Abstract
Preserving optimal mitochondrial function is critical in the heart, which is the most ATP-avid organ in the body. Recently, we showed that global deficiency of the nuclear receptor RORα in the “staggerer” (RORαsg/sg) mouse exacerbates angiotensin II-induced cardiac hypertrophy and compromises cardiomyocyte mitochondrial function. The mechanisms underlying these observations have not been defined. Here we present evidence that RORα regulates cardiomyocyte mitophagy using pharmacological and genetic gain- and loss-of-function tools, including a novel cardiomyocyte-specific RORα knockout mouse. Cardiomyocyte RORα is upregulated by hypoxia and the loss of RORα blunts hypoxia-induced mitophagy and broadly compromises mitochondrial function. We show that RORα is a direct transcriptional regulator of the mitophagy mediator caveolin-3 in cardiomyocytes and that increased expression of RORα increases caveolin-3 abundance and enhances mitophagy. Knockdown of RORα impairs cardiomyocyte mitophagy, but this defect can be rescued by caveolin-3 overexpression. Collectively, these findings reveal a novel role for RORα in regulating mitophagy through caveolin-3 and expand our currently limited understanding of the mechanisms underlying RORα-mediated cardioprotection.
Introduction
The heart consumes more ATP than any other organ by virtue of the requirement for constant cardiomyocyte contraction and relaxation. As such, maintenance of a large and optimally functional pool of mitochondria is essential to normal cardiac physiology. Compromised mitochondrial function leads to energy deprivation and enhanced oxidative stress, both of which are central to the pathobiology of heart failure (Zhou & Tian, 2018). The cardiac mitochondrial pool is preserved through careful orchestration of mitochondrial biogenesis and the clearance of dysfunctional mitochondria, chiefly through selective mitochondrial autophagy, or mitophagy (Kubli & Gustafsson, 2012). Nuclear receptors increasingly are recognized as central transcriptional regulators of these critical processes in the heart (Vega & Kelly, 2017).
The retinoic acid-related orphan nuclear receptor (ROR) subfamily consists of three members, RORα, RORβ, and RORγ (NR1F1-3), with highly tissue-specific and context-dependent expression. RORα plays well-described roles in regulating circadian rhythm, lipid metabolism, and T cell development (Cook et al, 2015). However, there was no recognized function for RORα in the heart prior to two recent reports from Pu and colleagues, who showed that RORα expression is protective in myocardial ischemia/reperfusion and diabetic cardiomyopathy (He et al, 2016; Zhao et al, 2017). We extended those findings in the angiotensin II (Ang II)-induced cardiac injury model, demonstrating that “staggerer” mice that globally lack functional RORα (RORαsg/sg) develop exaggerated myocardial hypertrophy and contractile dysfunction after Ang II exposure (Beak et al, 2019). Using both in vivo and in vitro approaches we made the novel observation that cardiomyocyte RORα deficiency was associated with decreased mitochondrial abundance, ATP depletion, and enhanced oxidative stress. The mechanisms underlying these findings and putative functions for RORα in uninjured cardiomyocytes remain unclear.
Here we use in vivo and in vitro approaches to identify a novel role for RORα in regulating cardiomyocyte mitophagy. The absence of RORα is associated with abnormal mitochondrial structure and function, due at least in part to defects in mitophagy mediated by impaired transcriptional regulation of caveolin-3. These new findings may account for our previous observation of energy deprivation in the hearts of RORαsg/sg mice and emphasize the emerging importance of RORα as a cardioprotective nuclear receptor.
Results
Mitochondria in RORαsg/sg mouse hearts are less abundant and structurally abnormal with fewer mitophagic events
Our recent study demonstrated that RORα protects cardiomyocytes against Ang II-induced hypertrophy by maintaining mitochondrial number and function (Beak et al., 2019). To explore the potential roles for RORα in preserving the cardiomyocyte mitochondrial pool, we examined wild type (WT) and RORαsg/sg hearts using transmission electron microscopy (TEM). We found that the mitochondria in RORαsg/sg hearts were 44% less abundant but 43% larger than in hearts from WT littermates (Figures 1A and B), broadly suggestive of defects in mitochondrial quality control. We also analyzed the TEM images for mitochondria engulfed in double membrane lysosomes, a finding that is indicative of active autophagy, and found 68% fewer of these structures in the RORαsg/sg than the WT hearts. (Figures 1C and 1D). Taken together, these findings suggest that defects in mitophagy may contribute to mitochondrial depletion and dysfunction in the absence of RORα.
Gene expression profiling suggests a defect in mitochondrial quality control in RORαsg/sg mouse hearts
To identify potential regulation of mitochondrial abundance and function by RORα, we performed microarray analysis using Agilent whole genome mouse oligo arrays. We identified 9653 genes that were differentially expressed in RORαsg/sg (n=4) and WT littermate (n=5) hearts. Of these genes, 3118 were >2.5-fold more abundant and 3340 were >2.5-fold less abundant in RORαsg/sg compared with WT hearts.
To analyze these microarray data for regulation of biologically meaningful pathways, we used the online Database for Annotation, Visualization and Discovery (DAVID) v6.8 (Huang da et al, 2009a, b). Filtering for transcripts that were at least 5-fold less abundant in RORαsg/sg hearts revealed a surprising number of pathways related to protein stabilization and degradation (Figure 2A). Transcripts in the Gene Ontology term “Protein Stabilization/Lysosomal protein stabilization” (“Any process involved in maintaining the structure and integrity of a protein and preventing it from degradation or aggregation”) were 2.9-fold less abundant in RORαsg/sg hearts compared to WT hearts (p = 1.2 × 10-7, Figure 2B). Numerous other significantly affected pathways (Supplemental Table 1) were either predicted by the known function of RORα in transcriptional regulation (“DNA templated transcription”, “mRNA processing”) or consistent with our previously published work (Beak et al., 2019) (“Sarcomere organization”, “muscle contraction”, “positive regulation of IκB kinase/NFκB signaling”). Focused analysis of the microarray data demonstrated that the expression of numerous genes associated with mitochondrial function, ATP synthase subunits and autophagy were significantly decreased in the RORαsg/sg mice compared with in WT (Figure 2C). Collectively our microarray analysis supported our published data indicating a role for RORα in mitochondrial maintenance and suggested that defects in protein quality control might contribute to mitochondrial dysfunction in RORαsg/sg mouse hearts.
To further explore at a transcriptional level whether alterations in mitochondrial biogenesis, dynamics, or autophagy might determine the compromised mitochondrial phenotype in the RORαsg/sg mouse hearts, we performed quantitative reverse transcriptase polymerase chain reaction (qRT-PCR) for selected genes critical to each process. Abundance of PGC1α, Gabpa, and TFAM (mitochondrial biogenesis, Figure 2D) was mildly reduced, as was expression of Opa1 (mitochondrial dynamics, Figure 2E). However, the most profound transcriptional differences were found in the autophagy-related genes autophagy and beclin 1 regulator 1 (Ambra1), autophagy related 7 (Atg7), Atg10, and beclin1 (Becn1) (Figure 2F). Using qRT-PCR, we also confirmed decreased abundance of multiple key electron transport genes, further corroborating the presence of a RORα-related mitochondrial defect (Supplemental Figure S1). These findings coupled with the morphological abnormalities seen on TEM (Figure 1) and transcriptome pathway analysis (Figure 2A) encouraged us to determine whether and how RORα regulates cardiomyocyte mitophagy.
RORα regulates expression of mitophagy mediators in normoxic and hypoxic conditions in vivo and in vitro
Mitophagy is carried out primarily through two well-defined pathways: PTEN-induced putative kinase 1 (PINK1) /Parkin and BCL2 interacting protein 3 (Bnip3)/NIX. Activation of either of these pathways can target mitochondria for autophagy by linking to microtubule associated protein light chain beta 3-II (LC3B-II) on autophagosome membranes (Moyzis et al, 2015). To better understand how RORα might affect mitophagy, we immunoblotted the mitochondrial fraction of WT and RORαsg/sg mouse hearts for these canonical mitophagy mediators and found that the absence of functional RORα was associated with markedly decreased abundance of mitochondrial Bnip3, PINK1, and LC3B-II (Figure 3A).
Oxygen tension is a potent regulator of mitochondrial function in the heart and protecting mitochondrial function is critical when the myocardium becomes ischemic (Cadenas et al, 2010). Under hypoxic conditions, such as myocardial infarction, mitophagy is upregulated to promote clearance of dysfunctional mitochondria. Failure to induce mitophagy can lead to exacerbated cardiomyocyte injury and impaired survival (Kubli et al, 2013). To assess the in vivo function of RORα in low oxygen tension, we subjected WT mice to hypoxia (10% O2 for 24 hours). Hypoxia led to nearly 2-fold upregulation of myocardial RORα mRNA expression, similar in magnitude to the classical hypoxia marker, Hif1α (Figure 3B). Immunoblots of heart lysates revealed that hypoxic upregulation of RORα was coupled with significant increases in LC3B and PINK1 expression, consistent with induction of mitophagy (Figure 3C).
Next we sought to determine whether pharmacological manipulation of RORα activity in vitro could recapitulate our genetic loss-of-function findings from our in vivo models. We exposed H9c2 rat ventricular myoblasts to hypoxia (1% O2) in the presence and absence of the RORα inverse agonist SR3335 or agonist SR1078 then quantified the number of phagolysosomes co-stained for LC3B (red) and the lysosomal stain LysoTracker (green) (Figure 3D). The RORα agonist SR1078 increased and the RORα inverse agonist SR3335 decreased phagolysosome events in both normoxia and hypoxia (Figure 3E). We then exposed normoxic and hypoxic neonatal rat ventricular myocytes (NRVMs) treated with SR3335 or SR1078 to bafilomycin A1 (BFA), which interferes with autolysosome acidification and autophagosome formation through inhibition of V-ATPase (Mauvezin & Neufeld, 2015), allowing measurement of autophagic flux as indicated by the ratio of LC3B-II/I (Sharifi et al, 2015). BFA increased the expression of the mitophagy adaptor Bnip3, as well as the LC3B-II/I ratio as expected in vehicle-treated NRVMs. SR3335 abrogated upregulation of Bnip3 and blunted the LC3B-II/I increase (Figure 3F) whereas SR1078 increased LC3B-II (Figure 3G), suggesting that RORα regulates hypoxic autophagic flux
Collectively, these findings suggest that upregulation of RORα in response to hypoxia facilitates cytoprotective autophagy, corroborating our morphological (Figure 1) and transcriptional (Figure 2) findings.
Cardiomyocyte mitophagy is compromised in vivo in a novel cardiomyocyte-specific RORα knockout mouse
The RORαsg/sg “staggerer” mouse is a well-validated genetic loss-of-function model, but it exhibits numerous extracardiac phenotypes that could potentially influence our study of mitophagy in the heart. Though our in vitro studies support cardiomyocyte autonomous functions for RORα, we created a cardiomyocyte-specific RORα knock-out (CMKO) mouse, breeding floxed RORα mice with αMHC-Cre mice, to extend those findings in vivo and to eliminate potential contributions from extracardiac phenotypes. Cre mRNA was detected only in the heart and RORα mRNA abundance was 80% lower in the CMKO than CMWT (Cre−flox+/+) hearts, expression in liver and kidney was unaffected, validating our breeding strategy. We found no compensatory changes in RORγ transcriptional expression (Figure 4A).
We then examined the effect of cardiomyocyte-specific RORα deletion on the in vivo expression of several proteins that are centrally involved in the induction and maintenance of mitophagy. RNA was isolated from hearts of CMWT and CMKO mice that were housed in normoxic (21% O2) or hypoxic (10% O2) conditions for 8 hours. Abundance of the transcripts encoding Atg10 and Atg7, autophagy receptor p62, and PINK1 was higher in hypoxic than normoxic CMWT hearts. However, the hypoxia-induced increase in these critical autophagy mediators was not observed in CMKO mouse hearts (Figure 4B). Immunoblotting demonstrated the expected increases in Parkin, p62, Bnip3, and LC3B-II in hypoxic CMWT hearts but these changes were blunted in CMKO hearts (Figure 4C). We next sought to examine the functional consequences of these differences, using immunofluorescent staining (Figure 4D) for LC3B (red) and the mitochondria marker heat shock protein-60 (Hsp60, green). In normoxic conditions, CMKO hearts exhibited 75% fewer mitophagy events than CMWT hearts (defined by LC3B and Hsp60 co-staining). Hypoxia induced an increase in mitophagy in both CMWT and CMKO hearts, but mitophagosomes were 70% less abundant in CMKO than CMWT (Figure 4E).
Collectively, these results substantiate our findings that RORα regulates mitophagy in the heart. They further indicate that the deficits in mitophagy exhibited by the global RORαsg/sg mouse most likely arise from cardiomyocyte autonomous effects of RORα.
Loss of RORα compromises hypoxic mitophagy and mitochondrial function in vitro
The previous findings encouraged us to further analyze the effect of RORα on mitophagy and mitochondrial function in hypoxic cardiomyocytes. To measure mitophagy in neonatal rat ventricular myocytes (NRVMs), we used mitochondrial-targeted Keima (mt-Keima), a fluorescent protein pH indicator that fluoresces red (543 nm) when introduced to the acidic environment of the lysosome but green (458 nm) at normal or basic pH (Shirakabe et al, 2016).
We infected NRVMs with lentivirus containing either short hairpin RNA (shRNA) against RORα or scrambled shRNA (shCtrl) and found that the mt-Keima fluorescence ratio (543/458 nm) underwent the expected increase under hypoxic conditions but was decreased in both normoxia and hypoxia by RORα knockdown (Figure 5A).
To expand upon our earlier in vivo discovery that RORαsg/sg mouse hearts have decreased ATP abundance (Beak et al., 2019), we used lentiviral shCtrl or shRORα in NRVMs for an in vitro loss-of-function approach. Knockdown of RORα in NRVMs decreased the functional mitochondrial mass, as measured by citrate synthase (Figure 5B) and succinate dehydrogenase (Figure 5C) activity. Oxygen consumption rate was markedly decreased by shRORα (Figure 5D). These metabolic insults collectively contributed to decreased ATP abundance in the absence of functional RORα (Figure 5E). Knockdown of RORα compromised mitochondrial membrane potential as measured by JC-1 (Figure 5F) and increased mitochondrial reactive oxygen species (ROS) generation as assayed by MitoSox (Figure 5G). Cell death, as measured by LDH release, was only modestly increased by shRORα (Figure 5H). Taken together, these findings further indicate that RORα contributes to optimal mitochondrial function in both normoxia and hypoxia, likely through maintenance of mitophagy.
RORα directly binds the caveolin-3 promoter to regulate hypoxic mitophagy
Caveolin-3 (Cav-3) enhances autophagy and protects mitochondrial function in hypoxic HL-1 cardiomyoblasts (Kassan et al, 2016). Hence, we hypothesized that RORα promotes mitophagy in the heart by regulating the expression of Cav-3.
Analysis of Cav-3 protein demonstrated that the level of Cav-3 protein was 34% lower in RORαsg/sg than WT hearts (Figure 6A). In the cardiomyocyte-specific RORα CMKO, Cav-3 mRNA (Figure 6B) and protein (Figures 6C and 6D) levels were similar to WT hearts in normoxic conditions. After WT and CMKO mice spent 8 hours in a hypoxia chamber (10% O2), Cav-3 mRNA and protein were increased 2-3-fold in WT hearts, but there was no change in Cav-3 abundance in CMKO hearts. These findings suggested that RORα may regulate Cav-3 expression in cardiomyocytes. Through promoter analysis, we found two consensus RORα binding sites (ROREs) in the proximal promoter of Cav-3. To determine whether the Cav3 gene is a direct transcriptional target gene of RORα in adult mouse ventricular myocytes, we performed chromatin immunoprecipitation (ChIP)-PCR using immunopulldown with an anti-RORα antibody or an anti-IgG antibody as a negative control. We found significant enrichment of the regions containing the two ROREs (−0.1 kb, and −0.5 kb regions), but not in a region of the promoter that does not contain an RORE (−4.2kb), indicating that RORα regulates Cav3 directly (Figure 6E).
Next, we examined the effect of RORα expression on Cav-3 expression in H9c2 cardiomyoblasts infected with a lentivirus containing doxycycline-inducible RORα4 (H9c2-Flag-RORα4-HA), the predominant RORα isoform in the heart (Supplemental Figure S2A). Doxycycline induced RORα4 in a dose-dependent manner (Supplemental Figure S2B) and RORα4 was localized largely to nuclei, as expected (Supplemental Figure S2C). The induction of RORα by doxycycline resulted in a 4.5-fold increase in Cav-3 expression (Figure 7A). The expression of Cav-3 was then compared in doxycycline-treated and untreated H9c2-Flag-RORα4-HA cells exposed to either hypoxia (1% O2) or normoxia (21% O2). We found that hypoxia enhanced Cav-3 expression, and this upregulation was significantly greater in doxycycline-treated cells in which RORα was induced (Figures 7B and 7C).To determine whether this increase in Cav-3 expression correlated with enhanced mitophagy we quantified co-localization of immunofluorescent LC3B (red) with mitochondria (green) and identified roughly 4 times more mitophagy events in the cells in which RORα was induced by doxycycline (Figure 7D). To determine whether Cav-3 is directly involved in the regulation of cardiomyocyte mitophagy by RORα we knocked down RORα in H9c2 cardiomyoblasts with shRNA then transfected these cells with an EGFP-Cav-3 plasmid. We then visualized mitophagy events by co-staining for LC3B (blue) and mitochondria (red), using EGFP to identify Cav-3 (Figure 7E). As expected, shRORα decreased mitophagy events, however forced expression of Cav-3 eliminated this defect such that mitophagy events were equivalent in shCtrl-Cav-3+ and shRORα-Cav3+ H9c2s. These results support the hypothesis that RORα directly regulates the expression of Cav-3 in cardiomyocytes and that RORα-mediated Cav-3 induction promotes mitophagy. Taken together, these findings provide a potential mechanism for the mitochondria-protective effects of RORα in the heart.
Discussion
In this study, we identify a novel role for the nuclear receptor RORα in the regulation of mitophagy, in part through transcriptional upregulation of Cav-3. Loss of RORα profoundly compromises cardiomyocyte mitochondrial function leading to energy deprivation, oxidative stress and cell death. These insults are particularly deleterious in the setting of hypoxia. The heart is the most ATP-avid organ in the body, necessitating optimal mitochondrial capacity. Nuclear receptors such as the peroxisome proliferator activated receptor (PPAR) and estrogen related receptor (ERR) families are central regulators of cardiomyocyte mitochondrial structure and function (Vega & Kelly, 2017), and the present findings identify RORα as an additional nuclear receptor that contributes to maintaining cardiac mitochondrial health. A role for RORα in regulating mitochondrial fission in the liver was proposed previously (Kim et al, 2017), but to our knowledge our present study is the first to demonstrate that RORα adaptively regulates mitophagy in any tissue.
We chose to use hypoxia as an in vivo and in vitro stressor to test the role of RORα in the cardiac response to stress because hypoxia is known to stimulate mitophagy and because the cellular response to hypoxia is critical to important clinical conditions such as ischemia-reperfusion injury and myocardial infarction. We found that RORα was upregulated by hypoxia in cardiomyocytes (Figures 3B and C) and that RORα knockdown broadly compromised cardiomyocyte mitochondrial function under hypoxic conditions (Figure 5). We also show that our novel mice with a cardiomyocyte-specific RORα deletion exhibit a failure to induce hypoxic mitophagy in vivo (Figure 4). Though the role of RORα in the cardiac response to hypoxia has not been studied previously, a complex interaction has been demonstrated in other tissues between RORα and hypoxia induced factor-1α (HIF-1α), the canonical regulator of cellular response to low oxygen tension. RORα regulates the transcription of hypoxia HIF-1α in endothelial cells (Kim et al, 2008). Interestingly, hypoxia increases RORα transcription in a hepatoma cell line by binding to an HRE (hypoxia response element) in the RORα promoter (Chauvet et al, 2004), indicating the potential for reciprocal regulation of RORα and HIF-1α. In future studies we will explore whether RORα regulates other critical elements of the cardiomyocyte response to hypoxia.
Our previous work demonstrated that RORα mitigated angiotensin II-induced cardiac hypertrophy through the regulation of cardiomyocyte STAT3 signaling (Beak et al., 2019). Here we identify a novel role for RORα in promoting cardiomyocyte mitophagy through enhancing Cav3 transcription. Like the other two members of the caveolin family of proteins, Cav-3, the predominant caveolin isoform in muscle cells, serves primarily as a scaffold for membrane trafficking and signaling within caveolae (Tang et al, 1996). However, Cav-3 also dissociates from caveolae to the cytosol in injured cardiomyocytes (Ratajczak et al, 2003) and to the mitochondrial membrane where it enhances respiratory capacity and mitigates oxidative stress (Fridolfsson et al, 2012). Cardiac specific overexpression of Cav-3 protects against hypertrophy induced by pressure loading (Horikawa et al, 2011) and ischemia-reperfusion injury (Tsutsumi et al, 2008). Cav-3 knockout mice exhibit impaired cardiomyocyte mitochondrial function (Fridolfsson et al., 2012) and enhanced susceptibility to injury (Horikawa et al, 2008). Recognized mechanisms for these protective effects include preservation of t-tubular calcium flux(Kong et al, 2019) and prevention of hypoxia-induced apoptosis.(Zhou et al, 2018)
As a nuclear receptor, RORα is thought to exert most of its effects through transcriptional regulation. RORα regulates caveolin-3 expression by directly binding the Cav3 promoter in skeletal muscle (Lau et al, 2004). The importance of caveolins in regulating autophagy has been established in cardiomyoblasts and other cell types. Cav-3 is upregulated by simulated ischemia-reperfusion in HL-1 cardiomyoblasts and knockdown of Cav-3 in HL-1s compromises autophagy and mitochondrial function, contributing to enhanced cell death.(Kassan et al., 2016) Cav-1 interacts with beclin-1 to induce autophagolysosome formation and protect against cerebral oxidative stress and ischemic injury (Nah et al, 2017). Here we show that RORα binds two sites in the Cav3 promoter to regulate Cav-3 expression (Figure 6E). Cav-3 abundance is lower in the hearts of RORαsg/sg mice (Figures 6A and 6B) and cardiomyocyte-specific RORα deletion is associated with a failure of the expected hypoxic induction of Cav-3 (Figures 6C and 6D). We also demonstrate that RORα overexpression in cardiomyocytes induces Cav-3 expression (Figures 7A-C) and promotes mitophagy (Figure 7D) in normoxic and hypoxic conditions. Forced expression of Cav-3 rescues mitophagy defects after RORα knockdown in cardiomyocytes (Figure 7E). Taken together these findings suggest that RORα and Cav-3 might represent part of a coordinated and protective response to hypoxia in the heart that includes enhanced mitophagy.
We began these investigations to expand upon our earlier finding that the global absence of functional RORα was associated with decreased mitochondrial number and ATP depletion in the heart (Beak et al., 2019). Here we show that knocking down RORα in cardiomyocytes reduces citrate synthase activity and respiratory capacity while compromising mitochondrial membrane potential and increasing oxidative stress, the first detailed characterization of how RORα affects mitochondrial function. Though our central conclusion is that RORα regulates mitophagy to preserve the cardiomyocyte mitochondrial pool, we also present evidence that RORα may be involved in other aspects of mitochondrial homeostasis. We show that the expression of several key regulators of mitochondrial biogenesis, PGC1α, Gabpa, and TFAM, was lower in RORαsg/sg than WT hearts (Figure 2D). PGC1α is more abundant in RORαsg/sg liver cells (Lau et al, 2008), but less abundant in skeletal muscle cells lacking functional RORα (Lau et al., 2004), indicating that this regulation is tissue-dependent. We also show that Opa1, a canonical regulator of mitochondrial fission, is reduced in RORαsg/sg hearts (Figure 2E), consistent with a previous report that RORα regulates mitochondrial dynamics in hepatocytes (Kim et al., 2017). As such, it is possible that induction of mitophagy alone does not account for the protective effects of RORα on mitochondrial function. Indeed, in future studies we will explore potential roles for RORα in the regulation of mitochondrial biogenesis and/or dynamics in cardiomyocytes.
In summary, our experiments indicate that RORα protects cardiomyocyte mitochondrial function through regulation of mitophagy. These findings provide mechanistic underpinning for recent studies demonstrating an emerging cardioprotective role for RORα in the setting of ischemia-reperfusion, diabetic cardiomyopathy and angiotensin II-induced heart failure. The RORα agonist SR1078 recently was shown to be well-tolerated and efficacious in an animal model of autism (Wang et al, 2016) and our present study indicates that this agonist is active in normoxic and hypoxic cardiomyocytes (Figures 3E and G). Taken together, these studies suggest that activation of RORα in vivo could represent a novel therapeutic approach to heart disease.
Sources of funding: BCJ: NIH (R01HL140067); Hugh A. McAllister Research Foundation. AMJ: Intramural Research Program of the National Institute of Environmental Health Sciences, the National Institutes of Health [Z01-ES-101586].
Materials and Methods
Experimental animals
Heterozygous RORαsg/sg mice on a C57BL/6J background were purchased from Jackson lab and maintained as previously described (Beak et al., 2019). Homozygous RORαsg/sg mice, the products of heterozygous breeding, and WT littermates were used in all experiments at 12-16 weeks of age. Homozygous cardiomyocyte-specific RORα knockout mice (RORα CMKO) were generated by crossing floxed RORα mice with LoxP sites flanking exon 9 (Anton Jetten, NIEHS, Durham, NC) with αMHC-Cre mice (Dale Abel, University of Iowa). All animal studies followed the NIH Guide for the Care and Use of Laboratory Animals and animal protocols were approved by the NIEHS Animal Care and Use Committee and the University of North Carolina Institutional Animal Care and Use Committee.
Transmission Electron Microscopy (TEM)
Cardiac sections were observed with a LEO EM910 TEM operating at 80 kV (LEO Electron Microscopy, Thornwood, NY) and photographed with a Gatan Orius SC1000 CCD Digital Camera and Digital Micrograph 3.11.0 (Gatan, Pleasanton, CA). Mitochondrial cross-sectional area was measured using NIH ImageJ at magnifications of ×5000 to ×10,000. The global scale was set according to the image-specific scale generated by the Gatan camera output. An average of 2500 mitochondria were analyzed from 30 fields from multiple levels from three hearts per mouse cohort.
Microarray analysis
Microarray analysis was performed to compare cardiac gene expression between WT (n=5) and RORαsg/sg (n=4) mice. RNA was amplified using NuGEN Ovation Pico WTA System (NuGEN). Gene expression analysis was conducted using Agilent Whole Mouse Genome 4×44 multiplex format oligo arrays (Agilent Technologies 014868) following the Agilent 1-color microarray-based gene expression analysis protocol. Starting with 500ng of total RNA, Cy3 labeled cRNA was produced according to manufacturer’s protocol. For each sample, 1.65ug of Cy3 labeled cRNA was fragmented and hybridized for 17 hours in a rotating hybridization oven. Slides were washed and then scanned with an Agilent Scanner. Data was obtained using the Agilent Feature Extraction software (v9.5), using the 1-color defaults for all parameters. The Agilent Feature Extraction Software performed error modeling, adjusting for additive and multiplicative noise. The resulting data were processed using the Rosetta Resolver® system version 7.2 (Rosetta Biosoftware, Kirkland, WA). In order to identify differentially expressed probes, analysis of variance (ANOVA) was used to determine if there was a statistical difference between the means of groups. In addition, we used a multiple test correction to reduce the number of false positives. Specifically, an error weighted ANOVA and Benjamini-Hochberg multiple test correction with a p value of p<0.01 was performed using Rosetta Resolver (www.rosettabio.com). The data presented in this publication have been deposited in the NCBI Gene Expression Omnibus at http://www.ncbi.nlm.nih.gov/geo/#GSE98892.
Hypoxia chamber
Mice were housed in a hypoxia chamber (Oxycycler) with internal dimension (76×51×51 cm) sufficient to hold 4 cages with 4 mice per cage as previously described (Zhang et al, 2020). Inflow rate (nitrogen mixed with oxygen or room air) was ~3.1 ft3/hr and 4 holes (0.7 cm diameter) were opened at the bottom of each of the chamber’s 3 sides. Drierite (#21909-5000, Acros, Fair Lawn, NJ) and soda lime (#36596, Alfa Aesar, Haverhill, MA) were placed at the bottom of the chamber in 12×10×5 cm trays (~260g and 200g, respectively). CO2 was measured using a Cozir Wide-Range sensor and GasLab software, (#CM-0123 CO2meter.com, Ormond Beach, Fl). The hypoxia chambers have internal fans to prevent gradients in composition of the atmospheres. CO2 was automatically recorded at 5 min intervals. FIO2 was lowered from 21% to 10% (hypoxia) in ~3 h. Control (21% O2, normoxia) mice were maintained in an identical chamber. Hypoxia chambers were generously provided by Dr. James Faber.
Histology and immunofluorescence microscopy
Mice were heparinized and the heart was perfused with 10 mL PBS followed by 20 mL of 4% PFA/PBS through a 23G butterfly needle, then excised and placed in 4% PFA/PBS for 24h prior to transfer to 70% EtOH. For immunohistochemistry, hearts were fixed overnight in 4% PFA/PBS, incubated in 30% sucrose/PBS, then frozen in O.T.C. medium (Tissue-Tek, Hatfield, PA). Frozen sections (10 microns), obtained with a Leica cryostat (Leica, Buffalo Grove, IL), were placed on glass slides, dried at room temperature, and then incubated with primary antibodies. After washing, the sections were incubated for 3 hours at room temperature with anti-mouse, anti-rabbit, anti-goat, or anti-rat Alexa Fluor-488, Fluor-594 or Fluor-647 conjugated secondary antibodies (1:1,000, Life Technologies, Grand Island, NY). Fluorescence was observed with a Zeiss LSM880 confocal microscope.
Quantitative real-time reverse transcriptase PCR (qRT-PCR)
Total mouse heart RNA was isolated with an RNeasy mini kit (Qiagen, Valencia, CA) or RNAqueous® Micro RNA isolation kit (Ambion, Austin, TX) following the manufacturer’s instructions. RNA was reverse-transcribed using a High-Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA). qRT-PCR reactions were carried out in triplicate in a LightCycler® 480 System (Roche, Indianapolis, IN, USA) using either probe/primer sets or SYBR Green I (see Primer List). Relative quantitation of PCR products used the ΔΔCt method relative to two validated reference genes (Tbp1 and Polr2a). All probes and primers were from Roche or Thermofisher.
Immunoblotting
Whole tissue or cell lysates were produced in RIPA buffer supplemented with PhosSTOP (Roche Diagnostics Corporation, Indianapolis, IN, USA) and protease inhibitor cocktail (Roche Diagnostics Corporation). Subsequently samples were incubated in 4× LDS sample buffer with 2% β-mercaptoethanol, for 10 min at 70 °C. SDS–PAGE and immunoblotting were performed using the 4-12% NuPage gel system (Life Technologies, Foster City, CA, USA). Membranes were blocked in 5% milk/TBS-Tween, incubated in primary antibody overnight at 4°C, then secondary HRP-conjugated antibodies for 1h at room temperature. Images were generated using Amersham ECL Select Western Blotting Detection Reagent (GE Healthcare life sciences, Marlborough, MA, USA) and the MultiDoc-It™ Imaging System (UVP gel image system, Upland, CA, USA).
Antibodies
RORα (GTX100029, 1:1000), SQSTM1/p62 (GTX629890, 1:1000), PINK1 (GTX107851, 1:1000), LC3B (GTX127375, 1:1000), Ambra1 (GTX55507, 1:1000), and HIF1α (GTX628480) were from Genetex (Irvine, CA, USA). VDAC1 (#4661, 1:000), HA (#3724, 1:1000), LC3B (#83506, 1:500), and Hsp60 (#12165, 1:1000) were from Cell Signaling Technology (Danvers, MA, USA). GAPDH (MAB374 clone 6C5, 1: 10,000, Millipore), Caveolin-3 (sc-5310, 1:1000), Parkin (sc-32282, 1:1000), and Bnip3 (sc-56167, 1:1000) were from Santa Cruz Biotechnology (Dallas, TX, USA). Polyclonal goat anti-rabbit IgG–horseradish-peroxidase (HRP; A9169, 1:5,000), polyclonal rabbit anti-mouse IgG–HRP (A9044, 1:5,000), polyclonal rabbit anti-goat IgG-HRP (A5420, 1:5,000) were from Sigma Aldrich (St. Louis, MO, USA).
Neonatal rat ventricular myocyte (NRVM) cultures, immunocytochemistry and lentiviral infections
Female Sprague-Dawley rats and newborn litters were from Charles River Laboratories (Wilmington, MA, USA). NRVMs were isolated as previously described (Simpson, 1983). Experiments were carried out after 36-48 hours of serum starvation in the presence of insulin, transferrin, and BrdU. NRVM hypoxia was induced by incubation in a 37°C hypoxia chamber with 1% O2 after infection with lentivirus containing either scrambled short hairpin RNA (shControl) or shRNA specifically targeting rat RORα (iO51217 or iV051217, abm, Richmond, BC, Canada).
shRORα target sequences: TGTCATTACGTGTGAAGGCTGCAAGGGCT, ACCTACAACATCTCAGCCAATGGGCTGAC, GGACTGGACATCAATGGGATCAAACCCGA, AGAGGTGATGTGGCAGTTGTGTGCTATCA
Mitochondrial fractionation
Mitochondria were isolated from frozen heart tissue using a Mitochondria isolation kit (ab110168, abcam). Tissue was briefly washed in isolation buffer, dried with Whatman filter paper, weighed, placed in a glass beaker, thoroughly minced then homogenized with a Dounce homogenizer. The homogenate was centrifuged at 1,000g for 10 min at 4°C. The supernatant was centrifuged at 12,000g for 15 min at 4°C then saved as crude cytosolic and nuclear fractions and the pellet was centrifuged again at 12,000g for 15min after resuspending in isolation buffer and protease inhibitor for further mitochondrial purification. The homogenized pellets were re-suspended in isolation buffer with protease and phosphatase inhibitor cocktails (Roche, Indianapolis, IN). The protein concentration was measured by BCA protein assay kit (Thermo Scientific, Waltham, MA).
ATP bioluminescence assay
ATP content was determined using a luciferin-luciferase ATP assay (ThermoFisher Scientific A22066) according to the manufacturer’s protocol. All reagents were placed in ice before the experiments were carried out. The ATP solution (100 nM) was prepared by mixing 50μl of a stock solution (5 mM) with 950 μl of deionized water (DW), and buffer in a microcentrifuge tube. The mixture was then transferred to a 96-well microplate for measurement of ATP bioluminescence. In brief, 100 μl of the ATP solution was mixed with 100 μl of the luciferin–luciferase reagent (10 mg/ml; reconstituted with DW) in each well of the microplate. ATP bioluminescence was measured immediately using a microplate luminometer. The integration time of the luminometer was set at 1 s with normal gain.
Mitochondrial membrane potential
Mitochondrial membrane potential in NRVMs was determined by 5, 5′, 6, 6′-tetrachloro-1, 1′, 3, 3′-tetraethylbenzimidazolylcarbocyanine iodide (JC-1) reduction. Cells were incubated with JC-1 (Cambridge, MA, Abcam) according to the manufacturer's protocol. In brief, serum-starved NRVMs were incubated in the normoxia chamber with 21% O2or in the hypoxia chamber with 1% after infection with shControl or shRORα. JC-1 2 μM was added to each well for 30 min. Cells were washed once with medium then analyzed by plate reader (CLARIOstar, BMG LABTECH, Germany). JC-1 green fluorescence was excited at 488 nm and emission was detected using a 530 ± 40 nm filter. JC-1 red fluorescence was excited at 488 nm and emission was detected using a 613 ± 20 nm filter.
Succinate dehydrogenase (SDH) activity assay
SDH activity was measured using an SDH Activity Colorimetric Assay Kit (MBS2540432) purchased from MyBiosource Co. (San Diego, CA). After correcting for the background, SDH activity in NRVM lysates was determined.
LDH release assay
For measurement of cytotoxicity, NRVMs were seeded onto 96-well plates. After treatment, 50 μl of culture medium from each well was collected and transferred to another 96-well culture plate. 50 μl of LDH reagent (Thermo Fisher scientific, C20300) was then added to each well and the solution incubated at room temperature for 30 min. After the reaction was complete, absorbance at 490 nm was measured by an ELISA reader.
Citrate synthase activity
Citrate synthase activity assay kit (MAK 193, Millipore sigma, St. Louis, MO) was used in NRVMs incubated at 21% or 1% O2 after infection with shControl- or shRORα-lentivirus.
Mitochondrial ROS measurement
The production of superoxide by the mitochondria of NRVMs incubated at 21% or 1% O2 after infection with shControl- or shRORα-lentivirus was measured using MitoSOX™ Red mitochondrial superoxide indicator (M36008, Thermo Fisher scientific).
Oxygen consumption rate (OCR) assay
OCR was measured using an extracellular oxygen consumption assay kit (ab197243; Abcam, Cambridge, UK) following the manufacturer’s protocol. NRVMs in a 96-well black plate were incubated at 21% or 1% O2 after infection with shControl- or shRORα-lentivirus in triplicates for each condition. A negative control was prepared with reaction buffer without cells. All wells were covered with two drops of pre-heated mineral oil to block the back diffusion of oxygen. OCR was calculated based on fluorescent signals (excitation of 380 nm/emission of 650 nm), measured in 1.5 min intervals for 1.5 h using a plate reader (CLARIOstar, BMG LABTECH, Germany). The experiment was repeated two times independently.
Keima with Mitochondrial Localization Signal
Keima with mitochondrial localization signal (Mito-Keima), a mitochondrially localized pH-indicator protein (Katayama et al, 2011), was generously supplied by Junichi Sadoshima (Rutgers New Jersey Medical School). The method used to detect lysosomal delivery of Mito-Keima has been described (Shirakabe et al., 2016). Briefly, NRVMs were infected with shControl- or shRORα-lentivirus for 24 hr then infected with mtKeima-virus for visualizing mitophagy (543 nm and 458 nm) under fluorescent microscopy. Relative fluorescent intensity (ratio of 543nm/458nm) was determined by Image J.
Treatment with RORα inverse agonist (SR3335) and agonist (SR1078)
NRVMs were treated with vehicle or RORα inverse agonist, SR3335 20μM (Kumar et al, 2011) or agonist, SR1078 2μM(Wang et al., 2016) for 6 hr and then incubated in a hypoxia chamber (1% O2) after pretreatment with BFA (100nM) for 2 hr.
Chromatin immunoprecipitation-PCR
To demonstrate direct transcriptional regulation of Caveolin-3 by RORα, adult cardiomyocytes isolated from 3-4 month old mice were washed in PBS containing protease and phosphatase inhibitor cocktail (PIC; Thermo Scientific) and then fixed in 1% formaldehyde at 25°C for 8 min for cross-linking and followed by quenching by 0.125 M glycine at 25°C for 8 min as previously described. (Beak et al., 2019) The amount of chromatin immunoprecipitated DNA (ChIPed-DNA) relative to each input DNA was determined by quantitative PCR in triplicate with primers amplifying consensus RORE (AGGTCA) or a nontargeting sequence in the mouse Caveolin-3 promoter.
Chromatin immunoprecipitation (ChIP)-PCR primers were:
Caveolin-3-4.2k, forward 5’-GCCAAGAGGGATCAGCAATA-3’ and reverse 5’-CAAGGGAAGGCATTTGACAT-3’;
Caveolin-3-0.5k, forward 5’-CCCCATGGCCTTAGTTTGAT-3’ and reverse 5’-TTATTTGCGTGCAAGTGAGC-3’;
Caveolin-3-0.1k, forward 5’-CAGGGTGGGAAGACTCTTGA-3’ and reverse 5’-GTCCCAGCTCTGTCTCTTGC-3’.
Generation of doxycycline-inducible H9c2 cell line and immunocytochemistry
An H9c2 cell line with doxycycline-inducible RORα4 was adapted from a published report. (Jeon et al, 2019) Briefly, RORα4 tagged with N-terminus FLAG and C-terminus HA (FLAG-RORα4-HA) was inserted into lentiviral vector pIND20, and viral soup was generated in HEK293T cells. H9c2 cells were infected with lentiviral pIND20-FLAG-RORα4-HA and stable cells were selected by 500 μg/ml of G418. Induction of RORα4 was examined by Western blot using HA antibody after treatment of doxycycline. For the Cav-3 rescue experiment, we infected H9c2s with shRNA against RORα or scrambled shRNA for 48hrs then transfected with an EGFP-Cav-3 plasmid (Addgene plasmid #68396) or empty EGFP plasmid and an mRaspberry-Mito-7 plasmid (Addgene #55931) to localize mitochondria. For visualization of mitophagy, H2c2s were stained with LC3B and LysoTracker (Thermo Fisher Scientific, L7525).
Statistics
All results are presented as mean ± SEM. Comparisons were made in GraphPad Prism (San Diego, CA) using unpaired t-test (2 groups) or two-way ANOVA (Type III sum of squares) with Tukey’s post-hoc analysis (4 groups).