Cardiac Autophagy Deficiency Attenuates ANP Production and Disrupts Myocardial-Adipose Cross Talk, Leading to Increased Fat Accumulation and Metabolic Dysfunction Free

Cross talk between the heart and peripheral tissues plays an important role in maintaining physiological homeostasis and is particularly evident when this axis is disrupted in individuals with heart failure (1,2). Increased autophagy is one mechanism known to protect the heart from stress-induced remodeling and dysfunction (3), and lack of autophagy can contribute to or enhance cardiac dysfunction (4).

Autophagy is an endogenous self-protection mechanism in cells, relying on lysosomes to degrade cytoplasmic aging or damaged proteins and organelles and thus prevent further damage to cells (4). The process of autophagy flux can be subdivided into multiple steps including the induction of autophagy, cargo recognition, autophagosome formation, autophagosome-lysosome fusion, and finally cargo degradation. Autophagy-related (Atg) proteins are involved in different steps of this pathway, with Atg7 playing an important role in the initiation of autophagosome formation. Atg7, an E1-like enzyme, activates ubiquitin-like proteins Atg8 and Atg12, which undergo conjugation to form complexes that regulate the formation of autophagosome membranes (5). Mice with Atg7 knockout are normal at birth but cannot survive beyond 1 day (6). Thus, in vivo studies with Atg7 have relied on tissue-specific or conditional knockout models (7,8). Current knowledge indicates that activation of autophagy is an important cardioprotective mechanism that limits the accumulation of misfolded proteins, improves mitochondrial dysfunction, and reduces oxidative stress, thereby limiting myocardial damage caused by stress such as ischemia-reperfusion injury (9). Impaired and deficient autophagy is associated with aging, obesity, and diabetes and is now well established as one contributory mechanism for development of cardiac dysfunction (4).

Cross talk between peripheral tissues is mediated by multiple factors including peptide hormones, metabolites, and nucleic acids (1). To date, little is known regarding the role of cardiac autophagy in determining cross talk between the heart and other organs. Cardiac natriuretic peptides (NP), atrial natriuretic peptide (ANP), and brain natriuretic peptide (BNP) are endocrine factors with well-established roles in blood pressure regulation (10) as well as proangiogenetic, antiatherosclerotic, and anti-inflammatory effects in the cardiovascular system (11). Importantly, cardiac NPs have also been shown to be important mediators of the cross talk between heart and adipose tissue (2). Indeed, cardiac NPs can stimulate adipocyte lipolysis and promote the “browning” of white adipose tissue (WAT) through increased respiration and thermogenesis leading to improved energy expenditure (12). In vivo experiments using whole-body and adipose tissue–specific deletion of NP clearance receptors demonstrated that enhanced ANP action can protect mice from diet-induced obesity and insulin resistance (12). ANP has also been suggested to work in concert with catecholamines in regulating lipid metabolism in adipose tissue (13,14).

In this study, we examined the role of autophagy in the cross talk between heart and adipose tissue through the generation of mice with cardiomyocyte-specific deletion of Atg7 (AKO). We hypothesized that disrupted heart-adipose cross talk due to defective cardiac autophagy and reduced ANP contributed to the obese and insulin-resistant phenotypes of AKO mice. Inducible autophagy deficiency in these mice allowed us to examine the resultant effects on peripheral metabolism and specifically alterations in adipose tissue. We observed that mice lacking cardiac autophagy accumulated fat mass, leading to increased body weight and insulin resistance.

Tamoxifen, type II collagenase, isoproterenol, fatty acid–free BSA, palmitic acid, and free glycerol determination kit were obtained from Sigma-Aldrich (St. Louis, MO). [1-¹⁴C]palmitic acid was purchased from PerkinElmer (Boston, MA), and D-[U-¹⁴C]glucose was purchased from American Radiolabeled Chemicals (St. Louis, MO). Antibodies used include β-actin (cat. no. 8457; Cell Signaling Technology), GAPDH (cat. no. 2118S; Cell Signaling Technology), adipose triglyceride lipase (ATGL) (cat. no. 2138; Cell Signaling Technology), hormone-sensitive lipase (HSL) (cat. no. 4107; Cell Signaling Technology), and phosphorylated HSL_Ser660 (cat. no. 4126; Cell Signaling Technology), UCP1 antibody (cat. no. ab23841; Abcam), ANP antibody (cat. no. STJ72680; St. John’s Laboratory), and Atg7 antibody (cat. no. sc-376212; Santa Cruz Biotechnology). ANP determination kit was from Thermo Fisher Scientific (cat. no. EIAANP), triglyceride analysis kit from BioVision (cat. no. K622), and cGMP kit from Thermo Fisher Scientific (cat. no. EMSCGMPL), and all were used following the manufacturer instructions.

All animal work was approved by the Animal Care Committee at York University. Animals were housed under a 12-h light/12-h dark schedule and provided with food and water ad libitum. Conditional cardiac depletion of Atg7 in cardiomyocytes was achieved by crossing αMerCreMer mice with Atg7^flox/flox mice; mouse strains are on the C57BL/6 background. All mice bred via this protocol were genotyped and assigned to experimental groups. Primers used for the genotyping of Atg7^flox/flox and αMerCreMer are listed below. DNA was extracted from ear tissue for genotyping.

Conditional Atg7 deletion was achieved by single tamoxifen (40 mg/kg) injection at the age of 7 weeks in male mice. Atg7^flox/flox Cre⁺ mice without tamoxifen injection were used as control animals. Animal body weights were monitored before tamoxifen injection (7 weeks) and weekly thereafter. One week before the induction of Atg7 deletion by tamoxifen injection, ALZET Osmotic Pumps (Model 2004) were subcutaneously implanted on the back of mice. Prior to implantation, pumps were put in sterile PBS at 37°C for at least 40 h. ANP solution or saline was injected into the pumps for subcutaneous implantation in mice under ketamine/xylaxine anesthesia and delivered at 0.05 μg/kg/min over 4 weeks.

At the end of treatment, animals were placed in the Comprehensive Lab Animal Monitoring System (CLAMS) from Columbus Instruments for 48 h for the measurement of in vivo metabolic parameters as previously described (15). Mice were allowed to acclimatize for 24 h prior to collection of data.

Glucose tolerance tests (GTT) and insulin tolerance tests (ITT) were performed in mice at the age of 10 weeks (3 weeks after corn oil or tamoxifen injection). Mice were fasted for 16 h (GTT) or 6 h (ITT) and then intraperitoneally injected with glucose (2 g/kg) or insulin (0.5 units/kg). Blood glucose levels were determined by Accu-Chek Aviva glucose meter at time points listed in Fig. 2.

All nuclear magnetic resonance (NMR) measurements were performed at 25°C on an AV III 700 NMR spectrometer operating with TopSpin software, version 3.2 (Bruker BioSpin, Karlsruhe, Germany) with a 5-mm TXI ¹H/¹³C/¹⁵N-Z cryo-probe. The data were acquired with use of the method developed and provided by the Metabolomics Innovation Centre (Edmonton, Canada). Briefly, each sample contained NMR buffer, 10% D₂O for field-frequency lock, and 2,2-dimethyl-2-silapentane-5-sulfonate for chemical shift reference. The spectra were acquired with the standard Bruker noesypr1d pulse program with 64,000 data points, a 1 s presaturation period, 100 μs mixing time, and 11.7 ppm spectral width. The spectra were processed with 0.3 Hz line broadening and baseline corrected. The spectra were automatically analyzed with use of the portal at www.magmet.ca.

Adipocyte isolation was performed as previously described (16). Briefly, epididymal (Epid) and subcutaneous inguinal (Sc Ing) fat depots were extracted and minced in Krebs-Ringer bicarbonate HEPES buffer (KRBH) prepared fresh on the day of each experiment from stock solutions of salts and buffers (stored at 4°C) to give the following final concentrations: 120 mmol/L NaCl, 4.8 mmol/L KCl, 2.5 mmol/L CaCl₂, 1.2 mmol/L KH₂PO₄, 1.2 mmol/L MgSO₄, 15 mmol/L NaHCO₃, 30 mmol/L HEPES, and type II collagenase (0.5 mg/mL). Before use, KRBH was gassed for 45 min with carbogen (95% O₂, 5% CO₂) and then BSA (3.5%) and glucose (5.5 mmol/L) were added. pH of the buffer was adjusted to 7.4 with NaOH. Minced tissues were incubated at 37°C with gentle agitation (120 orbital strokes/min) for ∼25–30 min. The digested tissue was then strained using a nylon mesh, and cells were transferred to 50-mL tubes, washed three times, and resuspended in KRBH containing 3.5% fatty acid–free BSA (KRBH-3.5% BSA). For distribution of an equal number of adipocytes in each treatment condition, cell diameters were measured and total cell numbers were determined as previously described (16).

Total RNA was obtained from atria or subcutaneous fat using the RNeasy Mini Kit (Qiagen Sciences, Germantown, MD) according to the manufacturer’s instructions. Total RNA was reverse transcribed using 0.5 μg poly-dT and 0.25 units reverse transcriptase at 42°C for 75 min and 95°C for 5 min. Quantitative real-time PCR analyses were performed as previously described (17). Briefly, cDNA and primers shown above were added to 20 μL reaction volume of the GoScript Reverse Transcription System (Promega, Madison, WI). PCRs were then performed with the CFX96 or CFX384 Touch Real-Time PCR Detection System (Bio-Rad Laboratories). Conditions were set to the following parameters: 3 min at 95°C followed by 40 cycles each of 15 s at 95°C and 40 s at 65°C. Commercial PCR array was used for the detection of adipose tissue fatty acid metabolic status (fatty acid metabolism [SAB target list] M384, cat no. 10034653; Bio-Rad Laboratories), in which 88 pairs of fatty acid metabolic-related gene primers were precoated onto a 384-well panel for detection with SYBR Green.

We normalized data by applying log transformation and scaling before statistical analysis, and a normality test was done with the normalized data. t test was performed in SPSS Statistics, version 25 (IBM, Armonk, NY) for identification of metabolites that differ between the wild-type (WT) and AKO groups. P value with cutoff of 0.05 was used for significance. False discovery rate (FDR) (adjusted P value) was acquired by correction of P values with the Benjamini-Hochberg method for multiple test adjustment. A volcano plot was plotted using log2 fold change and –log10 P value.

Fold change was calculated by comparison of the mean value in the AKO group to that in the WT group. For pathway analysis of quantitative PCR data, we subjected genes with their expression in WT and AKO groups to gene set enrichment analysis (GSEA) using Kyoto Encyclopedia of Genes and Genomes (KEGG) database in WebGestalt (WEB-based Gene SeT AnaLysis Toolkit) (18) under a Mus musculus background.

Western blot analysis was performed according to standard methods (19). Briefly, isolated tissues were homogenized in 25 mmol/L Tris-HCl and 25 mmol/L NaCl (pH 7.4), 1 mmol/L MgCl₂, 2.7 mmol/L KCl, 1% Triton-X, and protease and phosphatase inhibitors (0.5 mmol/L Na₃VO₄, 1 mmol/L NaF, 1 μmol/L leupeptin, 1 μmol/L pepstatin, and 20 mmol/L phenylmethylsulfonyl fluoride) buffer. Homogenates were centrifuged, and supernatant protein was quantified by Bradford protein assay. Sample (30 μg) was diluted 1:1 (v/v) with 2× Laemmli sample buffer, heated to 95°C for 5 min, resolved by SDS-PAGE, and transferred to polyvinylidene difluoride membranes. Western blotting was performed using primary antibodies at 1:1,000 dilution and secondary antibodies, and immune complexes were detected with enhanced chemiluminescence reagents (Bio-Rad Laboratories).

Glucose and palmitate oxidation was assessed by the production of ¹⁴CO₂ in adipocytes (5 × 10⁵ cells) isolated from the Epid and Sc Ing fat depots as previously described (16). Briefly, cells were incubated in KRBH-3.5% BSA containing either 0.2 μCi/mL [1-¹⁴C]palmitic acid and 200 μmol/L nonlabeled palmitate or 0.2 μCi/mL D-[U-¹⁴C]glucose and 5.5 mmol/L nonlabeled d-glucose for 1 h. Subsequently, the media were acidified with 0.2 mL H₂SO₄ (5 N), and the vials were maintained sealed at 37°C for an additional 1 h for the collection of ¹⁴CO₂ released from the cells and the media. The vials used for incubation had a centered isolated well containing a loosely folded piece of filter paper that was moistened with 0.2 mL 2-phenylethylamine/methanol (1:1, vol:vol) for the capture of ¹⁴CO₂. At the end of the incubation, the filter papers were removed and transferred to scintillation vials for radioactivity counting (16).

Lipolysis was measured in isolated Epid and Sc Ing adipocytes (5 × 10⁵ cells). For stimulation of lipolysis, adipocytes were incubated with isoproterenol (nonspecific β-agonist). Each assay was conducted in duplicates, and the samples were incubated for 60 min at 37°C with gentle agitation (80 orbital strokes/min). After incubation, a 200-μL aliquot of medium was taken from each vial for the determination of glycerol concentration.

Glucose incorporation into lipids in adipocytes was measured in isolated adipocytes from the Epid and Sc Ing fat depots as previously described (16). Briefly, 5 × 10⁵ cells were incubated in KRBH-3.5% BSA (containing 5 mmol/L glucose) with 0.2 μCi/mL D-[U-¹⁴C]glucose under basal or insulin-stimulated (100 nmol/L) conditions for 1 h at 37°C. Lipids were then extracted according to the method of Dole and Meinertz and assessed for radioactivity (16).

Data are expressed as mean ± SEM, with P < 0.05 considered statistically significant. Comparison between two groups was carried out by two-tailed nonparametric Mann-Whitney test or unpaired Student t test based on their normal distribution (Shapiro-Wilk) test results. One-way or two-way ANOVA was used to determine statistical significance among groups followed by Tukey test, where appropriate, with SPSS Statistics, version 25 (IBM).

The data sets generated during and/or analyzed during the current study are available from the corresponding author upon reasonable request.

Atg7^flox/flox Cre⁺ male mice were identified by genotyping of all mice bred under this protocol, and a representative example is shown in Fig. 1A and B. At the age of 7 weeks, tamoxifen (40 mg/kg) was injected intraperitoneally into Atg7^flox/flox Cre⁺ mice to induce cardiomyocyte-specific deletion of Atg7 (AKO mice); Atg7^flox/flox Cre⁺ mice without tamoxifen injection (WT) served as the control group. Atg7 protein level was determined 3 weeks later by Western blotting in ventricular tissue homogenates, and effective deletion of Atg7 protein was verified in AKO mice (n = 6) (Fig. 1C). We measured circulating triglyceride levels in these Atg7^flox/flox Cre⁺ mice with or without tamoxifen injection and found no significant alteration (data not shown). Prior to tamoxifen-induced Atg7 deletion, body weight did not differ between WT and Atg7^flox/flox Cre⁺ mice (n = 6 per group) (Fig. 1D). However, 3 weeks after Atg7 deletion was induced, the body weight was 1.3-fold higher in AKO than WT mice (n = 6) (Fig. 1E), while food intake did not differ between genotypes (n = 6) (Fig. 1F). Adiposity visibly increased in AKO mice, and there was a significant increase in subcutaneous inguinal (Sc Ing) and apparent increase in epididymal (Epid) fat masses of AKO mice, by 2.3- and 1.43-fold, respectively, in comparison with WT mice (n = 6) (Fig. 1G). No significant differences were found for interscapular brown adipose tissue mass (data not shown). Adipocyte thermogenesis was assessed through UCP1 expression, which was not detected in Sc Ing or Epid fat in WT or AKO mice (Fig. 1H).

The glucose excursion curves during GTT and ITT and quantitative area under curve analysis revealed that AKO mice developed insulin resistance (n = 3–4 per group) (Fig. 2A and B). Rates of oxygen consumption (n = 3–4 per group) (Fig. 2C and D) for AKO mice were 22% and 17% lower than for WT mice during the light and dark cycles, respectively. Ambulatory activity during the light cycle did not differ between AKO and WT mice; however, during the dark cycle AKO mice were significantly less active (47%) than WT mice (n = 3–4) (Fig. 2E and F). No significant differences were detected for respiratory exchange ratio (RER) between AKO and WT mice, although slightly higher values were recorded for AKO during the dark cycle (n = 3–4) (Fig. 2G and H).

Given the alterations observed in adiposity, GTT, and ITT, we used NMR to assess a group of 58 circulating metabolites in WT and AKO mice (n = 3 per group). The volcano plot (Fig. 3A) and heat map (Fig. 3B) indicate significant decreases in lactate (49%, P = 0.02, FDR = 0.67), 3-hydroxyisovaleric acid (79%, P = 0.04, FDR = 0.67), and choline (49%, P = 0.04, FDR = 0.67) in AKO mice compared with WT mice. Due to lactate’s direct association with metabolic analysis that we performed here, we then focused on lactate and used the ¹H spectra to also quantify relative plasma lactate levels. We found that circulating level of lactate was reduced by ∼50% in AKO mice (Fig. 3C).

Western blotting analysis revealed that ANP content was reduced in both atrial and ventricular tissues of AKO mice (Fig. 4A [immunoblots representative of n = 5 mice for atrial ANP and n = 3 for ventricle ANP]), which was accompanied by decreased circulating ANP levels in these animals (n = 4–6 per group) (Fig. 4B). To examine the significance of this change, we restored circulating ANP to normal levels in AKO mice via osmotic pump delivery of recombinant ANP. Indeed, after 3 weeks of pump implantation, circulating ANP (Fig. 4B) and body weight (n = 5–6) (Fig. 4C) of AKO mice were comparable with those of WT animals. As a biomarker of ANP action in adipose tissue, cGMP level was measured in Sc Ing fat, and we found that it was decreased in AKO mice and then restored to WT levels in AKO mice after ANP infusion (n = 4–6) (Fig. 4D).

Because adiposity was increased in AKO mice and this variable is partially determined by glucose and fatty acid metabolism within adipocytes, we assessed rates of glucose incorporation into lipids and glucose and fatty acid oxidation in adipocytes isolated from Sc Ing and Epid fat depots. Glucose incorporation into lipids did not differ between Sc Ing (Fig. 5A) and Epid (Fig. 5B) adipocytes under basal or insulin-stimulated conditions in AKO mice. Furthermore, ANP repletion did not alter this variable, indicating that lipogenesis was not affected by alterations in circulating ANP levels (n = 4–6) (Fig. 5A and B). Glucose oxidation was significantly increased (∼2.5-fold) in both Sc Ing (n = 3–5) (Fig. 5C) and Epid (n = 4–6) (Fig. 5D) adipocytes from AKO mice. However, replenishment of circulating ANP reduced glucose oxidation rates to values similar to those of WT mice in adipocytes from both fat depots (Fig. 5C and D). Fatty acid oxidation did not differ either among any of the conditions studied (n = 3–7) (Fig. 5E and F).

Since lipogenesis was not altered in AKO mice, we hypothesized that triglyceride breakdown could be suppressed in Sc Ing and Epid fat adipocytes of AKO mice. In fact, assessment of lipolysis in these cells revealed that the rate of isoproterenol-induced glycerol release was significantly reduced in Sc Ing (Fig. 6A) and Epid (Fig. 6B) adipocytes from AKO mice, with replenishment of circulating ANP resulting in a trend toward reversal of the suppression of lipolysis in the Sc Ing but not in Epid fat depot (n = 4–6) (Fig. 6A and B). We found that, in line with suppression of lipolysis, ATGL content was significantly reduced in the Epid fat depot of AKO mice (n = 4–6) (Fig. 6D), although its content in the Sc Ing fat depot was not significantly affected in these animals, even after ANP replenishment (n = 4–6) (Fig. 6C). Although not statistically significant, HSL_Ser660 phosphorylation also seemed to have been markedly reduced in both fat depots of AKO mice and reinfusion of ANP did not rescue this effect (n = 3–6) (Fig. 6E and F).

We used gene arrays to examine alterations in multiple genes related to lipid and carbohydrate metabolism in adipose tissue (n = 4) (Supplementary Fig. 1A). Gene expression fold change in AKO mice compared with WT mice is shown in a heat map (Supplementary Fig. 1A). However, there are no significant changes found in individual genes. For further investigation of the altered pathways, all genes with their expression in WT and AKO groups were subjected to GSEA using KEGG pathway database. Our analysis showed significant enrichment of a PPAR signaling pathway (P < 0.05) in AKO groups versus WT groups (n = 4) (Supplementary Fig. 1B).

Multiple peripheral tissues have now been characterized as endocrine organs, most notably adipose tissue, as well as more nontraditional examples like muscle and heart. So-called adipokines, myokines, and cardiokines secreted from these tissues act as important mediators of cross talk with other tissues throughout the body (1,20,21). Among these, decreased release of beneficial adipokines such as adiponectin or increased release of proinflammatory factors (such as lipocalin-2 and tumor necrosis factor-α) may be important in the occurrence and development of metabolic disorders (19,22,23). Importantly, cardiokines can regulate peripheral tissues and the development of metabolic disorders (1,24). Supporting this theory are recent studies that have shown that mitsugumin 53, a muscle/heart-produced factor, is a striatum-specific E3 ligase that can accelerate the degradation of ubiquitin-dependent insulin receptor and insulin receptor substrate 1, induce insulin resistance, and promote the development of metabolic syndrome and type 2 diabetes (25). An increasing number of studies have shown that NPs secreted by the heart can communicate with various organs, including adipose tissue, and participate in the regulation of systemic energy and substrate metabolism (12,26–29). However, it remains unclear how NP secretion is regulated and how this influences the development of obesity-related metabolic disorders.

Our study provides novel evidence uncovering the role of reduced ANP-mediated cross talk between autophagy-deficient hearts and adipose tissue. We observed that cardiac autophagy deficiency elicited impaired whole-body glycemic control and increased fat storage through alterations in ANP expression and secretion. This conclusion is supported by our findings that cardiomyocyte-specific deletion of Atg7 caused insulin resistance, as significantly higher glycemic excursions were observed in AKO compared with WT mice during GTT. We believe that our present study is the first to report the direct effects of autophagy deficiency on a reduction in ANP production by the heart. Finckenberg et al. (30) previously observed small changes in ANP expression in following caloric restriction in double transgenic rats harboring human renin and angiotensinogen genes. In this model, the authors observed protection against angiotension II–induced mitochondrial remodeling and cardiac hypertrophy and a decrease in cardiac ANP mRNA expression levels in caloric-restricted rats. This correlation is in keeping with our data, although it did not directly establish the link between myocardial autophagy and ANP. In addition, increased autophagy in this study was inferred by increased cardiac LC3-II levels, but it is well-known that this is not in itself a definitive measure of increased autophagy (31). The production of NPs can be regulated at multiple levels including transcriptional, translational, posttranslational, and clearance. We did not find significant changes in ANP mRNA levels between control and AKO animals (data not shown), and thus a cardiac autophagy defect most likely reduced NP levels through translational or posttranslational mechanisms. It is also possible that autophagy deficiency in heart may lead to altered secretion of additional cardiokines that can have effects on metabolic disease (32), and alteration in the myocardial secretome is certainly an area of research that warrants further investigation.

NPs are known to have pleiotropic biological effects (33), and it is possible that ANP-mediated cross talk with other organ systems could contribute to the effects we observed on glycemic control and adiposity. Here, we focused on direct effects on adipose tissue. Since ANP was known to act via its receptors on adipose tissue to modulate lipolysis, energy expenditure, adipokine release, and food intake (12,27–29), we tested whether Atg7 deletion could affect cardiac ANP content and release. We found that both myocardial and circulating levels of ANP were markedly reduced in AKO mice. Furthermore, levels of cGMP, a marker of ANP action in Sc Ing fat, and isoproterenol-stimulated lipolysis were reduced in Sc Ing and Epid adipocytes. Most importantly, administration of exogenous recombinant ANP through the implantation of an osmotic pump elevated cGMP levels in Sc Ing fat of AKO mice to values that were higher than those found in WT mice. This was accompanied by reversal of the suppressed isoproterenol-stimulated lipolysis found in Sc Ing adipocytes from AKO mice, although in Epid adipocytes this was not achieved. The molecular mechanisms behind these effects seem to, at least partially, involve reductions in content and activity of the lipolytic enzymes ATGL and HSL. These enzymes were distinctly affected in Sc Ing and Epid fat depots of AKO mice, which could explain the differences in lipolytic responses that we found in adipocytes isolated from these tissues. In fact, ATGL content was not significantly altered in Sc Ing fat, whereas ATGL in Epid fat was markedly reduced in AKO mice. Although not statistically different from WT mice, HSL_Ser660 phosphorylation seemed to be markedly downregulated in both fat depots of AKO mice, and exogenous ANP administration did not restore phosphorylation levels of this enzyme. We also tested whether increased adiposity could be due to enhanced lipogenesis in Sc Ing and Epid adipocytes. However, this did not seem to be the case, since glucose incorporation into lipids under either basal or insulin-stimulated conditions did not differ between WT and AKO mice, and exogenous administration of ANP had no effect on this variable. Therefore, it seems that it is the impaired breakdown of triglycerides in the WAT and not enhanced lipogenesis that leads to adipose tissue expansion in AKO mice.

Cardiac NPs have been reported to induce a brown fat thermogenic program in mouse and human adipocytes and affect substrate metabolism in WAT (14). In this context, we expected that with reduced circulating ANP in AKO mice, oxidation of glucose and fatty acids would be downregulated as a result of attenuated thermogenic activity in WAT. We measured UCP1 content in Sc Ing and Epid fat depots and could not detect UCP1 protein in either WT or AKO. Thus, alterations in adipocyte thermogenesis did not seem to have played a role in regulating adiposity in AKO. This was consistent with our findings that palmitate oxidation in isolated adipocytes did not differ under any of the conditions studied. However, it was surprising to find that rates of glucose oxidation were consistently elevated in adipocytes from both fat depots of AKO mice, a response that was fully prevented by the exogenous administration of ANP to AKO mice. Potentially, in the absence of the prolipolytic effects of ANP, adipocytes diverted glucose to oxidation as opposed to using it for lactate and glycerol production. Lactate serves as an important substrate for glyceroneogenesis, particularly under conditions of accelerated lipolysis, to sustain a high rate of fatty acid reesterification in adipocytes (16,34). However, because adipocytes from AKO mice had lower rates of lipolysis, the demand for fatty acid reesterification in these cells was likely reduced, allowing glucose to be diverted to oxidation instead of glycerol production. This could also partially explain the reduction in circulating levels of lactate in AKO mice. However, our conclusions are limited by the small sample size in our metabolic analysis and increasing the sample size could potentially uncover additional alterations in the metabolomic profile of AKO mice. This would also allow more rigorous interpretation from statistical analysis of the data; yet, we believe that this observational study provided an excellent and interesting platform to identify candidate metabolites for validation in future functional studies.

Besides alterations in rates of lipolysis and lipogenesis in the WAT, energy intake and whole-body energy expenditure also determine adiposity. We did not detect any differences in food intake between WT and AKO mice. However, when oxygen consumption and spontaneous physical activity were measured, it was clear that both variables were much lower in AKO than WT mice, which is consistent with reduced energy expenditure and increased adiposity in these animals. Even though statistical significance was not achieved, RER was found to be slightly higher in AKO than WT mice, mostly during the dark cycle, suggesting that glucose was the substrate of preference in these mice. With attenuation of lipolysis, it makes sense for glucose to supply a larger proportion of whole-body energy. However, because energy expenditure was markedly reduced, the higher contribution of glucose to whole-body energy metabolism was likely not sufficient to prevent elevated glycemia when AKO mice were challenged with GTT and ITT. Previous studies have demonstrated that mice with enhanced NP signaling in WAT were protected against diet-induced obesity and insulin resistance and that these mice also exhibited increased energy expenditure and higher thermogenic activity in BAT (12). It appears that in our model, in which circulating ANP and WAT cGMP levels were reduced, such protective effect was lost, thus leading to increased adiposity and reduced capacity to clear glucose from the blood.

While uncovering new knowledge on the role of cardiac autophagy in interorgan cross talk, our study has several limitations and also calls for additional follow-up experimental work. First, the relatively low n, typically 4–6, and the variability that exists in the data set are acknowledged. Nevertheless, our conclusions are based on statistically significant changes. We carefully refer to the cardiomyocyte-specific AKO mice generated here as autophagy deficient, since there will remain some degree of autophagic capacity occurring. This could also be viewed as a strength of the model, as it more closely reflects lower levels, and not complete loss, of autophagy, which occur during disease pathogenesis. Finally, the detailed cellular mechanisms via which cardiomyocyte autophagy deficiency regulates ANP production and secretion are not explored in the current study, and this certainly warrants further detailed investigation.

In summary, our findings provide evidence that increased adiposity and insulin resistance in AKO mice were caused by a combination of reduced energy expenditure and impaired ANP signaling and lipolysis in WAT. These data provide further evidence of interorgan cross talk likely to be of pathophysiological significance in cardiometabolic diseases.

Funding. This work was supported by an operating grant to G.S. from Canadian Institutes of Health Research and grants to E.S. from National Natural Science Foundation of China (81900779) and to R.B.C. from Natural Sciences and Engineering Research Council of Canada. G.S. also gratefully acknowledges support from Heart & Stroke Foundation of Ontario via a Career Investigator Award.

Duality of Interest. No potential conflicts of interest relevant to this article were reported.

Author Contributions. E.S. performed most of the experimental work, researched the work, and helped plan protocols and experiments and with writing the manuscript. D.D.E., S.J., D.S-K., and H.H. conducted the experimental work included in the figures. V.V. wrote, reviewed, and edited the manuscript. M.L. analyzed data and reviewed the manuscript. M.B.W. and R.B.C. contributed to planning the study, analyzing data, and editing the manuscript. G.S. designed the project, supervised the experimental work, wrote the manuscript, and provided funding. G.S. is the guarantor of this work and, as such, had full access to all the data in the study and takes responsibility for the integrity of the data and the accuracy of the data analysis.