Introduction
Autophagy is a ubiquitous catabolic process by which cells engulf and deliver damaged proteins and organelles to lysosomes for degradation, maintaining cellular homeostasis (Rubinsztein et al,
2011; Yamaguchi,
2019). Autophagy has been extensively implicated in regulating cardiac structure and function under various physiological and pathophysiological conditions, such as ischemic heart disease, heart failure, and heart hypertrophy (Bhuiyan et al,
2013; Kassiotis et al,
2009; Lavandero et al,
2015; Nakai et al,
2007; Riehle et al,
2013; Sciarretta et al,
2012; Xie et al,
2014), and altered autophagy has been involved in the development of some inherited cardiomyopathies, such as Danon’s disease in which autophagy is impaired secondary to the deficiency of lysosome-associated membrane protein 2 (LAMP-2) (Nascimbeni et al,
2017), left ventricular noncompaction (LVNC) secondary to PLEKHM2 mutations (Muhammad et al,
2015), and desmin-related cardiomyopathy (DRM) secondary to desmin mutations (Bhuiyan et al,
2013). In addition, impaired autophagy has been demonstrated to directly induce cardiomyopathy (Nakai et al,
2007; Taneike et al,
2010). As such, the mechanisms underlying the alteration of autophagic flux have been extensively investigated in hearts under various physiological and pathophysiological conditions (Sciarretta et al,
2015; Zhang et al,
2017). Although impaired mitophagy and protein quality control may contribute to mitochondrial and cardiac dysfunction, a gap in knowledge remains concerning mechanisms linking impaired autophagic flux and the development of cardiac dysfunction.
An important proposed mechanism underlying cardiac dysfunction in the setting of autophagic defects is the accumulation of damaged cytosolic materials, including organelles, that directly induce cellular cytotoxicity leading ultimately to cell death. Accumulation of damaged proteins (misfolded, polyubiquitinated, oxidized, or aggregated) impairs their function and those of other proteins to alter homeostatic signaling, metabolites, and ion fluxes (Bhuiyan et al,
2013; Nakai et al,
2007; Taneike et al,
2010). Dysfunctional mitochondria that accumulate in the face of reduced mitophagy induce ROS production and release of cytochrome
c to initiate apoptotic cell death (Bhuiyan et al,
2013; Dhingra et al,
2019; Nakai et al,
2007; Taneike et al,
2010; Wu et al,
2009). Increased release of mtDNA may activate inflammatory responses and cell death pathways in pressure-overloaded hearts, contributing to the development of heart failure (Oka et al,
2012; Wu et al,
2009).
In addition to removing damaged cytosolic materials, autophagy also selectively degrades specific protein(s) involved in metabolism or signal transduction by binding to the SQSTM1-motif that targets these peptides to the autophagic machinery. Pleiotropic functions of SQSTM1, also known as the ubiquitin-binding protein p62, have been described, including binding to MEKK3 and Raptor to regulate MTORC1 signaling, TRAF6 to regulate NF-κB signaling, MAP1LC3A (LC3) to regulate autophagic loading, and KEAP1 to regulate NRF2 signaling (Moscat et al,
2016). Impaired autophagy has been linked to altered function of these signaling pathways. In addition, impaired autophagy has been shown to deplete NAD
+ levels and thus cause cell death (Kataura et al,
2022; Wilson et al,
2023). This therefore leaves open the possibility that in addition to accumulation of damaged cytosolic materials, the disruption of the above-mentioned mechanisms could contribute to mitochondrial and cardiac dysfunction in the setting of autophagic deficiency. These additional mechanisms have not to date been rigorously evaluated in the context of heart failure.
In this study, we found that in the setting of impaired autophagy induced by ATG3 deficiency, hearts developed NAD+ deficiency attributed to accelerated NAD+ clearance, but not to decreased NAD+ salvage, de novo synthesis, or increased NAD+ consumption, which contributes to the development of mitochondrial and cardiac dysfunction and resultant heart failure. The acceleration of NAD+ clearance is due to induction of nicotinamide N-methyltransferase (NNMT) attributable to the activation of SQSTM1-NF-κB signal transduction. These data indicate that autophagy maintains mitochondrial and cardiac function in part by mediating SQSTM1-NF-κB signal transduction to control cellular levels of NAD+, a central gauge of nutrient status.
Discussion
In this study, we demonstrate that reducing autophagic flux accelerates NAM clearance leading to NAD+ deficiency, which results in mitochondrial and cardiac dysfunction in the heart. Specifically, impaired autophagic flux induces SQSTM1 accumulation that activates NF-κB signal transduction. The activated NF-κB binds the NNMT promoter to initiate NNMT transcription, which eventually increases NNMT protein content. As a key enzyme of NAD+ clearance, NNMT induction reduces NAD+ levels and antagonizes salvage synthesis of NAD+ from NAM, which precipitates mitochondrial and cardiac dysfunction.
Autophagic flux is a central homeostatic mechanism that maintains cellular homeostasis by degrading materials toxic to cardiomyocytes (Rubinsztein et al,
2011; Yamamoto et al,
2014). The detrimental effects of impaired autophagy on mitochondrial and cardiac function have been largely attributed to the accumulation of damaged proteins and organelles. Thus, most efforts have focused on why and how a defect in autophagic flux may occur under various pathophysiological conditions (Sciarretta et al,
2015; Zhang et al,
2017). Our study focused on identifying mechanisms arising from the impaired autophagic flux that could be independent of impaired cellular quality control in cardiomyocytes. By focusing on NAD
+ metabolism in cardiomyocytes, we observed that impaired autophagic flux reduces NAD
+ availability in cardiomyocytes, secondary to increased degradation, which underlies mitochondrial and cardiac dysfunction. Our data also supports a model in which autophagic flux via directional changes in SQSTM1 expression provides a novel regulatory circuit linking autophagy activation (usually induced by nutrient deficiency), with the regulation of NAD
+, an important sensor of cellular nutrient status.
NAD
+ functions as a cellular electron carrier in multiple metabolic pathways. It is also involved in other cellular processes, such as posttranslational modifications of proteins, for example, SIRT1-mediated deacetylation (Hsu et al,
2014). In failing hearts, NAD
+ levels decline and restoration of NAD
+ has been reported to ameliorate cardiac dysfunction (Diguet et al,
2018; Matasic et al,
2018). NAD
+ deficiency, therefore, has been considered one mechanism for heart failure (Yamamoto et al,
2014; Yano et al,
2015). NAD
+ deficiency in failing heart tissues has been mainly attributed to decreased NAD
+ salvage synthesis secondary, for example, to decreased expression of nicotinamide phosphoribosyltransferase (NAMPT), a rate-limiting enzyme in the NAD
+ salvage pathway that catalyzes the conversion of nicotinamide (NAM) to nicotinamide mononucleotide (NMN) (Diguet et al,
2018; Matasic et al,
2018), which is subsequently converted to NAD
+ by nicotinamide mononucleotide adenylyl transferases (NMNATs) (Tatsumi et al,
2019). In addition, the activation of NADases, PARP and Sirtuin families, causes NAD
+ deficiency in MEFs where autophagic flux is impaired (Kataura et al,
2022). Here, we show that in heart failure induced by reduced autophagic flux, NAD
+ deficiency results from increased NAM clearance as a novel mechanism of decreased NAD
+ salvage synthesis. We did not observe the changes in the expression of kinases that are involved in NAD
+ salvage synthesis including NAMPT and NMNAT1-3 or NADases including Sirtuin families, PARP1/2, and BST1. Of note, protein expression of CD38, one of the NADases was decreased. Furthermore, we did not observe the changes in the activities of PARP1/2 and SIRT1. In contrast to heart failure from other causes, this NAD
+ deficiency is due to induction of NNMT by SQSTM1-NF-κB signaling. Thus, although NAD
+ deficiency plays a critical role in the development of heart failure in the mouse model of
ATG3 deficiency, which blocks autophagic flux, the mechanisms underlying NAD
+ deficiency are distinct from other causes of heart failure. These data support the centrality of perturbed NAD
+ metabolism in heart failure but reveal additional mechanisms by which this might occur.
Of note, the affinity of NNMT to NAM is ~400 µM, several orders of magnitude lower than that of NAMPT with K
M of the low nanomolar range (Burgos and Schramm,
2008; Pissios,
2017), implying that the recycling of NAM back into NAD synthesis by NAMPT may predominate over the formation of MeNAM via NNMT. It is therefore unexpected that the moderate upregulation of NNMT protein in the present study may outcompete NAMPT to methylate NAM toward MeNAM, resulting in NAD
+ deficiency. One study has indicated that the coexistence of NNMT with NAMPT largely accelerates NAD
+ consumption to result in NAD
+ deficiency (Bockwoldt et al,
2019). However, the mechanisms by which NNMT competes with NAMPT to divert NAM toward MeNAM and whether NNMT could suppress NAMPT remains to be elucidated. Another interpretation is the release of NAM inhibition on NAD consumption enzymes to accelerate NAD
+ consumption flux (Bockwoldt et al,
2019). However, in this study, we did not observe increases in protein expression or activities of NAD
+-consuming enzymes. NNMT induction resulting in NAD
+ deficiency has also been observed in other tissues such as liver (Griffiths et al,
2021; Komatsu et al,
2018). Given the importance of NAD
+ metabolism in cellular homeostasis, it will be important in future studies, to illustrate how NNMT and NAMPT coordinately regulate NAD
+ consumption flux by diverting NAM.
Studies have shown that NAD
+ deficiency due to the inhibition of NAMPT blocks autophagic flux in cardiomyocytes, in part by SIRT1-mediated acetylation of autophagic mediators (Hsu et al,
2009; Hsu et al,
2014). In this study, we found that reduced autophagic flux may induce NAD
+ deficiency by accelerating NAM clearance without changing NAMPT protein expression. Thus, two distinct mechanisms of NAD
+ deficiency may cause cardiac dysfunction, which can be reversed by restoring the NAD
+ pool. Therefore, it is likely that autophagy and NAD
+ metabolism regulate each other, and the consequence or direction of this regulation might be context-dependent.
Autophagic flux has been reported to maintain mitochondrial structure and function by degrading dysfunctional mitochondria via mitophagy (Lee et al,
2012). Impaired autophagic flux has been associated with accumulation of dysfunctional mitochondria that may be characterized by impaired O
2 consumption, decreased ATP production, increased ROS generation and cytochrome
c release that may precipitate apoptotic cell death (Nakai et al,
2007; Taneike et al,
2010; Wu et al,
2009). In this study, we observed decreased mitochondrial respiratory capacity that precedes cardiac dysfunction in ATG3-deficient hearts. Furthermore, we observed reduced mitochondrial content, which likely resulted in part from reduced mitochondrial biogenesis, given the reduced expression of transcriptional regulators of mitochondrial biogenesis such as PPARGC1A, NRF1, and NRF2. In addition, we observed evidence of inhibited mitophagy. Thus, impaired mitophagy and decreased mitochondrial biogenesis may simultaneously contribute to mitochondrial dysfunction when autophagic flux is impaired. Crosstalk between NAD
+ availability and PPARGC1A signaling, or the regulation of NAD
+ metabolic pathways by PPARGC1A have been described (Tran et al,
2016). What is clear, is that the well-documented relationships between NAD
+ metabolism and mitochondrial dysfunction (Kraus et al,
2014; Lee et al,
2019), or between autophagy and mitochondrial function is likely quite complex. A widely accepted mechanism is increased mitochondrial protein acetylation, attributed to NAD
+ deficiency and reduced sirtuin activity (Horton et al,
2016). Increased acetylation of mitochondrial proteins has been implicated not only as a mechanism for perturbing mitochondrial energy homeostasis, but also for promoting mitochondrial MPTP opening (Karamanlidis et al,
2013). However, the relationship between PPARGC1A acetylation and mitochondrial biogenesis (Nemoto et al,
2005; Rodgers et al,
2005) is complex and incompletely understood, and whether or not acetylation of PPARGC1A alters its cellular functions is unclear. Thus, the precise mechanisms linking impaired autophagic flux or decreased NAD
+ availability with PPARGC1A expression or activity as observed in this study remains to be elucidated and will require additional study.
Many studies have demonstrated complex crosstalk between autophagy, mitochondria, and lipid metabolism (Civitarese et al,
2010; Dong and Czaja,
2011; Gumucio et al,
2019; Lee et al,
2013; Rambold et al,
2015; Schulze et al,
2016; Singh et al,
2009; Zhang et al,
2018), with impairment of any of these pathways perturbing the others. Thus, the coexistence of impaired autophagy, mitochondrial dysfunction, and impaired lipid metabolism often occurs, and is often accompanied by lipid accumulation (Civitarese et al,
2010; Dong and Czaja,
2011; Gong et al,
2015; Gumucio et al,
2019; Lee et al,
2013; Rambold et al,
2015; Schulze et al,
2016; Singh et al,
2009; Zhang et al,
2018). Consistent with this view, we observed altered fat metabolism in autophagy defective hearts, namely decreased fat oxidation in concert with evidence of myocardial lipid accumulation. Thus, ATG3-deficient hearts could be more susceptible to lipotoxicity. Although impaired mitochondrial metabolism could account for this phenomenon, the possibility also exists that depression of the NAD
+ pool might alter fat metabolism in other ways and will be the subject of future studies.
In conclusion, we describe a novel mechanism by which impaired autophagic flux induces mitochondrial dysfunction in cells and hearts and heart failure in vivo. Autophagic flux maintains cardiac and mitochondrial function by mediating SQSTM1-NF-κB-NNMT signal transduction that controls the cellular levels of NAD
+. Thus, the present study identifies a previously unrecognized regulatory pathway linking NAD
+ metabolism and autophagic flux. This pathway may be physiologically relevant linking nutrient deprivation, autophagy induction, and the maintenance of the cellular NAD
+ pool. Conversely, impaired autophagy, by promoting depletion of the NAD
+ pool, may contribute to tissue dysfunction such as heart failure. Consistent with previous studies (Diguet et al,
2018), this study provides strong experimental evidence to further support that boosting NAD
+ metabolism represents a potential therapeutic strategy for ameliorating mitochondrial dysfunction and improving contractile function in heart failure (Diguet et al,
2018). Recent reports of small molecule NNMT inhibitors, which increase cellular NAD
+ levels leading to metabolic benefits such as the reversal of diet-induced obesity and prevention of muscle senescence (Neelakantan et al,
2019; Neelakantan et al,
2018; Sampson et al,
2021) supports the viability of modulating NNMT activity to achieve these goals.
Methods
Animal studies
The study protocol was approved by the Institutional Animal Care and Use Committees of the University of Utah, the Carver College of Medicine of the University of Iowa, and the David Geffen School of Medicine at the University of California, Los Angeles. Mice were maintained at the central animal facility with 12-h dark and 12-h light cycle, temperature at ~23 °C, and humidity at 40 to 60%. Mice had free access to water and food. All mice for experiments in this study were male on the C57BL6/J background. Female mice were used only for breeder cages and calculation of the percentage of cAtg3 mice (Appendix Table S
1). Mice were starved for 5–6 h before sacrifice. Mice were anesthetized by chloral hydrate or 2% isoflurane gas with an inflow rate of 1 mL/min, and then hearts were immediately removed and rinsed in ice-cold phosphate-buffered saline (PBS, 10010023, ThermoFisher Scientific, Waltham, MA) before being snap-frozen in liquid nitrogen.
The
ATG3loxP/loxP mice (Cai et al,
2018),
SQSTM1loxP/loxP mice (Okada et al,
2009), and mice expressing Cre recombinase under the control of the α-myosin heavy chain promoter (
MYH6-Cre) (Abel et al,
1999) were previously described. Constitutive cardiac-specific
ATG3 KO mice were generated by crossing
ATG3loxP/loxP mice with
MYH6-Cre mice. Age-matched
ATG3loxP/loxP mice without Cre were used as control mice. Constitutive cardiac-specific
SQSTM1 KO mice were generated by crossing
SQSTM1loxP/loxP mice with
MYH6-Cre mice. Age-matched
SQSTM1loxP/loxP mice without Cre were used as controls.
Mice expressing a tamoxifen-inducible Cre recombinase under the control of the inducible cardiac-specific MYH6 promoter (MYH6-MerCreMer; B6.FVB(129)-A1cfTg(MYH6-Cre/Esr1*)1Jmk/J) were obtained from The Jackson Laboratory (00565, Bar Harbor, ME). For inducible cardiac-specific ATG3 deletion, 6-week-old mice harboring MYH6-MerCreMer and ATG3loxP/loxP genes were intraperitoneally injected with tamoxifen at a dose of 100 µg/g body weight/day for 5 consecutive days. Age-matched ATG3loxP/loxP mice without MYH6-MerCreMer were injected with the same amount of tamoxifen as control mice.
Cardiomyocyte isolation
Adult mouse cardiomyocytes were isolated as described in our previous study (Harris et al,
2022; Zhang et al,
2020). Briefly, adult mouse hearts were cannulated through the aorta and then perfused on a Langendorff system for ~5 min with Buffer A (113 mM NaCl, 4.7 mM KCl, 0.6 mM KH
2PO
4, 0.6 mM Na
2HPO
4, 1.2 mM MgSO
4, 10 mM HEPES, 12 mM NaHCO
3, 10 mM KHCO
3, 30 mM taurine, 10 mM 2,3-butanedione monoxime [BDM; Sigma-Aldrich, B0753], 5.5 mM glucose, pH 7.0). The hearts were then perfused with a digestion buffer (Buffer A with the addition of Type II collagenase [Gibco, 17101015] at 300 U/mL and CaCl2 at 50 μM). The digestion times were determined by the hardness of the hearts and the rate of perfusion. Once the hearts were soft and the perfusion rate markedly increased, the ventricles were immediately dissected into small pieces in ice-cold Buffer A containing BSA (Research Products International,
A30075) at 100 mg/mL. The cardiomyocyte suspension was then centrifuged at 2000×
g for 3 min at 4 °C. The supernatant was discarded, and the cardiomyocyte pellet was then washed twice by resuspension in ice-cold Buffer A containing BSA at 100 mg/mL and centrifugation for 3 min at 4 °C. After washing, cardiomyocyte pellets were homogenized in ice-cold King homogenization buffer (pH 7.2) containing 50 mM HEPES, 10 mM sodium pyrophosphate, 10 mM sodium fluoride, 2 mM EDTA, 2 mM sodium orthovanadate, 1% Triton X-100, and 10% glycerol, supplemented with Halt™ Protease and Phosphatase Inhibitor Cocktail, EDTA-free (78447, ThermoFisher Scientific) at a ratio of 1:100. Then lysates were centrifuged at 20,000×
g for 30 min at 4 °C, and the supernatants collected for immunoblotting.
Transthoracic echocardiography
For assessment of LV fractional shortening, mice were anesthetized with 0.5–2% isoflurane gas with an inflow rate of 1 mL/min and placed on a heated stage (37 °C). Chest hair was then removed with a topical depilatory agent before echocardiography and pre-warmed ultrasonic gel was applied. B-mode images in parasternal long- and short-axis projections (at the papillary muscle level) were obtained using a Vivid 7 Pro ultrasound machine with a 13 MHz linear probe (GE Medical Systems, Boston, MA) by an experienced operator blinded to mouse identity. Fractional shortening (%) was calculated as [(LVDd − LVDs)/LVDd]*100, where LVDd is the LV dimension at diastole and LVDs is the LV dimension at systole (Riehle et al,
2011).
For assessment of LV ejection fraction (%), mice were sedated with Midazolam (0.3 mg/kg. Body weight) (NDC 0409-2305-17, Pfizer, New York, NY), and chest hair was then removed with a topical depilatory agent before the echocardiogram. Echocardiography was performed by an experienced operator blinded to mouse identity. M model images were obtained using a Vevo 2100 ultrasound machine with a 30 MHz linear array transducer (Visual Sonics, Bothell, Washington). Ejection fraction [%] was calculated as [(LVDd3 – LVDs3)/LVDd3] * 100. Echocardiography data were analyzed by an operator in a blinded fashion.
Histological analysis
Myocardial fragments were stained with H&E (047223.22, ThermoFisher Scientific; and HT110116, Sigma-Aldrich, St. Louis, MO) or Masson’s trichrome (Sigma-Aldrich, HT15-1KT) as previously described (Riehle et al,
2013). Stained slides were analyzed as previously described (Mandarim-de-Lacerda,
2003).
TUNEL staining
TUNEL staining was conducted in heart tissue using In Situ Cell Death Detection Kits (12156792910, Roche, Little Falls, NJ) following the supplier’s protocol.
Genomic DNA isolation and the measurement of DNA damage
Genomic DNA was isolated using a genomic DNA isolation kit (ab65358, Abcam, Waltham, MA), and the measurements of DNA damage were performed using a DNA damage-AP sites-Assay kit (ab211154, Abcam). The operations were completed according to the manufacturer’s manuals.
Oil red O staining
Heart samples were embedded in OCT on dry ice, and then sections were cut at ~8 μm thickness. After being fixed in 4% formalin in PBS, the sections were stained with Oil Red-O followed by lightly staining nuclei with hematoxylin (047223.22, ThermoFisher Scientific). The areas were randomly selected, and images were captured by bright-field microscopy.
Starvation
Mice were deprived of food for 48 h starting at 8:00 AM, and then mice were immediately euthanized, and tissues harvested. Random-fed mice were used as controls.
TGs measurement
TGs in heart tissues were measured using a Colorimetric Assay Kit (Cayman Triglyceride Colorimetric Assay Kit (10010303, Cayman, Ann Arbor, MI). The measurements were performed according to the manufacturer’s manual.
PARP activity assay
PARP activity was measured using a PARP/Apoptosis universal colorimetric assay kit (4677-096-K, R&D Systems, Minneapolis, MN), and the measurements were performed according to the manufacturer’s manual.
Assay of NAD+ levels
NAD+ levels in cells and heart tissues were measured using a NAD/NADH assay kit (ab6548, Abcam), and the measurements were performed according to the manufacturer’s manual.
Cell culture experiment
H9c2 cells (rat embryonic cardiomyoblasts; CRL-1446, ATCC, Manassas, VA) were maintained in high-glucose DMEM (11965-092, ThermoFisher Scientific) supplied with 10% FBS (97068-085, VWR, Radnor, PA). For siRNA knockdown of
Atg3,
SQSTM1,
NNMT, and
RELA (NF-κB p65), H9c2 cells were infected with siRNAs against
Atg3,
SQSTM1 (6399, Cell Signaling Technology, Danvers, MA; or paired sequences as indicated),
NNMT (L-101014-02-0005, GE Dharmacon), and RELA (6261S, Cell Signaling Technology), or scramble siRNA at 80–100 nM for 48–72 h together with Lipofectamine 2000 (11668500, Thermofisher Scientific) under nutrient-replete conditions. The sources of silencers or paired sequences are listed in Appendix Table S
4.
For NNMT overexpression, H9c2 cells were infected with an adenovirus encoding GFP or NNMT (kind gift from Dr. Pavlos Pissios, Beth Israel Deaconess Medical Center, Boston, MA; Appendix Table S
5) (Hong et al,
2015), and further cultured in high-glucose DMEM with 10% FBS for an additional 48 h before harvest. The cells and medium were collected, respectively. The NAD
+ levels and its related metabolites were measured using LC-MS methods as described in previous studies (Trammell et al,
2016a,
2016b).
For SQSTM1 and ATG7 overexpression, H9c2 cells were infected with 5 μg/mL plasmid encoding
HA-FLAG (10792, Addgene, Watertown, MA),
HA-SQSTM1 (28027, Addgene), or
ATG7 (24921, Addgene) for 60 h (Appendix Table S
5) together with Lipofectamine 2000 (11668500, Thermofisher Scientific). Then, chloroquine (CQ, C6628, Sigma-Aldrich) at 20 µM was added to H9c2 cells for ~24 h. under nutrient-replete conditions.
For Torin1 (10997, Cayman) treatment, Torin1 at 100 nM was added to H9c2 cells for ~16 h under nutrient-replete conditions.
An NF-κB reporter kit (60614, BPS bioscience, San Diego, CA) was used for the measurement of NF-κB luciferase activity in cells. The experiment was performed according to the manufacturer’s manual. Briefly, H9c2 cells were transfected with the NF-κB luciferase reporter vector or non-luciferase vector as control. After 24 h of transfection, H9c2 cells were treated with Veh (PBS) or CQ at 20 µM for an additional 24 h in the nutrient-replete medium. A dual-luciferase assay was performed based on the dual-luciferase reporter assay system (E1960, Promega, Madison, WI).
For cell starvation, H9c2 cells were washed with PBS twice, and cells were cultured in Hank’s Balanced Salt Solution (HBSS, 1.26 mM CaCl2, 0.5 mM MgCl2, 0.4 mM MgSO4, 0.4 mM KCl, 0.35 mM NaHCO3, 8 mM NaCl, 0.05 mM Na2HPO4) supplemented with 0.5 mM pyruvate (P8574, Sigma-Aldrich).
Autophagosome isolation
The protocol for autophagosome isolation from H9c2 cells was modified from protocols described in a previous report (Yao et al,
2019). In brief, H9c2 cells were first transfected with adenovirus encoding either GFP or GFP-tagged LC3, and then cells encoding GFP-LC3 were treated with either vehicle (Veh) or CQ at 20 µM for 16 h under nutrient-replete conditions. Cells were collected in the ice-cold separation buffer containing 250 mM sucrose (Sigma, S0389), 1 mM ethylenediaminetetraacetic acid (EDTA; E58100, Research Products International, Mt Prospect, IL), 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES; Research Products International, H750303), pH 7.4, with addition of Halt
TM protease & phosphatase inhibitor cocktail at 1:1000 (ThermoFisher Scientific, 78446). Then, cell lysates were homogenized using a Dounce glass homogenizer, followed by centrifugation at 1000×
g for 10 min at 4 °C to eliminate nuclei. The post-nuclear supernatant fractions were further centrifuged at 17,000×
g for 20 min at 4 °C. Then, the supernatant fractions were collected as cytosolic fractions, and the pellet fractions containing autophagosomes were washed twice in PBS.
After washing, the pellet fractions were resuspended in the separation buffer, and primary antibody against GFP (55494, Cell Signaling Technology) was added to the pellet fractions, followed by incubation overnight at 4 °C. Next, protein A agarose beads (Millipore-Sigma, 16-125) were added and incubated for 1 h at 4 °C. The complexes were washed in ice-cold PBS for up to three or four times. Then, immunoblotting and qPCR were conducted, respectively.
For immunoblotting, following the addition of loading buffer (ThermoFisher Scientific, LC2676), the complexes were boiled at 85 °C for 5 min. Two controls, antibody (AB) no lysate and cytosolic fraction no AB were performed. For AB no lysate, the primary antibody against GFP was directly added to the separation buffer followed by the IP protocol as described above. For no AB, cytosolic fractions were boiled in loading buffer at 85 °C for 5 min. Immunoblotting was conducted as described below.
For qPCR, after washing, the pellet fractions were resuspended in TRizol reagent (ThermoFisher Scientific, 15596018), and total RNA in the pellet fractions was extracted and purified with the RNeasy kit (Qiagen, 74104). Quantitative real-time PCR was performed using SYBR Green (ThermoFisher Scientific, 2012607) with RNU6-1 as a reference.
Mitochondrial isolation
Mitochondria were isolated from fresh WT and cAtg3-KO mouse hearts. Fresh whole hearts were cut into pieces using a scissor in ice-cold buffer (pH 7.2) containing 70 mmol/L sucrose, 210 mmol/L d-mannitol, 1 mmol/L EGTA, 0.5% fatty acid-free BSA, and 5 mmol/L HEPES, and then homogenized using a Wheaton 903475 overhead Stirrer at speed of 8–10 for ~0.5 to 1 min on ice. The homogenates were then centrifuged at 4000×g for 4 min at 4 °C. After centrifugation, the pellets were discarded, and the supernatants were further centrifuged at 10,000×g for 10 min at 4 °C. The pellets were saved as the mitochondrial fraction, and the supernatant was saved as the cytosolic fraction. The mitochondrial fractions were then lysed in ice-cold King homogenization buffer (pH 7.2) containing 50 mM HEPES, 10 mM sodium pyrophosphate, 10 mM sodium fluoride, 2 mM EDTA, 2 mM sodium orthovanadate, 1% Triton X-100, and 10% glycerol, supplemented with Halt™ Protease and Phosphatase Inhibitor Cocktail, EDTA-free (78447, ThermoFisher Scientific) at a ratio of 1:100. Then lysates were then centrifuged at 20,000×g for 30 min at 4 °C, and the supernatants were collected for immunoblotting.
Oxygen consumption analysis
Oxygen consumption was measured in H9c2 cells using a Seahorse XF Analyzer (Agilent Technology, Santa Clara, CA) as described before (Dott et al,
2014). H9c2 cells were seeded in Seahorse cell culture plates (Agilent Technology, Santa Clara, CA). Basal oxygen consumption rates (OCR) and extracellular acidification rates (ECAR) were measured to establish baseline rates. The cells were then treated with the following compounds in succession: 1 µg/mL oligomycin (Sigma, O4876), 0.5 µg/mL FCCP (Sigma, C2920), and 1 µg/mL rotenone (Sigma, R8875). Mitochondrial respirometry by Seahorse was conducted by an operator in a blinded fashion.
For the measurement of oxygen consumption in permeabilized cardiac fibers, left ventricular (LV) subendocardial muscle fibers were used to measure oxygen consumption rates and ATP synthesis as previously described (Boudina et al,
2005). Oxygen consumption rates were determined in the presence of palmitoyl-carnitine (Sigma, P1645) at 0.02 mM (V
O), 1 mM ADP (A2754, Sigma) (V
ADP), and 1 μg/mL of the ATP synthase inhibitor oligomycin (A7699, Sigma) (V
Oligo). ATP levels were measured in fibers after the addition of 1 mM ADP.
Chromatin immunoprecipitation (CHIP) assay
ChIP assays were performed as described previously with minor modification (Gao et al,
2015). Chromatin was extracted from mouse hearts using the SimpleChIP kit according to the manufacturer’s instructions (9004S, Cell Signaling Technology). Briefly, tissues were cross-linked with 1% formaldehyde (F10800, Research Products International) for 10 min and then lysed by micrococcal nuclease (EN0181, ThermoFisher Scientific) and sonicated using a sonic dismembrator five times for 15 s each time. RELA (NF-κB p65) protein was immunoprecipitated from precleared lysates with Protein A/salmon sperm DNA agarose beads (16-157, Millipore-Sigma,). DNA was released from protein-DNA complexes by proteinase K (115 879 001, Roche) digestion and then subjected to qPCR using power SYBR green kit. The RELA binding region of the
NNMT promoter was amplified using the following primers: forward, 5′-TAG CCA TGA GCT CGT TTC CT-3′, and reverse, 5′-CAG AAG CCC TGA GTC CAG AG-3′. ChIP-qPCR data were normalized to total chromatin.
Metabolomics analysis
Mice were euthanized, and hearts were immediately removed, rinsed in ice-cold PBS, and frozen in liquid nitrogen. Glycolytic and citric acid cycle metabolites in hearts were detected using GC-MS as previously described (McClain et al,
2013). This assay was conducted by an operator in a blinded fashion.
In vivo NAD+ flux metabolomics
Mice were intraperitoneally injected with heavy-isotope-labeled nicotinamide riboside (NR) (
13C, D double-labeled NR,
13C, D-NR) as previously described (Trammell et al,
2016a,
2016b). 0, 30, and 60 min after intraperitoneal injection of
13C, D-NR, mice were euthanized, and hearts were harvested. For metabolomics analyses, hearts were immediately removed, freeze-clamped and rapidly submerged in liquid nitrogen. NAD
+ levels and its related metabolites were measured using LC-MS methods as described in previous studies (Trammell et al,
2016a,
2016b).
NMN administration
β-nicotinamide mononucleotide (NMN; N3501, Sigma) was prepared in PBS and injected intraperitoneally to 3-week-old mice at 500 mg/kg/d for 7 days. PBS (pH 7.4) was injected intraperitoneally as controls. Mice were euthanized 6 h. after the last injection, and hearts were immediately removed, rinsed in ice-cold PBS, and then frozen in liquid nitrogen. ATP, ADP, AMP, NAD
+, and NADH were measured using LC-MS methods as previously reported (Shibayama et al,
2015).
Administration of 5-amino-1-methylquinolinium iodide
5-amino-1-methylquinolinium iodide (NNMTi; 42464-96-0, Tocris, Minneapolis, MN) was first dissolved in DMSO and then resuspended in vehicle (0.2% CMC, 0.25% Tween 80, and 2% DMSO). The solution was sterilized through a 0.45-µm filter. NNMTi was injected intraperitoneally to mice at 5 mg/kg twice/day for 3 days. The vehicle (0.2% CMC, 0.25% Tween 80, and 2% DMSO) was injected intraperitoneally as control. Twelve hours after the last injection of NNMTi, mice were subjected to echocardiography under isoflurane anesthesia as described in “Transthoracic echocardiography”.
Measurement of the activities of citrate synthesis (CS), carnitine palmitoyltransferase (CPT), glyceraldehyde 3-phosphate dehydrogenase (GAPDH), and Sirt1
For measurements of CS activity, hearts were homogenized in ice-cold homogenization buffer containing 20 mM HEPES (Research Products International) and 10 mM EDTA (Research Products International) at pH 7.4. Homogenates were subjected to two freeze/thaw cycles to liberate CS from the mitochondrial matrix and then diluted to an approximate final protein concentration of 1 μg/μL. The reaction was performed in 1 mL of reaction buffer containing 20 mM HEPES (H7523, Sigma), 1 mM EDTA, 220 mM sucrose (S0389, Sigma), 40 mM KCl (Sigma, P3911), 0.1 mM 5,5′-dithio-bis (2-nitrobenzoic acid) (DTNB, Sigma, D8130), and 0.1 mM acetyl-CoA (pH 7.4 at 25 °C) (A1625, Sigma). The reaction was started by the addition of 0.05 mM oxaloacetate (O4126, Sigma) and monitored at 412 nm for 3 min using an Epoch Microplate Spectrophotometer (BioTek Instruments, Inc. Winooski, VT). The result was normalized to protein content.
For CPT activity measurement, mitochondria were isolated from fresh heart tissue as previously described (Riehle et al,
2011). Mitochondria were assayed in 1 mL of reaction buffer containing 20 mM HEPES (H7523, Sigma), 1 mM EGTA (E3889, Sigma), 220 mM sucrose (S0389, Sigma), 40 mM KCl (P3911, Sigma), 0.1 mM DTNB (D8130, Sigma), 1.3 mg/ml BSA, and 40 μM palmitoyl-CoA (P9716, Sigma). The reaction was started by the addition of 1 mM carnitine (C0283, Sigma) and monitored at 412 nm for an additional 4 min using an Epoch Microplate Spectrophotometer (BioTek Instruments). The CPT activity was normalized to mitochondrial protein levels.
For measurements of GAPDH and SIRT1 activities, heart tissues were homogenized and GAPDH and SIRT1 activities were measured according to the manufacturer’s manuals (ab204732 and ab156065, Abcam). Exogenous NAD+ (10127965001, Sigma-Aldrich) was added to the reaction buffer to a final concentration of 10 mM.
Administration of colchicine (Col) and chloroquine (CQ) to mice
For Col (C9754, Sigma) treatment, Col was dissolved in PBS and injected intraperitoneally in mice at 18, 6, and 2 h before heart harvest at doses of 4, 1, and 1 µg/g body weight, respectively. For CQ treatment, CQ was dissolved in PBS and injected intraperitoneally in mice at 48, 24, and 2 h. before heart harvest at doses of 30, 30, and 50 µg/g body weight, respectively. After the treatments above, mice were euthanized, hearts were immediately removed and rinsed in ice-cold PBS, and then frozen in liquid nitrogen.
Quantitative real-time PCR
Total RNA was extracted from hearts or H9c2 cells using TRIzol reagent and purified with the RNeasy kit. Quantitative real-time PCR was performed using SYBR Green with ROX as an internal reference dye. The expression level was normalized to the transcript levels of the
RPL13A,
GAPDH,
RNU6-1, or
ACTB. The primer sequences are listed in Appendix Table S
2.
Immunoblotting
For immunoblotting, proteins were resolved on SDS-PAGE and analyzed using the LI-COR Odyssey Imager (LI-COR). The following primary antibodies were used (Appendix Table S
3): ATG3 (A3231), MAP1LC3A (LC3, L8918), TUBA4A (alpha-Tubulin, T5168) from Sigma-Aldrich; ACTB (beta-Actin, 3700), SQSTM1 (p62, 5114), ATG7 (8858), Acetylated-Lysine (9441), FOXO1 (2880), GAPDH (2118), RELA S536 (NF-κB p65 S536, 3036), RELA (NF-κB p65, 6956), MTOR (2971), MTOR S2448 (4517), AMPKα T172 (2531), AMPKα (2793), P70S6K T389 (9206), P70S6K (2708), ULK1 (8054), Sirtuin antibody sampler kit (9787), SIRT4 (69786), PARP1 (9532), GFP (55494), Poly/Mono-ADP Ribose (83732), and PRKN (Parkin, 4211) from Cell Signaling Technology; Ac-FKHR (D-19, sc-49437), PPARGC1A (PGC1α, H-300, sc-13067), GAPDH (SC-32233), SIRT1 (B-10, sc-74504), and TOMM20 (TOM20, sc-17764) antibodies from Santa Cruz Biotechnology (Santa Cruz Biotechnology, Dallas, TX); CD38 (60006-1-lg), BST1 (CD157, 16337-1-AP), NMNAT1 (11399-1-AP), NMNAT3 (13261-1-AP), PARP2 (2055-1-AP), and NNMT antibody (15123-1-AP) and PINK1 antibody (23274-1-AP) from Proteintech (Rosemont, IL); NMNAT2 (PA5-115662) from Invitrogen (ThermoFisher Scientific); SDHA (ab14715) antibody from Abcam.
Statistical analysis
Statistical analyses were performed using GraphPad Prism 8. All data are presented as mean ± SEM. Statistical significance (P < 0.05) was determined by one-way ANOVA followed by Bonferroni’s multiple comparison test. An unpaired t test was used to compare between two groups.