manuscript.
REFERENCES
- 1. Wang K, Gheblawi M, Nikhanj A, Munan M, MacIntyre E, O'Neil C, Poglitsch M, Colombo D, Del Nonno F, Kassiri Z, Sligl W, Oudit GY. Dysregulation of ACE (angiotensin-converting enzyme)-2 and renin-angiotensin peptides in SARS-CoV-2 mediated mortality and end-organ injuries. Hypertension 79: 365–378, 2022. doi: 10.1161/HYPERTENSIONAHA.121.18295. [DOI] [PubMed] [Google Scholar]
- 2. Chen L, Li X, Chen M, Feng Y, Xiong C. The ACE2 expression in human heart indicates new potential mechanism of heart injury among patients infected with SARS-CoV-2. Cardiovasc Res 116: 1097–1100, 2020. [Erratum in Cardiovasc Res 116: 1994, 2020]. doi: 10.1093/cvr/cvaa078. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 3. Munoz-Fontela C, Dowling WE, Funnell SGP, Gsell PS, Riveros-Balta AX, Albrecht RA, et al. Animal models for COVID-19. Nature 586: 509–515, 2020. doi: 10.1038/s41586-020-2787-6. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 4. Chan JF-W, Zhang AJ, Yuan S, Poon VK-M, Chan CC-S, Lee AC-Y, Chan W-M, Fan Z, Tsoi H-W, Wen L, Liang R, Cao J, Chen Y, Tang K, Luo C, Cai J-P, Kok K-H, Chu H, Chan K-H, Sridhar S, Chen Z, Chen H, To KK-W, Yuen K-Y. Simulation of the clinical and pathological manifestations of Coronavirus Disease 2019 (COVID-19) in a golden Syrian hamster model: implications for disease pathogenesis and transmissibility. Clin Infect Dis 71: 2428–2446, 2020. doi: 10.1093/cid/ciaa325. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 5. Huang J, Hume AJ, Abo KM, Werder RB, Villacorta-Martin C, Alysandratos KD, Beermann ML, Simone-Roach C, Lindstrom-Vautrin J, Olejnik J, Suder EL, Bullitt E, Hinds A, Sharma A, Bosmann M, Wang R, Hawkins F, Burks EJ, Saeed M, Wilson AA, Mühlberger E, Kotton DN. SARS-CoV-2 infection of pluripotent stem cell-derived human lung alveolar type 2 cells elicits a rapid epithelial-intrinsic inflammatory response. Cell Stem Cell 27: 962–973.e7, 2020. doi: 10.1016/j.stem.2020.09.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 6. Tschöpe C, Ammirati E, Bozkurt B, Caforio ALP, Cooper LT, Felix SB, Hare JM, Heidecker B, Heymans S, Hubner N, Kelle S, Klingel K, Maatz H, Parwani AS, Spillmann F, Starling RC, Tsutsui H, Seferovic P, Van Linthout S. Myocarditis and inflammatory cardiomyopathy: current evidence and future directions. Nat Rev Cardiol 18: 169–193, 2021. doi: 10.1038/s41569-020-00435-x. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 7. Angeles Montero-Fernandez M, Pardo-Garcia R. Histopathology features of the lung in COVID-19 patients. Diagn Histopathol (Oxf) 27: 123–127, 2021. doi: 10.1016/j.mpdhp.2020.11.009. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 8. Oudit GY, Kassiri Z, Jiang C, Liu PP, Poutanen SM, Penninger JM, Butany J. SARS-coronavirus modulation of myocardial ACE2 expression and inflammation in patients with SARS. Eur J Clin Invest 39: 618–625, 2009. doi: 10.1111/j.1365-2362.2009.02153.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 9. Tucker NR, Chaffin M, Bedi KC Jr, Papangeli I, Akkad AD, Arduini A, Hayat S, Eraslan G, Muus C, Bhattacharyya RP, Stegmann CM; Human Cell Atlas Lung Biological Network; Margulies KB, Ellinor PT; Human Cell Atlas Lung Biological Network Consortium Members. Myocyte-specific upregulation of ACE2 in cardiovascular disease: implications for SARS-CoV-2-mediated myocarditis. Circulation 142: 708–710, 2020. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 10. Tavazzi G, Pellegrini C, Maurelli M, Belliato M, Sciutti F, Bottazzi A, Sepe PA, Resasco T, Camporotondo R, Bruno R, Baldanti F, Paolucci S, Pelenghi S, Iotti GA, Mojoli F, Arbustini E. Myocardial localization of coronavirus in COVID-19 cardiogenic shock. Eur J Heart Fail 22: 911–915, 2020. doi: 10.1002/ejhf.1828. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 11. Cheung CY, Poon LLM, Ng IHY, Luk W, Sia S-F, Wu MHS, Chan K-H, Yuen K-Y, Gordon S, Guan Y, Peiris JSM. Cytokine responses in severe acute respiratory syndrome coronavirus-infected macrophages in vitro: possible relevance to pathogenesis. J Virol 79: 7819–7826, 2005. doi: 10.1128/JVI.79.12.7819-7826.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 12. Del Valle DM, Kim-Schulze S, Huang H-H, Beckmann ND, Nirenberg S, Wang B, Lavin Y, Swartz TH, Madduri D, Stock A, Marron TU, Xie H, Patel M, Tuballes K, Van Oekelen O, Rahman A, Kovatch P, Aberg JA, Schadt E, Jagannath S, Mazumdar M, Charney AW, Firpo-Betancourt A, Mendu DR, Jhang J, Reich D, Sigel K, Cordon-Cardo C, Feldmann M, Parekh S, Merad M, Gnjatic S. An inflammatory cytokine signature predicts COVID-19 severity and survival. Nat Med 26: 1636–1643, 2020. doi: 10.1038/s41591-020-1051-9. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 13. Han H, Ma Q, Li C, Liu R, Zhao L, Wang W, Zhang P, Liu X, Gao G, Liu F, Jiang Y, Cheng X, Zhu C, Xia Y. Profiling serum cytokines in COVID-19 patients reveals IL-6 and IL-10 are disease severity predictors. Emerg Microbes Infect 9: 1123–1130, 2020. doi: 10.1080/22221751.2020.1770129. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 14. Viveiros A, Rasmuson J, Vu J, Mulvagh SL, Yip CYY, Norris CM, Oudit GY. Sex differences in COVID-19: candidate pathways, genetics of ACE2, and sex hormones. Am J Physiol Heart Circ Physiol 320: H296–H304, 2021. doi: 10.1152/ajpheart.00755.2020. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 15. South AM, Diz DI, Chappell MC. COVID-19, ACE2, and the cardiovascular consequences. Am J Physiol Heart Circ Physiol 318: H1084–H1090, 2020. doi: 10.1152/ajpheart.00217.2020. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 16. Hoffmann M, Kleine-Weber H, Schroeder S, Kruger N, Herrler T, Erichsen S, Schiergens TS, Herrler G, Wu NH, Nitsche A, Müller MA, Drosten C, Pöhlmann S. SARS-CoV-2 cell entry depends on ACE2 and TMPRSS2 and is blocked by a clinically proven protease inhibitor. Cell 181: 271–280.e8, 2020. doi: 10.1016/j.cell.2020.02.052. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 17. Patel VB, Clarke N, Wang Z, Fan D, Parajuli N, Basu R, Putko B, Kassiri Z, Turner AJ, Oudit GY. Angiotensin II induced proteolytic cleavage of myocardial ACE2 is mediated by TACE/ADAM-17: a positive feedback mechanism in the RAS. J Mol Cell Cardiol 66: 167–176, 2014. doi: 10.1016/j.yjmcc.2013.11.017. [DOI] [PubMed] [Google Scholar]
- 18. Lambert DW, Yarski M, Warner FJ, Thornhill P, Parkin ET, Smith AI, Hooper NM, Turner AJ. Tumor necrosis factor-alpha convertase (ADAM17) mediates regulated ectodomain shedding of the severe-acute respiratory syndrome-coronavirus (SARS-CoV) receptor, angiotensin-converting enzyme-2 (ACE2). J Biol Chem 280: 30113–30119, 2005. doi: 10.1074/jbc.M505111200. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 19. Haga S, Yamamoto N, Nakai-Murakami C, Osawa Y, Tokunaga K, Sata T, Yamamoto N, Sasazuki T, Ishizaka Y. Modulation of TNF-alpha-converting enzyme by the spike protein of SARS-CoV and ACE2 induces TNF-alpha production and facilitates viral entry. Proc Natl Acad Sci USA 105: 7809–7814, 2008. doi: 10.1073/pnas.0711241105. [DOI] [PMC free article] [PubMed] [Google Scholar]
- 20. Viveiros A, Gheblawi M, Aujla PK, Sosnowski DK, Seubert JM, Kassiri Z, Oudit GY. Sex- and age-specific regulation of ACE2: insights into severe COVID-19 susceptibility. J Mol Cell Cardiol 164: 13–16, 2022. doi: 10.1016/j.yjmcc.2021.11.003. [DOI] [PMC free article] [PubMed] [Google Scholar]