Flies were maintained at 25°C, 60% relative humidity and 12 h light/dark cycles. Female flies (∼1 week of age) were reared on the standard cornmeal and yeast-based diet (0.8% cornmeal, 10% sugar, and 2.5% yeast) or the HFD (standard cornmeal and yeast-based diet supplemented with 20% w/v coconut oil). To induce gene expression in the GeneSwitch lines, RU food was made by adding 100 ul of 200 μM RU486 (Mifepristone, Cayman Chemical #100063171) onto the surface of fly food. Fly stocks used in this study were as follows: Hand4.2-Gal4 (a gift from Rolf Bodmer), TinCΔ4-Gal4 (a gift from Rolf Bodmer), Hand. myo-Gal4 (a gift from Hui-Ying Lim), Hand-GS-Gal4 (a gift from Rolf Bodmer), da-GS-Gal4 (a gift from Véronique Monnier), UAS-rictor RNAi (#1-BDSC #31527, #2-BDSC #36584), rictor ^Δ24 (a gift from Jongkyeong Chung), UAS-rictor-OE (a gift from Ernst Hafen), UAS-Akt RNAi (BDCS #82957), UAS-Drp1 RNAi (#1-BDSC #51483, #2- BDSC #27682), UAS-Drp1-OE (BDSC #51647), Drp1 ^[CR00300] (BDCS #79236). yw ^R flies (a gift from Eric Rulifson) were used as control or wild-type flies.

About 20 fly hearts per condition were lysed with 20 mM Tris-HCl (pH 8.0), 100 mM NaCl buffer supplied with protease inhibitor at 4°C for 20 min. After centrifugation at 14,000 rpm for 30 min, the supernatants were denatured with Laemmli sample buffer at 95°C for 5 min. Proteins were separated by Mini-PROTEAN^® TGX Precast Gels (Bio-Rad Laboratories, Inc.), transferred to PVDF membranes, immunoblotted with the following primary and secondary antibodies, and visualized with Pierce ECL Western Blotting Substrate. The antibodies used in western blot were: anti-phospho-Akt (Ser473) (Cell signaling technology #4060, 1:1000), anti-Akt (pan) (Cell signaling technology #4691, 1:2000), anti-beta-actin (Cell signaling technology #4967, 1:2000), Peroxidase AffiniPure Donkey Anti-Rabbit IgG (Jackson ImmunoResearch # 711-035-152, 1:5000).

Each fly heart sample was dissected in Schneider’s Drosophila medium (ThermoFisher #217-20024) and fixed with 4% paraformaldehyde (Electron Microscopy Sciences #15710) for 20 min at room temperature. Then, the samples were washed with 1x Phosphate Buffered Saline with 0.3% Triton X-100 (Fisher scientific #151-500) (0.3% PBST). After blocking the samples with 5% normal donkey serum (NDS, Jackson ImmunoResearch #017-000-121) for 1 h under room temperature (diluted in 0.3% PBST), the samples were stained with primary antibody diluted in 5% NDS for 16–24 h at 4°C. On the next day, after washing with 0.3% PBST, the samples were incubated in secondary antibody for 1–2 h at room temperature. The samples were then washed and mounted on the slides using ProLong Diamond Antifade Mountant (Thermo Fisher #P36961). The samples were imaged with an FV3000 Confocal Laser Scanning Microscope (Olympus). The following antibodies were used in immunostaining: anti-phospho-Akt (Ser473) (Cell signaling technology #4060, 1:200), anti-ATP5A1 for mitochondrial staining (Thermo Fisher #439800, 1:200), anti-Drosophila Drp1 (a gift from Leo Pallanck, 1:200), anti-mouse IgG-Alexa Fluor^® 488 (Jackson ImmunoResearch #715-545-150, 1:400); anti-rabbit IgG-Alexa Fluor^® 594 (Jackson ImmunoResearch #711-585-152, 1:400). DAPI (Fisher scientific #NC1526409) or Hoechst 33342 (Thermo Fisher #H3570) was used for nuclear staining. F-actin was stained with Phalloidin-CF^®647 (Biotium #00041T, 1:40).

The images were processed and analyzed by using Olympus cellSens Dimensions software and ImageJ/Fiji. For phospho-Akt quantification, the area around the nucleus of cardiomyocytes was selected as the region of interest (ROI), the mean intensity of the fluorescence was quantified using the count and measure function in cellSens Dimensions software. For mitochondrial diameter measurement, the A2 or A3 segment of the heart was selected as the ROI. Then the fluorescent threshold was adjusted to highlight mitochondria, followed by mitochondrial size correction using the object split tool in cellSens Dimensions. The maximum diameter of each mitochondrion in the ROI was quantified using the count and measure function. The maximum diameter is defined as the longest distance between two boundary points of the object. The proportion of elongated mitochondria with a maximum diameter greater than 2 μm was presented in each figure. See Supplementary Figure S1 for the workflow of mitochondrial diameter quantification. The colocalization between Drp1 and mitochondrial marker ATP5A1 in each ROI was measured by using the colocalization function of the cellSens and presented as Pearson correlation coefficient values.

Mitochondrial complex I and IV activities were measured according to a previous method (Spinazzi et al., 2012). To isolate mitochondria, about 20 flies were first homogenized in a glass dunce grinder containing sucrose homogenization buffer (20 mM Tris-HCl, 40 mM KCl, 2 mM EDTA). The fly homogenate was cleared by centrifugation at 600 x g for 10 min at 4°C twice. The supernatant was transferred to a new centrifuge tube each time. After centrifugation at 14,000 × g for 10–20 min, the mitochondrial-enriched fraction in the pellet was resuspended with 10 mM Tris-HCl. Mitochondrial extracts were frozen–thawed for three times prior to the complex I and IV activity assay. For complex I activity assay, about 300 μL of complex I assay buffer (50 mM of potassium phosphate buffer pH 7.5, 0.5 mg/ml of bovine serum albumin, 0.3 mM of potassium cyanide, 0.1 mM of NADH) and 4 μg of mitochondrial extracts were mixed in 96-well plates for each sample. Six μl of 10 mM ubiquinone (Sigma #C7956) was added to each sample to initiate the reaction. Complex I inhibitor rotenone was added separately to calculate complex I specific activities. The absorbance at 340 nm was recorded for 20 min using an Agilent BioTek Gen5 plate reader. In complex IV activity assay, 200 μL Complex IV activity buffer (80ul ddH2O, 100ul 0.1 M ph7.0 potassium phosphate buffer, and 150 μL 0.5 mM reduced cytochrome C) and 0.5 μg of mitochondrial extracts were mixed in 96-well plates for each sample. Complex IV inhibitor potassium cyanide was used to obtain complex IV specific activities. The absorbance at 550 nm was recorded for 5 min using an Agilent BioTek Gen5 plate reader. The complex I and IV activities were calculated by subtracting the activity in the presence of inhibitors from the total activity following the below equation:

Complex I and IV activity (μmol min⁻¹ mg⁻¹) = (Δ Absorbance/min × 1,000)/[(extinction coefficient × volume of sample used in ml) × (sample protein concentration in mg ml⁻¹)].

The fly hearts were prepared according to previously described semi-automatic optical heartbeat analysis (SOHA) (Fink et al., 2009). The high-speed heart contraction movies at the rate of 100 frames per second were captured using the Hamamatsu ORCA-Flash4.0 digital CMOS camera (Hamamatsu Photonics) and the Olympus BX51WI microscope with a 10X water immersion lens. The cardiac contractile defects, including the non-contractility and partial conduction block, were analyzed according to a previous study (Ocorr et al., 2009; Birse et al., 2010). For a non-contractile heart, the heart shows weak or no contractile. The heart with partial conduction block beats at a different rate in different regions of the heart. Fractional shortening was calculated as (diastolic diameters—systolic diameters)/diastolic diameters.

GraphPad Prism was used for statistical analysis. Unpaired two-tailed Student’s t test was performed to compare the mean value between control and treatment groups. Two-way ANOVA followed by Tukey’s multiple comparison test was used to examine the effects of diet and gene knockdown on mitochondrial diameter/function and cardiac function. Chi-square test was performed to examine the relationship between diet (or genotype) and contractile defects. The outliers were excluded using robust regression and outlier removal method (Q = 10%) prior to the data analysis.

To investigate the role of mTORC2 in HFD-induced mitochondrial fission in Drosophila hearts, we first examined how HFD treatment regulates mTORC2 activity. Wild-type flies were fed on 20% (w/v) coconut oil for 2 days before the analysis of Akt phosphorylation, a commonly used marker for mTORC2 activation metabolism (Saxton and Sabatini, 2017). Similar to a previous study (Diop et al., 2015), we found that short-term HFD treatment (2 days) significantly induced Akt phosphorylation in cardiomyocytes compared with normal diet (ND) treatment via immunostaining analysis (Figures 1A,B). This finding was further verified by western blot analysis on dissected fly hearts (Figures 1C,D). Together, these results suggest that HFD treatment induces mTORC2 activity in fly hearts.

Consistent with the previous studies (Filippi et al., 2017; Chen et al., 2018; Tong et al., 2019), we found that HFD treatment induced mitochondrial fragmentation in wild-type fly hearts (Figures 1E,F). Mitochondrial fragmentation was determined by counting the proportion of elongated mitochondria with a maximum diameter greater than 2 μm in a selected ROI. To determine the role of mTORC2 in HFD-induced mitochondrial fragmentation, we monitored mitochondrial morphology in fly hearts with the knockdown of rictor, the key subunit of mTORC2 (Figures 1E,F). Hand4.2-Gal4 was used to drive heart-specific gene knockdown. Two different rictor RNAi lines were used to eliminate genetic background effects. We found that both rictor knockdown lines showed no HFD-induced mitochondrial fragmentation (Figures 1E,F). It is known that Hand4.2-Gal4 drives expression in both cardiomyocytes and pericardial cells. To determine the cardiac autonomous role of rictor, we performed rictor knockdown using two cardiomyocyte-specific drivers, Hand. myo-Gal4 (Liu et al., 2019) and TinCΔ4-gal4 (Chang et al., 2020). As shown in Supplementary Figures S2A,B, cardiomyocyte-specific knockdown of rictor also blocked HFD-induced mitochondrial fragmentation, suggesting rictor regulates mitochondrial fission in a cardiomyocyte-dependent manner. Furthermore, to avoid the potential effects of constitutive gene knockdown on heart development, we examined adult-onset knockdown of rictor using a GeneSwitch heart driver, Hand-GS-Gal4. We found that 5∼7 days of rictor knockdown in adult flies significantly blocked HFD-induced mitochondrial fragmentation (Supplementary Figure S2C). Lastly, since HFD increases mTORC2 activity, we wonder if overexpressing rictor alone is sufficient to induce mitochondrial fission. As shown in Figure 1G, overexpression of rictor did not alter mitochondrial size and morphology. Taken together, these results suggest that mTORC2 is required for HFD-induced mitochondrial fission.

Given the vital role of rictor in HFD-induced mitochondrial fission in fly hearts, we next asked whether rictor is involved in HFD-induced mitochondrial and cardiac dysfunction. To monitor mitochondrial function, we first measured the complex I activity. Complex I is the key component of the electron transport chain that is responsible for ATP production. We found that the complex I activity was reduced upon short-term HFD treatment (2 days), which might be due to reduced complex I abundance in fragmented mitochondria upon HFD treatment (Figure 2A). Interestingly, HFD treatment did not down-regulate complex I activity in rictor loss-of-function mutants, even though the complex I activity of rictor mutants was lower than wild-type flies before HFD treatment (Figure 2A). In addition, we measured complex IV activity to assess other electron transport processes and mitochondria function. We found that the complex IV activity was not affected by either HFD treatment or rictor mutants (Figure 2B). These results suggest that not all electron transport processes are altered by short-term HFD treatment, and mTORC2 specifically targets the complex I activity potentially through the regulation of mitochondrial fission upon HFD treatment.

Next, we carried out semi-automatic optical heartbeat analysis (SOHA) to determine whether mTORC2 is required for cardiac protection under HFD treatment. We fed control flies and rictor knockdown flies on either ND or HFD for 7 days prior to the SOHA analysis, as there are no obvious cardiac defects found upon short-term (2 days) HFD treatment (Birse et al., 2010). The incidence of abnormal contractility (e.g., partial conduction block and non-contractility) and altered fractional shortening were monitored accordingly to a previous study (Birse et al., 2010). In wild-type flies, HFD treatment slightly (but not significantly) increased the incidence of abnormal contractility (X ² (1, N = 43) = 2.633, p > 0.1)) (Figure 2C). In contrast, knockdown of rictor significantly increased the incidence of contractile defects (especially partial conduction block) upon HFD treatment (X ² (1, N = 43) = 5.135, p < 0.01)) (Figure 2C). Cardiac-specific knockdown of rictor also significantly decreased fractional shortening upon HFD treatment, a common measurement for heart muscular contractility (Figure 2D). Taken together, these results suggest that mTORC2 is required for cardiac protection upon HFD treatment, which might be due to its positive role in mitochondrial fission.

Next, we investigated the mechanisms for how mTORC2 controls mitochondrial fission under HFD treatment. Given that Akt phosphorylation is induced under HFD treatment, we decided to test whether mTORC2 regulates mitochondrial fission through its downstream effector Akt. We knocked down Akt in fly hearts using Hand4.2-Gal4 driver and monitored mitochondrial fragmentation upon HFD treatment. Surprisingly, we found that knockdown of Akt did not block HFD-induced mitochondrial fragmentation (Figures 3A,B), which suggests that Akt is not responsible for HFD-induced mitochondrial fission. In addition, we examined whether Akt knockdown affects mitochondrial and cardiac function. Unlike rictor knockdown, we found that Akt knockdown did not alter the down-regulation of complex I activity and fractional shortening by HFD treatment (Figures 3C,D). Thus, mTORC2 may control mitochondrial fission through a different downstream target.

To explore the factors involved in rictor-regulated mitochondrial fission upon HFD treatment, we examined the fission protein, Drp1. The correlation between mTORC2 activity and Drp1 activation has been previously reported (Kim et al., 2016). Since the recruitment of Drp1 to the mitochondria is one of the essential steps of mitochondrial fission, we co-stained Drp1 and a mitochondrial marker (ATP synthase subunit ATP5A1) to monitor the Drp1 recruitment to mitochondria under HFD treatment. As expected, HFD treatment increased the colocalization of Drp1 and mitochondrial marker ATP5A1, while rictor knockdown blocked HFD-induced mitochondrial localization of Drp1 (Figures 4A,B).

Similar to rictor knockdown, we found that knockdown of Drp1 in fly hearts blocked HFD-induced mitochondrial fragmentation (Figures 4C,D). Two Drp1 RNAi lines were used. We also showed that Drp1-mediated regulation of mitochondrial fission is cardiac autonomous by knocking down Drp1 using two cardiomyocyte-specific drivers, Hand. myo-Gal4 and TinCΔ4-gal4 (Supplementary Figures S2D,E). Furthermore, adult-onset knockdown of Drp1 also blocked HFD-induced mitochondrial fragmentation (Supplementary Figure S2F). To determine whether Drp1 acts as the downstream effector of mTORC2 to regulate HFD-induced mitochondrial fission, we conducted an epistasis analysis using a combined fly line, Hand-GS-Gal4; rictor RNAi, and crossed it with control or Drp1 overexpression flies. As shown in Figure 4E, knockdown of rictor alone blocked HFD-induced mitochondrial fragmentation, while overexpression of Drp1 restored HFD-induced mitochondrial fragmentation in rictor knockdown flies. Together, our findings suggest that mTORC2 regulates HFD-induced mitochondrial fission through Drp1 recruitment to mitochondria.

To further examine the role of Drp1 in the regulation of HFD-induced mitochondrial and cardiac dysfunction, we measured complex I and IV activities, contractile defects, and fractional shortening of the Drp1 knockdown and mutant flies. Similar to rictor mutants, HFD treatment did not reduce complex I activity of Drp1 loss-of-function (Figure 5A), while the complex IV activity was not affected by both HFD treatment and Drp1 mutants (Figure 5B). Furthermore, heart-specific knockdown of Drp1 exaggerated the detrimental effects of HFD on cardiac contractile function (X ² (1, N = 50) = 7.837, p < 0.01) (Figure 5C) and fractional shortening defects (Figure 5D). Taken together, these results suggest that Drp1 might be an important downstream effector of mTORC2 in protecting hearts under HFD treatment.