Introduction

Cells must appropriately allocate available nutrients to optimize their metabolic programs− for energy extraction and for the generation of building blocks that enable cell growth. The balance between these processes not only maintains cellular health under varying nutritional conditions but also plays an important role in determining cell fate and function (Baker & Rutter, 2023; DeBerardinis & Thompson, 2012; Ghosh-Choudhary et al., 2020; Metallo & Vander Heiden, 2013). For example, hepatocyte metabolism changes considerably between fed and fasted conditions. In the fed state, hepatocytes use the majority of their nutrients to synthesize proteins, lipids, and glycogen, which results in increased cell size and liver biomass (Kast et al., 1988; Reinke & Asher, 2018; Sinturel et al., 2017). To meet the energy demands of other tissues during fasting, the liver undergoes a metabolic shift to produce glucose from biosynthetic precursors, thereby decreasing hepatocyte and liver size (Lang et al., 1998; Sinturel et al., 2017).

The metabolic pathways that support biosynthetic metabolism can be inappropriately activated in diseases such as cancer and heart failure to promote pathological growth. For example, many of the primary oncogenic adaptations in tumors prioritize anabolic metabolism over ATP production, which facilitates rapid cell proliferation (Lunt & Vander Heiden, 2011; Vander Heiden et al., 2009; Zhu & Thompson, 2019). Metabolic rewiring during heart failure similarly results in greater biosynthetic potential and less efficient ATP production, which in post-mitotic cardiomyocytes leads to increased cell size and insufficient cardiac pumping (Bornstein et al., 2024; Henry et al., 2024; Weiss et al., 2023).

The fate of pyruvate, which is primarily generated from glucose via glycolysis in the cytoplasm, is a critical node that can determine the balance between energetic and biosynthetic metabolism (Baker & Rutter, 2023; Yiew & Finck, 2022). In most differentiated cells, the majority of pyruvate is transported into the mitochondria via the mitochondrial pyruvate carrier (MPC) complex, a heterodimer composed of MPC1 and MPC2 proteins (Bricker et al., 2012; Herzig et al., 2012). Once in the mitochondria, pyruvate is converted to acetyl-CoA by the pyruvate dehydrogenase complex (PDH), fueling the tricarboxylic acid (TCA) cycle and supporting efficient ATP production. In cancer, stem, and other proliferative cells, more pyruvate is converted to lactate and exported from the cell, a process that regenerates NAD⁺, a cofactor necessary for glycolytic flux (Baker & Rutter, 2023; Lunt & Vander Heiden, 2011; Zhu & Thompson, 2019). In some specialized cells, including hepatocytes, pyruvate is imported into mitochondria but converted to oxaloacetate, which can feed the TCA cycle but also serves as a precursor for glucose production via gluconeogenesis (Holecek, 2023b; Jitrapakdee et al., 2008). We and others have demonstrated that these alternative fates of pyruvate−energy generation, cell proliferation, or glucose production−differentially impact the metabolic and fate decisions of multiple cell types in varying contexts (Bensard et al., 2020; Cluntun et al., 2021; Wei et al., 2022; Yiew & Finck, 2022). Loss of MPC function, which shifts pyruvate metabolism toward lactate production and thus expedites glycolysis and the production of biosynthetic precursors, has been shown to increase cell proliferation in mouse and Drosophila intestinal stem cells as well as in various tumors (Bensard et al., 2020; Schell et al., 2017; Zangari et al., 2020). MPC expression is also reduced in human and mouse models of heart failure, and genetic deletion of the MPC in cardiomyocytes is sufficient to induce hypertrophy and heart failure (Cluntun et al., 2021; Fernandez-Caggiano et al., 2020; McCommis et al., 2020; Zhang et al., 2020). Conversely, MPC overactivation or overexpression restricts intestinal stem cell proliferation and limits the growth of cardiomyocytes under hypertrophic stimuli, with excess mitochondrial pyruvate fueling the TCA cycle (Bensard et al., 2020; Schell et al., 2014; Schell et al., 2017). These observations suggest that mitochondrial pyruvate metabolism is central to cell proliferation as well as the size and specialized functions of post-mitotic cells.

The mechanisms regulating cell size include well-characterized cellular signaling pathways and transcriptional programs (Grewal, 2009; Liu et al., 2022; Lloyd, 2013). The CDK4-Rb pathway monitors cell size in proliferating cells by coupling cell growth with cell division (Amodeo & Skotheim, 2016; Ginzberg et al., 2018; Tan et al., 2021; Zhang et al., 2022). In response to insulin and other growth factors, the PI3K/Akt pathway activates mTORC1, leading to increased biosynthesis of proteins, lipids, and nucleotides, and consequently, increased cell size (Gonzalez & Hall, 2017; Gonzalez & Rallis, 2017; Saxton & Sabatini, 2017). The pro-growth transcription factor Myc drives a gene expression program that enhances metabolic activity and protein synthesis, resulting in larger cells (Baena et al., 2005; Dang, 1999; Iritani & Eisenman, 1999; Stine et al., 2015; van Riggelen et al., 2010). However, we only partially understand how metabolic pathways regulate physiological (or pathophysiological) growth, particularly in cells that have distinct and specialized roles in organismal metabolism.

Here, we investigated whether pyruvate metabolism influences biosynthetic capacity and cell size, using the Drosophila fat body as a model of the mammalian liver. We found that MPC overexpression and increased mitochondrial pyruvate transport restrict cell growth and limit protein synthesis in larval fat body cells and in spheroids of human liver-derived cells. Higher MPC expression resulted in smaller cells, not by increasing TCA cycle flux as observed in other cell types, but instead by redirecting mitochondrial pyruvate metabolism towards gluconeogenesis. A key driver of this metabolic rewiring is a reduced cellular redox state, which disrupts the biosynthesis of TCA cycle-derived amino acids, such as aspartate and glutamate, ultimately reducing protein synthesis. These observations highlight how cells with specialized functions, like hepatocytes, employ distinct metabolic adaptations to respond to organismal demands under varying nutritional conditions.

Results

Increased mitochondrial pyruvate transport reduces the size of Drosophila fat body cells

The Drosophila fat body is functionally analogous to mammalian white adipose tissue and liver, serving as a buffer to store excess nutrients in fat droplets and glycogen and deploy them to support the animal with fuel during times of fasting (Arrese & Soulages, 2010; Musselman et al., 2013). During larval development, fat body cells halt cell division and dramatically increase in size during the third instar stage (from 72 to 120 hours After Egg Laying (AEL)) (Edgar & Orr-Weaver, 2001; Zheng et al., 2016) (Fig. 1a-c). As a first step to understand the metabolic programs that enables this rapid cell growth, we performed RNA-sequencing of the Drosophila fat body across this developmental period. We observed a time-dependent change in mRNAs encoding proteins that have well-characterized roles in supporting cell growth, including components of the insulin and mTORC1 signaling pathways and the Myc transcriptional network which correlated with increased cell size (Supplementary Fig. 1a). Amongst metabolic genes, we observed modest differences in those that function in amino acid synthesis and fatty acid metabolism (Supplementary Fig. 1b). The abundances of mRNAs encoding proteins involved in glycolysis, oxidative phosphorylation (OxPhos), and the TCA cycle were distinctly altered in fat bodies during development (Supplementary Fig. 1b). Since pyruvate metabolism is the central node connecting carbohydrate metabolism and the TCA cycle, we studied the abundances of mRNAs that encode proteins that regulate pyruvate metabolism (Fig. 1d). We found that the expression of genes that link pyruvate to the TCA cycle were reduced including, Pyk, the Drosophila pyruvate kinase homolog, which converts phosphoenolpyruvate into pyruvate; Mpc1 and Mpc2, which encode the two subunits of the MPC, which transports pyruvate into the mitochondrial matrix; as well as Pdha and Dlat, which encode subunits of PDH complex (Fig. 1e). In contrast, some mRNAs encoding proteins that regulate pyruvate abundance were upregulated such as Pepck and Pepck2, which make phosphoenolpyruvate from oxaloacetate (Supplementary Fig. 1c). Based on these data, we hypothesized that the suppression of mitochondrial pyruvate metabolism, which is gated by the action of the MPC, might support the rapid cell growth observed in fat body cells.

Figure 1:

To test this hypothesis, we prevented the downregulation of the MPC during larval development (Fig. 1e) by ectopically expressing Mpc1 and Mpc2 (termed “MPC+” Supplementary Fig. 2a, 2b). The sustained expression of the MPC (both Mpc1 and Mpc2) throughout the fat body significantly reduced the rate of cell growth compared to a GFP-expressing control (Supplementary Fig. 2c). Given the important role of the fat body in controlling organismal growth, we wanted to assess the cell-autonomous effects of MPC expression using mosaic analysis in individual fat body cells (Supplementary Fig. 2d) (Ito et al., 1997). We generated GFP-labeled clones with expression of the MPC (MPC+), which we confirmed by immunofluorescence (Supplementary Fig. 2e, f). The MPC-expressing, GFP-positive cells were significantly smaller in size compared with either mock clones (control) or neighboring GFP-negative cells within the same tissue. Mpc1 knock down (Mpc1-KD) clones, in contrast, were marginally larger (Fig. 1f-g). These results demonstrate that sustained expression of the MPC in developing fat body cells is sufficient to prevent cell growth in a cell-autonomous manner.

If the effects of MPC expression were related to the mitochondrial transport of pyruvate, then limiting the production of pyruvate should mitigate these effects. Therefore, we measured cell size in larvae raised on either a normal (9% sugar) diet or a diet with no added sugars, which limits the production of pyruvate from glucose and fructose. Limiting dietary sugars significantly reduced the size of control clones but increased the size of MPC-expressing clones. Notably, the size of MPC-expressing fat body clones was comparable to that of control clones when larvae were grown in sugar-limited medium, suggesting that limiting pyruvate synthesis abolishes the effect of MPC expression on fat body cell size (Fig. 1h, i). Mpc1-KD clones were again larger than control cells and their size was unaffected by the sugar-limited diet. Inhibiting MPC activity by feeding larvae a normal diet supplemented with the MPC inhibitor UK5099 ameliorated the cell size effects of MPC expression (Fig. 1j and Supplementary Fig. 2g). These results indicate that pyruvate transport into mitochondria inversely correlates with the size gain in fat body cells. These observations also suggest that the suppression of mitochondrial pyruvate import, and metabolism is required for the rapid cell size expansion observed in the fat body during larval development.

The size of a cell is determined by its content of proteins, nucleic acids, and lipids (Bjorklund, 2019; Lloyd, 2013; Schmoller & Skotheim, 2015). To understand how MPC expression affects the abundance of these macromolecules, we dissected fat bodies from control and fat body-wide MPC-expressing larvae at 120 hours AEL and quantified DNA, RNA, triacylgylcerides, and protein. Control and MPC+ fat bodies had equivalent DNA content (Fig. 1k, Supplementary Fig. 2h) and similar levels of EdU incorporation (Supplementary Fig 2o, p), suggesting that MPC expression does not impact DNA endoreplication. RNA and lipid content were modestly decreased in MPC+ fat bodies compared with control tissues (Fig. 1k, Supplementary Fig. 2i, j), although the number of lipid droplets was higher with MPC expression (Supplementary Fig 2l-n). In contrast, MPC expression dramatically decreased protein abundance in fat bodies (Fig. 1k, Supplementary Fig. 2k), and reduced protein synthesis as assessed by staining for the puromycin analog, O-propargyl-puromycin (OPP) (Deliu et al., 2017; Villalobos-Cantor et al., 2023) (Fig. 1l, m). These data suggest that MPC-mediated mitochondrial pyruvate import decreases protein synthesis, which likely contributes to the reduced size of MPC-expressing cells. Conversely, in developing fly larvae, repression of the MPC and the subsequent decrease in mitochondrial pyruvate appear to provide a metabolic mechanism to support a rapid expansion in cell size.

Growth factor signaling pathways are hyperactivated in MPC expressing cells

The best understood mechanisms that govern cell size involve conserved signaling and transcriptional networks (Bjorklund, 2019; Grewal, 2009; Lang et al., 1998; Lloyd, 2013). For example, the mTORC1 pathway coordinates both extracellular and intracellular growth regulatory signals to dictate the synthesis and degradation of macromolecules, including proteins, lipids, and nucleic acids (Gonzalez & Hall, 2017; Gonzalez & Rallis, 2017; Saxton & Sabatini, 2017). mTORC1 increases protein synthesis through the phosphorylation of several proteins, including S6 kinase (S6K) and 4EBP1 (Fig. 2a). Since MPC-expressing clones were smaller in size and had reduced protein synthesis compared with control clones, we assessed mTORC1 activity in the fat body using our mosaic expression system. As a control, we confirmed that S6k (the Drosophila gene encoding S6K) over-expressing clones were larger in size and had elevated phospho-S6 staining compared with wild-type clones (Fig. 2b, c). Surprisingly, MPC+ clones also had elevated phospho-S6 staining (Fig. 2b, c), suggesting that despite their small size, these cells have high mTORC1 activity. Over-expression of Rheb, which is an upstream activator of mTORC1, resulted in clones that were larger than wild-type controls and which had increased phospho-4EBP1 staining (Fig. 2d, e). Again, even though the MPC+ clones were smaller in size, we observed robust p-4EBP1 staining, indicating that mTORC1 is hyperactive in these cells.

Figure 2:

Gene set enrichment analysis of RNA-sequencing data from MPC-expressing fat bodies showed enrichment for signatures associated with pro-growth signaling (Supplementary Fig. 3a). We assessed markers of several of these pathways in MPC+ clones. In addition to the mTORC1 pathway, Myc increases cell growth by regulating the transcription of ribosome subunits and biosynthetic metabolic genes. We found that MPC+ clones had an elevated abundance of the Myc protein, both in the cytoplasm and nucleus, (Fig. 2f, g), suggesting that the growth-promoting Myc transcriptional program is active in these cells. MPC+ clones also had reduced expression of the transcription factor Foxo, consistent with its downregulation by pro-growth signaling pathways (Supplementary Fig. 3b). Finally, we stained for phospho-eIF2α to assess the activity of the integrated stress response, which restricts global protein synthesis. Starvation robustly induced the integrated stress response in control clones, but MPC+ clones exhibited no evidence of activation of this pathway under normal growth conditions (Fig. 2h, i). Collectively these data indicate that conventional pro-growth pathways are activated in MPC-expressing clones, which is incongruent with the small size of the clones. This suggests that mitochondrial pyruvate metabolism controls cell size via alternative molecular mechanisms.

We next performed genetic epistasis analysis to better understand the relationship between MPC expression and the mTORC1, PI3K, and Myc pathways. Activation of the PI3K or mTORC1 pathways, via over-expression of PI3K, increased the size of control but not MPC-expressing fat body cells (Supplementary Fig. 3c). Similarly, activation of mTORC1 via knock down of its inhibitor, Tuberous Sclerosis Complex 1 (Tsc1), increased the size of control cells but had no effect on MPC+ cell size (Supplementary Fig. 3d). Over-expression of Myc, on the other hand, increased the size of control and MPC+ fat body clones to a similar degree (Supplementary Fig. 3e). Knock down of Myc was sufficient to decrease cell size to a similar extent as MPC expression; however, Myc knock down had no additional effect on cell size in MPC+ clones (Supplementary Fig. 3f). These results suggest that neither mTORC1, PI3K, nor Myc are epistatic to MPC and suggest that MPC likely acts independently of these canonical pathways to regulate the size of fat body cells.

Increased mitochondrial pyruvate transport reduces the size of HepG2 cells and spheroids

Since the Drosophila fat body has many features reminiscent of the mammalian liver, we tested whether MPC expression might similarly restrict cell size in cells derived from this tissue. We engineered HepG2 cells, which were originally isolated from a hepatocellular carcinoma, to express epitope tagged-MPC1 and MPC2 in a doxycycline-inducible manner. We observed expression of ectopic MPC1 and MPC2 starting at four hours after doxycycline treatment, which increased over the duration of the time course, peaking at 24 hours post-induction (Fig. 3a). We stained doxycycline-treated cells with phalloidin and measured cell size using microscopy images taken at two-hour intervals. Induction of MPC expression coincided with a significant reduction in cell size, which was first apparent six hours post-doxycycline treatment and was sustained for the remainder of the time course (Fig. 3b). Doxycycline had no effect on the size of control HepG2 cells (EV). Treatment with the MPC inhibitor UK5099 for 24 hours markedly increased the size of control cells and partially reversed the small size phenotype of MPC-expressing cells (Fig. 3c). We also assessed cell volume by analyzing a 3D reconstruction of confocal images of phalloidin-stained HepG2 cells and found that the cell volume was lower in MPC+ cells (Supplementary Fig. 4a).

Figure 3:

We have observed that the physiological consequences of altered pyruvate metabolism are more apparent in cells grown in a three-dimensional culture environment (Schell et al., 2014; Wei et al., 2022). Using our doxycycline-inducible cells, we found that MPC expression resulted in spheroids that were significantly smaller than control spheroids as assessed by microscopy (Fig. 3d, e). Cells from MPC-expressing spheroids were also smaller as shown by flow cytometry (Fig. 3f, g). When compared with control (EV) spheroids, expression of MPC had no effect on the number of cells per spheroid, on the cell cycle phase distribution, or on the number of apoptotic or necrotic cells (Supplementary Fig. 4b, 4c, and 4d). These analyses suggest that the smaller size observed in MPC expressing spheroids is not due to effects on cell proliferation or cell death. Consistent with our observations in the Drosophila fat body, protein content was lower in MPC-expressing HepG2 spheroids compared with EV controls, whereas there was no difference in the abundances of DNA, RNA, or triacylgylcerides (Fig. 3h and Supplementary Fig. 4e-h). To directly assess the effect of MPC expression on protein synthesis, we induced the MPC in HepG2 cells and treated cells acutely with a low concentration of puromycin to label nascent proteins. We found that MPC expression decreased the incorporation of the amino acid analog L-homopropargylglycine (HPG) (Shen et al., 2021; Tom Dieck et al., 2012) into nascent proteins, as assessed by fluorescence microscopy (Fig. 3i, j). Protein synthesis was similarly reduced in MPC-expressing HepG2 spheroids (Fig. 3k, l). These observations were further supported by the decreased abundance of the short-lived, destabilized GFP (d2GFP) (Li et al., 1998; Pavlova et al., 2020) in MPC-expressing HepG2 spheroids compared with controls. This reduction in the level of d2GFP was prevented by treatment with the MPC inhibitor UK5099 (Fig. 3m). Together these data suggest that increased transport of pyruvate into mitochondria, mediated by MPC expression, reduces protein synthesis and cell size in both fly and mammalian models.

Excess mitochondrial pyruvate promotes gluconeogenesis

We have previously shown that loss of the MPC reduces the contribution of glucose and pyruvate to the TCA cycle (Bensard et al., 2020; Cluntun et al., 2021). To investigate how MPC expression impacts carbohydrate metabolism in HepG2 cells, we performed metabolic tracing using ¹³C-glucose (Fig. 4a). We observed that MPC expression reduced the labeling fraction of the glycolytic intermediates 3-phosphoglycerate (Supplementary Fig. 5a) and pyruvate (Supplementary Fig. 5b), as well as alanine (Supplementary Fig. 5c), which is derived from pyruvate, all of which suggests a reduction in glycolysis in these cells. Once imported into mitochondria, glycolytic-derived pyruvate has two major fates: conversion to acetyl-CoA by PDH or to oxaloacetate by pyruvate carboxylase (PC). These fates can be differentiated by assessing the abundances of M+2 (derived from PDH) and M+3 (derived from PC) TCA cycle intermediates (Fig. 4a). We found that both M+2 and M+3 isotopomers of TCA cycle intermediates were modestly increased following MPC expression (Fig. 4b and 4c; Supplementary Fig. 5d-g). The fact that we observe increases in TCA cycle intermediates despite decreased glycolytic labeling in pyruvate suggests that the apparent labeling through PDH and PC might be underestimating the effect on metabolic flux through these enzymes. Thus, it appears that MPC expression increases the activity of both enzymes that utilize pyruvate. Typically, when TCA cycle flux is increased, one observes an increase in activity of the electron transport chain (ETC) to oxidize the resulting NADH. However, MPC expression had no impact on ETC activity as assessed by measuring oxygen consumption (Fig. 4d). The implications of elevated NADH production without a concomitant increase in NADH oxidation will be discussed in Fig. 5.

Figure 4:

Figure 5:

To probe the impact of increased flux through the PC and PDH reactions, we conducted genetic epistatic analysis in fat body cells. We found that Drosophila fat body clones in which we over-expressed Pcb (the Drosophila gene encoding pyruvate carboxylase) were significantly smaller than controls and were equivalent in size to MPC+ clones (Fig. 4e, f). Clones expressing both the MPC and Pcb were even smaller (Fig. 4e, f). Conversely, knock down of Pcb (Pcb-KD) in MPC-expressing fat body clones completely rescued the cell size phenotype (Fig. 4e, f and Supplementary Fig. 6a). Knock down of Pcb in MPC-expressing HepG2 cells also eliminated the small cell size phenotype (Supplementary Fig. 6b). These data suggested that the small size of cells expressing the MPC is likely due to increased flux through the PC reaction. Consistent with this, knock down of the PDH-E1 (Drosophila gene Pdha) or PDH-E2 (Drosophila gene Dlat) subunits of PDH, which should divert mitochondrial pyruvate, so it is preferentially used by PC, also resulted in smaller fat body cells (Supplementary Fig. 6c-d). In contrast, activating PDH by knocking down the inhibitory PDH kinases (mammalian PDKs, Drosophila gene Pdk) (Stacpoole, 2017; Wang et al., 2021), which should promote the flux of pyruvate through PDH and away from PC, rescued cell size in MPC-expressing fat body clones (Supplementary Fig. 6c, d). Similarly, in MPC-expressing HepG2 cells, activation of PDH via treatment with the PDK inhibitors DCA and AZD7545 was sufficient to restore cell size (Supplementary Fig. 6e) (Stacpoole, 2017; Wang et al., 2021). These analyses suggest that the reduction in cell size observed with MPC expression is due to PC-mediated metabolism of pyruvate.

The product of PC, oxaloacetate, has three major metabolic fates: 1) feeding the TCA cycle via citrate synthase as discussed above, 2) conversion to aspartate by glutamatic-oxaloacetic transaminase 2 (GOT2), and 3) conversion to phosphoenolpyruvate by PEPCK2 leading to gluconeogenesis, synthesis of glucose (Jitrapakdee et al., 2008; Kiesel et al., 2021). Sustained expression of the MPC in fat bodies increased the concentration of glucose in this tissue (Fig. 4g) as well as in larval circulation (known as hemolymph) (Fig. 4h), suggesting that MPC expression increases glucose production in fat body cells. Next, we tested whether PEPCK2-mediated gluconeogenesis was elevated in these cells compared with controls. We used ¹³C-lactate to trace the ¹³C-labeling of phosphoenolpyruvate and resultant glucose synthesis via gluconeogenesis (Fig. 4i). We found that the relative abundance of M+3 phosphoenolpyruvate was higher in MPC-expressing cells compared with controls (Fig. 4j), suggesting increased activity of PEPCK2. The gluconeogenic pathway employs a series of biochemical reactions to convert phosphoenolpyruvate into glucose, which is then excreted from the cell. This production of M+6 glucose from lactate was also higher in MPC-expressing HepG2 cells (Fig. 4k). To test whether gluconeogenesis contributes to the small size phenotype of MPC-expressing cells, we knocked down enzymes in the pathway (Drosophila phosphoenol carboxykinase, Pepck2, and fructose bisphosphatase, Fbp) and assessed cell size in Drosophila and HepG2 models. Knockout of PEPCK2 in MPC-expressing HepG2 cells resulted in smaller spheroids (Fig. 4l) and cells (Supplementary Fig. 6f). Knock down of either Pepck2 or Fbp partially rescued the size defect in MPC-expressing Drosophila fat body clones (Fig. 4n, o Supplementary Fig. 6g), and Pepck2 knock down also increased protein synthesis in these clones (Fig. 4p, q). Collectively, these data suggest that the cell size and protein synthesis phenotypes observed in MPC-expressing cells require PC-mediated gluconeogenesis, and this relationship is found in both the Drosophila fat body and in HepG2 cells.

A redox imbalance impairs protein synthesis and cell growth in MPC-expressing cells

We were intrigued by the observation that MPC expression exerted its impact on cell size through PC and not via increased flux through PDH. We did not observe any differences in the abundances of PDH1a, phosphorylated (inactive) PDH1a, PC, PEPCK2, or G6PC proteins in MPC-expressing HepG2 cells (Supplementary Fig. 7a, b). We therefore hypothesized that the shift in pyruvate metabolism in response to MPC expression might be driven by changes in the abundances of enzyme cofactors, metabolite regulators, or cellular redox balance. PC and PDH show reciprocal regulation by such factors: PC utilizes ATP as a substrate and is allosterically activated by acetyl-CoA, whereas PDH is inhibited by ATP, acetyl CoA, and NADH (Chai et al., 2022; Sugden & Holness, 2011) (Supplementary Fig. 7c). In addition, several reactions in gluconeogenesis require ATP or NADH (Hausler et al., 2006; Siess et al., 1977) (Supplementary Fig. 7c). Upon induction of MPC expression in HepG2 cells, we observed increased abundance of acetyl-CoA (Supplementary Fig. 7d), a higher ATP to ADP ratio (Supplementary Fig. 7e), a greater abundance of NADH without an increase in NAD⁺, and thus an increase in the cellular NADH/NAD⁺ ratio (Supplementary Fig. 7f-h). To determine how MPC expression impacts the subcellular distribution of redox factors, we separated cytoplasmic and mitochondrial fractions from control or MPC-expressing Drosophila fat bodies and measured the NADH/NAD⁺ ratio. We found that MPC expression increased the NADH/NAD⁺ ratio in both the cytoplasm and mitochondria (Fig. 5a). We observed a similar increase in the NADH/ NAD⁺ ratio in both fractions in MPC-expressing HepG2 spheroids (Fig. 5b). These results suggest that the increased abundances of acetyl-CoA, ATP, and NADH in MPC-expressing cells could promote the rewiring of mitochondrial pyruvate metabolism through PC and support gluconeogenesis.

To test the hypothesis that ATP or NADH concentrations might affect cell size in MPC-expressing Drosophila fat body clones or HepG2 cells, we utilized pharmacological or genetic modulation of these molecules. Treatment with gramicidin, which decreases the ATP to ADP ratio (Xue et al., 2022), did not alter the size of MPC-expressing HepG2 cells (Supplementary Fig. 7i). We used several orthogonal approaches to reduce the cellular NADH/NAD⁺ ratio in the MPC-expressing systems and measured their effect on cell size. Co-expression of the Drosophila nicotinamide mononucleotide adenylyl transferase (Nmnat), which increases NAD⁺ biosynthesis, almost normalized the small size phenotype of MPC-expressing clones in the Drosophila fat body (Fig. 5c, d). We observed a similar rescue of cell size following expression of the Ciona intestinalis alternate Complex I enzyme NADH dehydrogenase (NDX) (Gospodaryov et al., 2020) or the yeast NADH dehydrogenase (NDI) (Sanz et al., 2010), both of which oxidize NADH to NAD⁺ without concomitant proton translocation and energy capture (Fig. 5e, f). Expression of NDI in HepG2 cells also mitigated the effect of MPC expression on spheroid size (Supplementary Fig. 7j, k), as did treatment with duroquinone (Merker et al., 2006) (Supplementary Fig. 7l), which oxidizes NADH to NAD⁺. To extend these investigations to three dimensional culture, we supplemented the growth medium of MPC-expressing HepG2 spheroids with the NAD⁺ precursors nicotinamide riboside (NR) or nicotinamide mononucleotide (NMN), both of which recovered spheroid size (Fig. 5g, h).

Since MPC expression reduced protein synthesis in both Drosophila fat bodies and HepG2 cells, we tested how cellular redox status might contribute to this phenotype in both systems. Expression of NDX, which lowers the NADH/NAD⁺ ratio, increased translation in MPC-expressing Drosophila fat body clones (Fig. 5i, j). In HepG2 spheroids, boosting NAD⁺ biosynthesis by supplementing growth media with NR or NMN partially rescued the abundance of destabilized GFP (Fig. 5k). These results suggest that the elevated NADH/ NAD⁺ ratio in MPC-expressing cells limits protein synthesis and that normalizing that ratio increases protein synthesis and cell size.

Reduced amino acid abundance impairs size of MPC-overexpressing cells

Given the reduced protein synthesis observed upon MPC expression, we assessed amino acid concentrations in HepG2 using steady-state metabolomics. We found that MPC expression reduced the abundance of most amino acids (Fig. 6a). To test whether the low abundances of amino acids contribute to the smaller size of MPC-expressing cells, we supplemented the growth media of HepG2 spheroids with excess non-essential amino acid (NEAAs)—either two or three times the recommended dilution of a commercially available amino acid cocktail that includes glycine, L-alanine, L-asparagine, L-aspartate, L-glutamate, L-proline, and L-serine. MPC-expressing spheroids grown with excess NEAAs were comparable in size to controls (Fig. 6b, c). In parallel, we provided Drosophila larvae with food containing excess (5x) amino acids from 72 to 120 hours AEL, which partially rescued cell size in MPC-expressing fat body clones (Fig. 6d, e). To genetically augment intracellular amino acids, we expressed the amino acid importer slimfast (Colombani et al., 2003) in control or MPC-expressing fat body clones and found that it prevented the decrease in cell size (Fig. 6f, g).

Figure 6:

To determine which amino acid(s) contribute to the cell size effects of MPC expression, we cultured control and MPC-expressing HepG2 cells in media supplemented with an excess of each individual amino acid from the NEAA cocktail. Treatment with excess glycine, L-alanine, or L-serine had no effect on cell size (Fig. 6h). However, the size of MPC-expressing cells was normalized by supplementation with L-aspartate, L-glutamate, or L-proline (Fig. 6h), all of which are derived from TCA cycle intermediates. Supplementation with either L-aspartate or L-glutamate also rescued the small size phenotype of MPC-expressing HepG2 spheroids (Supplementary Fig. 8a, b). Increasing L-aspartate uptake by over-expressing the aspartate transporter SLC1A3 also recovered size of HepG2 spheroids (Supplementary Fig. 8c, d). In addition, treatment with excess (3x) NEAA partially restored the abundance of d2GFP in MPC-expressing HepG2 spheroids (Supplementary Fig. 8e), suggesting rescued protein synthesis.

Like glutamate and proline, aspartate is derived from a TCA cycle intermediate, specifically via transamination of oxaloacetate by glutamic-oxaloacetate transaminase 2 (GOT2). Since GOT2 and PEPCK2 both use oxaloacetate as a substrate, we hypothesized that knocking down GOT2 might phenocopy MPC expression by driving PEPCK2-mediated conversion of oxalacetate into phosphoenolpyruvate and suppressing aspartate biosynthesis. Knock down of Got2 (the Drosophila gene encoding GOT2) reduced cell size in Drosophila fat body clones and phenocopied MPC-expressing cells (Fig. 6i, j). Similarly, both GOT2 knock out and MPC-expressing HepG2 cells were smaller than EV(Supplementary Fig. 8f). Expression of the MPC in these GOT2 knock down systems had no significant impact on cell size (Fig. 6i, j and Supplementary Fig. 8f), suggesting that the effects of MPC expression and GOT2 knock down act similarly to limit amino acid synthesis and cell size. We next performed the reciprocal experiment by over-expressing Got2 to favor the production of aspartate from oxaloacetate. Got2 expression normalized cell size in both MPC-expressing fat body clones and in HepG2 spheroids (Fig. 6i, j and Supplementary Fig. 8g, h). The efflux of aspartate from mitochondria into the cytoplasm is a critical component of the malate-aspartate shuttle, which is a major redox shuttle in human cells. To test whether increasing the abundance of aspartate would ameliorate the high NADH/NAD⁺ ratio observed in MPC-expressing cells, we supplemented growth media with exogenous aspartate and assessed cellular redox status. We found that excess aspartate reduced the cellular NADH/NAD⁺ ratio in these cells such that it was comparable with control cells (Fig. 6k). We observed similar results when we treated these cells with NMN or UK5099 (Fig. 6k). Moreover, all these treatments increased protein synthesis in MPC-expressing cells (Fig. 6l, m).

Mitochondrial pyruvate import reduces the size of rat primary hepatocytes

Although HepG2 cells exhibit some features of hepatocytes, they are a transformed, immortalized, and proliferative hepatocellular carcinoma cell line. We wanted to test whether the MPC expression phenotypes that we observed in Drosophila fat bodies and HepG2 cells could be recapitulated in a more physiologically relevant mammalian model. We chose primary rat hepatocytes, which have been used extensively to interrogate hepatocyte cell metabolism and signaling and also have the advantage of being genetically tractable. We expressed MPC1 and MPC2 in cultured primary rat hepatocytes (Fig. 7a), and, consistent with our results in other systems, we found that expression of MPC reduced cell size (Fig. 7b, c) and decreased protein synthesis (Fig. 7d). MPC-expressing primary hepatocytes also had a higher NADH to NAD⁺ ratio in both the cytoplasmic and mitochondrial fractions (Fig. 7f). We assessed gluconeogenesis in primary hepatocytes by quantifying glucose in the culture media following incubation with the gluconeogenic precursors, pyruvate, and lactate. Glucose production was higher in MPC-expressing hepatocytes compared with controls (Fig. 7e). Treatment with UK5099 eliminated this effect and reduced glucose production to a similar rate in both cells (Fig. 7e). These results demonstrate that augmented mitochondrial pyruvate in hepatocytes, and related cells in Drosophila, drives a metabolic program that results in an increased NADH/NAD⁺ ratio. This scenario results in accelerated gluconeogenesis, decreased protein synthesis, and educed cell size.

Figure 7:

Discussion

We investigated whether pyruvate metabolism influences biosynthetic capacity and cell size in the Drosophila fat body and in HepG2 spheroids. During the third instar phase of Drosophila larval growth, when cells are rapidly expanding in size, we observed a profoundly decreased expression of the MPC in liver-like fat body cells. We found that this rewiring of pyruvate metabolism is essential for cell growth as forced maintenance of MPC expression resulted in dramatically smaller cells. By combining Drosophila genetic analyses and metabolomics studies in HepG2 cells and spheroids, we demonstrated that excess mitochondrial pyruvate elevates the cellular NADH/NAD⁺ ratio and redirects carbohydrate metabolism to favor gluconeogenesis over glycolysis (Fig. 7g). This shift reduces the availability of oxaloacetate for aspartate and glutamate biosynthesis, triggering a broader imbalance in amino acid abundances within the cell. We conclude that altering the fate of pyruvate to support biomass accumulation is required for the cell size expansion that occurs during fat body development. We speculate that this phenomenon also applies to the mammalian liver, which is the closest analog of the Drosophila fat body, as both HepG2 cells and primary rat hepatocytes show similar effects following ectopic MPC expression.

Why does simply reorienting the metabolism of pyruvate have this profound effect on cell growth? Our data suggest that a central mediator of the phenotype is an elevated NADH/NAD⁺ ratio, which likely results from MPC expression driving an acceleration of the TCA cycle, as evidenced by an increase in the abundances of M+2 isotopomers of TCA cycle intermediates.

Although the increase in labeled succinate, fumarate, and malate is modest, it occurs despite a reduction in glycolytic labeling. This suggests that less labeled pyruvate feeds the TCA in MPC-expressing cells compared with controls and that we are likely underestimating the actual increase in flux. The TCA cycle generates NADH and appears to do so more actively in cells with ectopically expressed MPC. However, our oxygen consumption data suggest that the oxidative phosphorylation system in these cells does not have the capacity to increase its activity in response to the enhanced availability of mitochondrial pyruvate and an increased NADH/NAD⁺ ratio. As a result, the increased TCA cycle flux and limited ETC activity together elevate the NADH/NAD⁺ ratio in both the mitochondria and cytoplasm, disrupting cellular redox balance leading to a rewiring of cellular metabolism.

This redox situation causes two distinct but related perturbations that appear to both contribute to decreased cell growth. First, we observed a clear depletion of amino acids and evidence of decreased synthesis of amino acids that are primarily derived from TCA cycle intermediates (aspartate, glutamate, and proline). A reduced NAD⁺ pool impairs the capacity of cells to synthesize aspartate (Birsoy et al., 2015; Sullivan et al., 2015; Sullivan et al., 2018), which is used to synthesize glutamate and proline and which plays a crucial role in maintaining redox balance in both the mitochondria and cytoplasm (Alkan et al., 2018; Holecek, 2023a; Lieu et al., 2020; Wei et al., 2020; Yoo et al., 2020). Replenishing any of these TCA cycle-derived amino acids via genetic or nutritional increases in their availability was sufficient to reverse the effect of ectopic MPC expression on cell size. Thus, amino acid depletion is a key driver of the small size phenotype. Second, the increased NADH/NAD⁺ ratio also drives a particular metabolic program that favors the conversion of mitochondrial pyruvate to oxaloacetate and a subsequent increase in gluconeogenesis. This program is enforced by allosteric regulation via NADH, acetyl-CoA, and ATP, which act on PC and several enzymes of glycolysis and gluconeogenesis. It also appears to be important for the small size of MPC-expressing cells since loss of any of several steps along the gluconeogenic pathway, particularly PC and PEPCK2, mitigates or eliminates the cell size phenotype observed with ectopic MPC expression in fat body cells and HepG2 spheroids. It is important to note that gluconeogenesis is a critical function of these cells, and its rate is carefully controlled in response to varying physiological stimuli. Surprisingly, constitutive MPC expression is sufficient to supersede the control enacted by physiological and hormonal signals and enact the loss of biomass and impaired cell size.

We have previously shown that loss of the MPC enhanced the stem cell identity and proliferation of intestinal stem cells, whereas ectopic MPC expression had the opposite effect (Bensard et al., 2020; Schell et al., 2017). Such observations are consistent with our findings that MPC expression inversely correlates with biomass accumulation. More recent data demonstrated that enhanced MPC activity prevented the increase in cell size that occurs in cardiomyocytes in response to hypertrophic signaling (Cluntun et al., 2021; Fernandez-Caggiano et al., 2020). In these cases, the fate of mitochondrial pyruvate that determines cell growth is its oxidation in the TCA cycle. In contrast, our studies of hepatocytes and related cells described herein, demonstrate that the fate of pyruvate that suppresses growth in cell size is not oxidation by the TCA cycle, but rather the production of glucose, which starts with its conversion to oxaloacetate by PC in the mitochondria. Cardiomyocytes have low expression of PC compared with liver, where this enzyme (as well as others including PEPCK2) serves a vital role during fasting by producing glucose, which fuels the brain and other organs that require glucose for their survival and function (Hatting et al., 2018; Petersen et al., 2017; Rui, 2014). This is an example of how the unique metabolic physiology of specialized cells plays a critical role in maintaining organismal homeostasis.

One conclusion from the data presented herein is that a cell’s metabolic program plays a decisive role in determining the growth of that cell. It was striking how rapidly expression of the MPC impacted cell size in the HepG2 model. Essentially, as soon as MPC over-expression became detectable, the population of cells started to show a decrease in size. Notably, the decrease in cell size following ectopic MPC expression occurred despite the upregulation of multiple pro-growth signaling networks. It is unclear why these networks are hyperactivated in this context as there is no evidence from our data indicating any nutrients are in excess abundance. We hypothesize that there may be regulators of cell size that recognize when cells are inappropriately small and engage these pathways to increase cell size. mTORC1, PI3K, and Myc pathways typically promote biomass accumulation and increased cell size but fail to do so when mitochondrial pyruvate is elevated. MPC expression reduced the abundance of amino acids, and this appears to play a dominant role to impair protein synthesis and prevent the cell growth effects expected following hyperactivation of the mTORC1 and Myc pathways. Thus, our data suggest that the metabolic fate of pyruvate can override canonical pathways that mediate cell size—such as mTORC1 signaling.

We demonstrate that the appropriate partitioning of pyruvate metabolism maintains the redox state of a cell to support the accumulation of biomass that is necessary for its specialized function. Increased mitochondrial pyruvate metabolism in cells from the fly “liver-like” fat body disrupts these processes, causes cells to perform excess gluconeogenesis, and prevents cell growth. As a result, the Drosophila larvae became hyperglycemic and experience developmental delay. The abundances of MPC1 and MPC2 are upregulated in mouse livers during starvation and in high-fat diet conditions, which correlates with increased rates of gluconeogenesis in both circumstances. Conversely, loss of the MPC in liver impairs gluconeogenesis (Gray et al., 2015; McCommis et al., 2015; Yiew & Finck, 2022). Moreover, liver dysfunction in diabetes and hepatic steatosis are driven by reductive stress and an elevated NADH/NAD⁺ ratio (Goodman et al., 2020; Jokinen & Luukkonen, 2024). We are intrigued by the possibility that the fate of pyruvate might have profound consequences on the redox state and gluconeogenic capacity of the mammalian liver, including by functioning as part of the metabolic milieu that drives unrestrained gluconeogenesis in diabetes. Our discovery that mitochondrial pyruvate regulates the cellular redox state, thereby controlling biosynthesis, offers insights for developing therapeutic strategies for these and other diseases.

Methods

Drosophila Strains and Handling

Drosophila melanogaster stocks were maintained at 25°C on semi-defined fly food composed of 20 g agar, 80 g baker’s yeast, 30 g sucrose, 60 g glucose, 0.5 g MgSO4, 0.5 g CaCl2, 11 ml tegosept and 6 ml propionic acid. This was base medium for all Drosophila experiments, but specific fly food modifications are mentioned in text and figure legends. To induce clones in fat bodies, synchronized eggs were transferred to 29°C for 4 days until dissection. For experiments involving genetic manipulation of all fat body cells, tubGal80^ts20 was used to restrict activity of CG-Gal4 at 18°C for 120 hours (57 hours equivalent time at 25°C) and larvae were shifted to 29°C till dissection at specified time points.

Following fly stocks were procured from Bloomington stock center UAS-MPC1-P2A-MPC2 (28812582), CG-Gal4; tubGal80^ts20, hs-Flp1.22 (BDSC, #77928), Act>CD2>Gal4, UAS-GFP (BDSC, #4413), Act>CD2>Gal4, UAS-RFP (BDSC, #30558), UAS-S6K^STDETE (BDSC, #6914), UAS-Rheb^PA (BDSC, #9689), UAS-Myc^OE (BDSC, #9675), UAS-Myc^RNAi (BDSC, #25783), UAS-PI3K93E^.Excel (BDSC, #8287), UAS-PI3K93E^A2860C (BDSC, #8288), UAS-TSC1^RNAi (BDSC, #31314), UAS-hPC (BDSC, #77928), UAS-PC^RNAi (BDSC, #56883), UAS-PEPCK2^RNAi (BDSC, #36915), UAS-FBPase^RNAi (BDSC, #51871), UAS-Pdk^RNAi (BDSC, #35142), UAS-NMNAT (BDSC, #37002), UAS-CintNDX (BDSC, #93883), UAS-ScerNDI1 (BDSC, #93878), UAS-Slif (BDSC, #52661), UAS-GOT2^RNAi (BDSC, #78778). From VDRC UAS-MPC1^RNAi(KK) and from NIG UAS-Pdha^RNAi, UAS-Dlat^RNAi (PDH E2) were purchased.

Generation of Overexpression and CRISPR Mutant fly stock

Pc and Pepck2 deletion fly stocks were generated using CRISPR-Cas9 as described (Gratz et al., 2014). We targeted specific nucleotide sequences of the genes of interest through homology-directed recombination using two guide RNAs and inserted a dsRed construct that expresses in adult eyes to facilitate the selection of mutant flies. We deleted exon 1 to exon 5 using guide RNAs for Pc^-KO: 5′ guide ATACATTTAAGTCCTAGGC; 3′ guide TCGATTGATCCTGGAAACA.

Pepck2^-KO, being a single exon coding sequence, we generated complete deletion by using guides 5′ guide AAAGGGTGCACATCTGTGA; 3′ guide TTTGGGGCGTGGCCTAGAC. The plasmids were injected into y[1] M{GFP[E.3xP3]=vas-Cas9.RFP}ZH-2A w[1118] (BDSC, #55821) fly stock embryos (Bestgene) and one Pc-KO and five Pepck2-KO flies were picked up, confirmed for the dsRed expression in adult eyes and used in subsequent experiments.

To overexpress Got2, Got2 cDNA was amplified from RNA extracted from larval fat bodies using primer 5’ GAATTC ATGAGTAGAACCATTATTATGACGCTTAAGGAC, 3’ CTCGAGCTTGGTAACCTTGTGTATGCTCTCAGCCAGG. The cDNA was then cloned into a pUAST-aatB plasmid using EcoRI and XhoI restriction enzymes, and the construct was injected into pBac{yellow[+]-attp-9A}VK00005 (BDSC, #9725) embryos to obtain the insertion

Mosaic Analysis and Phalloidin Staining for Cell Size Analysis

Fat body clones were induced by the leaky expression of heat shock flippase 1.22 during embryonic stages, and the 2D size of fat body cells was analyzed at 120 hours after egg laying (AEL) using fluorescence microscopy. Cell size analysis was conducted as reported earlier (Toshniwal et al., 2019). Fat bodies were dissected from larvae at the specified time points in 1X PBS buffer (pH 7.2, Invitrogen, #10010049) and fixed in 8% paraformaldehyde (Sigma Aldrich, # P6148) for 1 hour at room temperature (RT). The tissues were then washed twice with 0.1% PBT (0.1% Triton X-100 (Sigma Aldrich, # X100) in 1X PBS buffer) for 10 minutes each. Subsequently, the tissues were incubated with Rhodamine Phalloidin (Thermo Scientific, # R418) at a 1:400 dilution in 1X PBS buffer for 2 hours at room temperature (RT). After staining, the tissues were washed once with 0.1% PBT and then with 1X PBS before being mounted in DAPI-supplemented VectaShield (Vector Labs, #H1200). Representative images were captured using a Laser Scanning Confocal Microscope (LSM 880, Carl Zeiss). For cell size analysis, images were captured with a fluorescence microscope (Carl Zeiss, Axio Vision) at 20X magnification, focusing on a plane where all nuclei were in focus. The 2D areas of fat body cells were measured using FIJI software. Cell membranes stained with Rhodamine Phalloidin were traced using the freehand tool, and the area of approximately 20-25 GFP-positive cells from fat bodies collected from about five animals was measured as one replicate. All cell size analyses were conducted in a blinded manner.

Immunofluorescence on Fat Body Clones

Larval fat bodies at 120 hours after egg laying (AEL) at 29°C were dissected in 1X PBS buffer (pH 7.2), fixed in 8% paraformaldehyde for 1 hour at room temperature (RT), and then washed three times in 0.1% PBT (0.1% Triton X-100 in 1X PBS buffer) for 10 minutes each. The tissues were blocked in 10% normal goat serum (NGS, Jackson ImmunoResearch Laboratories, #005-000-121) in 0.1% PBT for 1 hour at RT, followed by incubation with primary antibodies overnight at 4°C. Secondary antibody incubation was performed for 2 hours at RT. Following three to four washes in 0.3% PBT (10 minutes each) at RT, the tissues were mounted in DAPI-supplemented Vectashield (Vector Laboratories, #H1200). Images were captured using a Laser Scanning Confocal Microscope (LSM 880, Carl Zeiss).

Image Acquisition and Analysis

For all experiments, confocal images were captured using the same laser power and identical settings, with z-stacks of the dissected tissue taken at 1 μm intervals along the Y-axis. The images were analyzed using Fiji software, where a similar number of z-stacks focusing on the nuclei were projected at mean intensities. Using the freehand tool, cell membranes were outlined, and the mean fluorescence intensity (mean gray value) for each cell was recorded. To account for background fluorescence, the average mean gray values of the background (measured from regions of interest [ROI] of the same size) were subtracted from the recorded mean gray values of the cells. The resulting mean gray values, adjusted for background fluorescence, were then normalized, and the percent normalized mean gray values (with standard deviation, SD) of GFP-negative and GFP-positive cells were plotted.

Measurement of Protein Synthesis in Fat Bodies

Protein synthesis in Drosophila fat bodies was analyzed using the Click-iT Plus OPP Alexa Fluor 594 kit (Molecular Probes, #C10457). Fat bodies were dissected at 120 hours AEL in Shields and Sang M3 Insect Media (SSM3, Sigma, #S8398) and incubated with 5 mM O-propargyl-puromycin (OPP) in SSM3 media for 30 minutes at RT. Tissues were then washed three times with 1X PBS for 10 minutes each and fixed in 8% paraformaldehyde for 1 hour at RT. After fixation, the tissues were washed twice with 0.1% PBT (0.1% Triton X-100 in 1X PBS) supplemented with 0.5% bovine serum albumin (BSA) for 10 minutes each. The OPP was developed for 30 minutes at RT using the Click-iT reaction mixture, which included 88 μl of OPP Reaction Buffer, 20 μl of Copper (III) Sulfate (component D), 2.5 μl of component B, and 100 μl of Buffer Additive (component E), following the manufacturer’s instructions. The tissues were then washed twice in Reaction Rinse Buffer for 10 minutes each, followed by two washes in 0.1% PBT supplemented with 0.5% BSA for 10 minutes each at RT. After a final wash in 1X PBS, the fat bodies were mounted in DAPI-supplemented Vectashield. Images were captured using a confocal microscope (LSM 880, Zeiss).

LipidTOX Staining in Larval Fat Body Tissues

At 120 hours AEL, 3rd instar larvae were dissected in 1X PBS. The fat bodies were fixed in 8% paraformaldehyde for 1 hour at room temperature. After fixation, tissues were rinsed twice with 1X PBS. They were then incubated for 30 minutes in a 1:100 dilution of LipidTOX Red (Invitrogen, #H34351) in PBS, followed by two additional rinses with PBS. The tissues were mounted in DAPI-supplemented Vectashield and imaged using a confocal microscope. To quantify lipid droplet size, the diameters of lipid droplets from 35 fat body clones were measured using FIJI software.

EdU Incorporation in Fat Bodies

At 120 hours AEL, fat bodies were dissected in 1X PBS and incubated with 5 μM 5-ethyl-2’-deoxyuridine (EdU) in 1X PBS for 30 minutes at room temperature. The tissues were then washed three times with 1X PBS for 10 minutes each, followed by fixation in 8% paraformaldehyde for 1 hour at room temperature. After fixation, the tissues were washed with 0.1% PBT supplemented with 0.5% bovine serum albumin. EdU detection was performed using Click-iT Plus EdU Alexa Fluor 594 (Molecular Probes, #C10639) for 30 minutes at room temperature, according to the manufacturer’s instructions. Following development, the fat bodies were mounted in DAPI-supplemented Vectashield. Images were captured using a confocal microscope (LSM 880, Zeiss).

RNA Isolation and RNA Sequencing

To collect fat body tissues, w¹¹¹⁸ flies were mated at 25°C. The following day, flies were starved for one hour, and eggs were collected every 2 hours. The first collection was discarded, and the subsequent two collections were incubated at 25°C. After 20 hours, early hatched 1st instar larvae were discarded, and 1st instar larvae collected over a 2-hour period were kept at 25°C. Fat bodies from 10 male larvae at the specified time points were dissected and preserved in 1X PBS. For RNA isolation from MPC-overexpressing fat bodies at 120 hours AEL, RNA was extracted and purified using the NucleoSpin RNA kit (Takara Bio USA, Inc., #740955.50) with on-column DNA digestion, as per the manufacturer’s instructions. Four independent samples for each time point were prepared for sequencing.

Library preparation for poly(A)-selected RNAs was carried out using the Illumina RNA TruSeq Stranded mRNA Library Prep Kit with oligo-dT selection. Sequencing was performed using the Illumina NovaSeq Reagent Kit v1.5 (150 bp paired-end reads) at the High-Throughput Genomics Core Facility at the University of Utah. The raw sequencing data were analyzed using the BDGP6.28 genome and gene feature files. Differentially expressed genes were identified using DESeq2 version 1.30.0 with a 5% false discovery rate. RNA quality control, library preparation, and sequencing were performed by the University of Utah Huntsman Cancer Institute High Throughput Genomics and Bioinformatics Shared Resource. RNA-seq data from this study are available at NCBI GEO.

QPCR

500 ng RNA was used to make cDNA using Superscript II reverse transcriptase (Molecular Probes, #18064-022), dNTP (Molecular Probes, #18427-088) and oligodT (Molecular Probes, #18418012). QPCR analyses were performed on cDNA as described using PowerUp SYBR Green Master Mix (Applied Biosystems, #2828831) on QuantStudio 7 Flex (Applied Biosystems) instrument. Fold changes in transcript level were determined using the ΔΔCt method. Transcript levels were normalized to rp49. Each experiment was performed using 4–5 independent samples. Following primers were used to do qPCR. List of primers is provided in Appendix 3

Hemolymph Glucose and fat body glucose measurement

Glucose concentrations in larval hemolymph and fat bodies were measured as previously described (Ugrankar-Banerjee et al., 2023). To isolate hemolymph, 10 third instar larvae were selected from culture tubes, thoroughly washed to remove any food residues, and then dried. Hemolymph was collected by bleeding the larvae on a parafilm strip using Dumont 5 forceps (Fine Science Tools, #11254-20). Two microliters of the colorless hemolymph were transferred to a 96-well plate and mixed with 200 μl of Autokit Glucose reagent (Wako, #997-03001). For measuring intercellular glucose, fat bodies were dissected from 10 larvae per genotype and homogenized in approximately 300 microliters of ice-cold 1X PBS using a 29G1/2 syringe. The lysates were inactivated at 70°C for 10 minutes, then centrifuged at maximum speed at 4°C. Thirty microliters of the lysate were mixed with 170 μl of Autokit Glucose reagent. The plates were incubated at 37°C for 30 minutes, and absorbance was measured at 600 nm. Glucose concentrations were determined based on the absorbance values recorded for glucose standards.

HepG2 Cells – Knock down and Overexpression Strategies

HepG2 cells were purchased from ATCC and maintained in EMEM supplemented with 10% FBS and 1% PenStrep at 37°C in a 5% CO₂ atmosphere. For inducible overexpression of human MPC1 and MPC2, the HA-MPC2-P2A-T2A-MPC1-FLAG sequence was cloned into the pLVX-TetOne-Zeocin vector. Lentiviral particles were generated using Gag-Pol, pMD2.G, and VSVG packaging plasmids.

Viral particles were produced by co-transfecting 293T cells with the respective packaging plasmids using polyethylenimine (PEI, Sigma, #765090) as the transfection reagent at a 3:1 mass ratio of PEI to DNA. The virus-containing medium was collected 48 hours post-transfection, filtered through a 0.45 μm filter, and added to HepG2 cells cultured in normal medium, along with polybrene (Sigma, #P1240000) at a concentration of 10 μg/ml. Transduced cells were selected with 10 μg/ml Zeocin (Gibco, #R25001) for 1 week, and the level of overexpression was assessed by western blotting.

For PC over-expression, PC coding sequence was amplified from mRNA isolated from HepG2 cells using primers 5’ GAATTC ATGCTGAAGTTCCGAACAGTCCATGGG, 3’ GGATCCCTCGATCTCCAGGATGAGGTCGTCACC and cloned into pLenti-CMV-Blast. Similarly, SLC1A3 cDNA was amplified with primers 5’ GGATCC ATGACTAAAAGCAATGGAGAAGAGC, 3’ CTACATCTTGGTTTCAATGTCGATGG and GOT2 coding sequence was amplified using primers 5’ GGATCC ATGGCCCTGCTGCACTCCGG, 3’ TCTAGACTTGGTGACCTGGTGAATGGCATGG and cloned in pLenti-CMV-Blast (addgene, #17486). The coding sequences of NDI (addgene, #72876) and d2GFP (addgene, #115665) were also cloned into pLenti-CMV-Blast vector. Cells were selected on 3 μg/ml blasticidin (Gibco, A1113903) for 3 days. To generate knock out cells, following gRNAs were used. PCg5e-GAAGCCTATCTCATCGGCCG CGG, PCg6e-CGAAGTCCGCTCGCTCAGAG AGG, PEPCK2g2e-ATCTCCACTAAGCACTCGCA GGG, PEPCK2g3e-CATGCGTATTATGACCCGAC TGG, GOT2ga-GAGTGGCCGGGTAAGCTGAGCAG AGG, GOT2gb-GGAGTGGACCCGCTCCGGAACAG TGG. The guides were annealed and clones in lentiCRISPRv2 with blasticidin (addgene, #83480) resistance using BsmBI.

HepG2 Cells and 2D Cell Size Analysis

HepG2 cells were cultured on 12 mm coverslips in a 24-well plate at a low density of 10,000 cells per well in Human Plasma-Like Medium (Gibco, #A4899101) supplemented with 10% FBS (Sigma, F0926) and 1% PenStrep (Thermo, #15140). The next day, treatments were initiated as described in the figure legends, including 1 μg/ml doxycycline (Sigma, #D5207), 10 μM UK5099 (Sigma, PZ0160-5MG), 10 mM glycine (Sigma, #G7403), 5 mM alanine (Sigma, #A7469), 5 mM asparagine (Sigma, #A4159), 5 mM aspartic acid (Sigma, #A7219), 5 mM glutamic acid (Sigma, #G8415), 5 mM proline (Sigma, #P5607), 5 mM serine (Sigma, #S4311), 10 μM AZD7545 (MedChemExpress, #HY-16082), 1 mM dichloroacetate (Sigma, #347795), 100 nM duroquinone (Sigma, #D223204), and 2 nM gramicidin (Sigma, #G5002).

For time-course experiments in Figure 3, cells were fixed at 2, 4, 6, 8, 10, 12, 18, and 24 hours after doxycycline treatment using 4% paraformaldehyde in 1X PBS for 20 minutes at RT. For all other experiments, cells were fixed 24 hours after doxycycline treatment. Following fixation, cells were washed once with 0.1% PBT (0.1% Triton X-100 in 1X PBS) for 10 minutes and incubated with Rhodamine Phalloidin at a 1:400 dilution in 1X PBS buffer for 20 minutes at RT. After a couple of washes in 1X PBS, cells were mounted in DAPI-supplemented VectaShield.

Images were captured using a fluorescence microscope (Carl Zeiss, Axio Vision) focusing on the plane of the cellular nuclei at 20X magnification, where all nuclei were in focus, and the 2D area of HepG2 cells was measured using FIJI software. All cell size analyses were conducted in a blinded manner.

For 3D cell volume analysis, images were captured using a Laser Scanning Confocal Microscope (LSM 880, Carl Zeiss). The red fluorescence signal was used for 3D reconstruction, and cell volume was measured using an ImageJ Macro code in FIJI.

Measurement of Protein Synthesis in HepG2 Cells

HepG2 cells were cultured in either 2D monolayers or spheroid forms. At the desired time point, cells were treated with 20 μg/ml puromycin (Sigma, #P4512) in Human Plasma-Like Medium (HPLM) for 30 minutes at 37°C. Following treatment, cells were washed with 1X PBS, and proteins were extracted using 1X RIPA buffer at 4°C. Protein concentrations were quantified, and 15 μg of total protein was separated by SDS-PAGE using standard methods. Proteins were then transferred onto a nitrocellulose membrane, and puromycin-labeled peptides were detected using an anti-puromycin [3RH11] antibody (Kerafast, #EQ0001) followed by incubation with an appropriate secondary antibody.

Protein synthesis in HepG2 cells was also analyzed using the Click-iT HPG Alexa Fluor 594 kit (Molecular Probes, #C10429). HepG2 cells were grown on 12 mm coverslips at a density of 10,000 cells per well and treated with 1 μg/ml doxycycline for 24 hours. Cells were then incubated with 50 μM L-homopropargylglycine (HPG) in methionine-free DMEM (Gibco, #21013024) for 30 minutes at 37°C. After incubation, cells were washed twice with 1X PBS for 2 minutes each and fixed in 4% paraformaldehyde for 20 minutes at room temperature.

For OPP staining, appropriately fixed cells were washed twice with 0.1% PBT (0.1% Triton X-100 in 1X PBS) supplemented with 0.5% bovine serum albumin (BSA) for 10 minutes each. HPG was detected using the Click-iT reaction mixture, which included 88 μl of OPP Reaction Buffer, 20 μl of copper solution (component D), 2.5 μl of component B, and 100 μl of Buffer Additive (component E), following the manufacturer’s instructions. Cells were washed twice in Reaction Rinse Buffer for 10 minutes each, followed by two washes in 0.1% PBT supplemented with 0.5% BSA for 10 minutes each at room temperature. After a final wash in 1X PBS, the coverslips were mounted in DAPI-supplemented Vectashield. Images of the cells were captured using a confocal microscope (LSM 880, Zeiss).

Analysis of Spheroids

HepG2 cells were cultured in ultra-low attachment 96-well plates (Costar, #7007) at a density of 10,000 cells per well in Human Plasma-Like Medium (HPLM) supplemented with 10% FBS and 1% penicillin-streptomycin. Treatments were applied as specified in the figure legends, including 1 μg/ml doxycycline, 5 mM aspartate, 5 mM glutamate, 10 μM UK5099, MEM non-essential amino acids (Gibco, #11140050), 100 nM nicotinamide riboside (Sigma, #SMB00907), and 10 μM NMN (Sigma, #N3501). Cells were incubated for 6 days at 37°C with CO2 and O2. Brightfield images of the spheroids were captured using a Zeiss Axio Observer Z1 microscope, and spheroid size was measured using FIJI software.

To quantify cell numbers, 12 spheroids from each condition (EV or MPC+) were pooled, dissociated by trypsinization, and the number of cells was counted using a CellQuant system (Bio-Rad).

For cell cycle analysis, cells were stained using Vybrant® DyeCycle™ Violet stain (Molecular Probes, #V35003). After pooling and dissociating 12 spheroids from each condition (EV or MPC+), cells were centrifuged and resuspended in 200 μL of 5 μM Vybrant® DyeCycle™ Violet stain in EMEM supplemented with 10% FBS. The staining was performed by incubating the cells at 37°C for 30 minutes, protected from light. Samples were analyzed using a BD Celesta flow cytometer with ∼405 nm excitation and ∼440 nm emission. The resulting FCS files were processed in FlowJo software, where the forward scatter of singlets was recorded, and median data were used to plot graphs. Vybrant dye staining was employed to assess the distribution of cells in the G1, S, and G2/M phases.

For apoptosis detection, an Annexin V/PI staining kit (Molecular Probes, #V13241) was used, with 5 μM camptothecin (MedChemExpress, #HY-16560) serving as a positive control. Sixteen spheroids were collected, dissociated with trypsin as described above, and resuspended in 200 μL of 1X annexin-binding buffer. Cells were incubated with 1 μl of Alexa Fluor™ 488 Annexin V (Component A) and 0.2 μl of 100 μg/ml PI working solution at room temperature for 15 minutes. Stained cells were then analyzed using a BD Canto flow cytometer, measuring fluorescence emission at 530 nm and 575 nm (or equivalent) with 488-nm excitation.

Biomolecule Separation and Measurement

For this experiment, biomolecules were extracted and measured from both Drosophila larvae fat bodies and HepG2 cell spheroids. Fat bodies were dissected from 10 male larvae at 120 hours after egg laying (AEL) and collected in 150 μl of 1X PBS. The samples were lysed by performing three freeze-thaw cycles. A total of 24 spheroids were homogenized in 150 μl of radioimmunoprecipitation assay (RIPA) buffer (50 mM Tris-HCl, 1% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 150 mM NaCl, and 2 mM EDTA).

Lysates were centrifuged at 7500 × g for 5 minutes to remove debris. The resulting supernatant was used for the following measurements. Triglyceride Measurement-For triglyceride analysis, 30 μL of the supernatant was incubated for 10 minutes at 75°C. Following this, 10 μl of homogenate from the spheroids or 2 μL from fat bodies was added to 200 μl of Triglycerides Reagent (Thermo Fisher Scientific, #TR22421). The mixture was incubated for 10 minutes at 37°C in 96-well microplates with gentle shaking. Absorbance was measured at 550 nm using the Synergy Neo2 multimode plate reader (BioTek).

Protein Measurement-Protein concentrations were determined by mixing 2 μl of the supernatant with 200 μl of BCA Protein Assay Reagent (Thermo Scientific, #23225). The mixture was incubated for 30 minutes at 37°C with gentle shaking in 96-well microplates. Absorbance was measured at 560 nm.

RNA and DNA Separation- The remaining 100 μl of the supernatant was processed for RNA and DNA separation using TRIzol reagent (1 ml; Invitrogen, # 15596026). To separate RNA, 0.2 ml of chloroform was added to the sample, followed by centrifugation at 12,000 × g for 15 minutes at 4°C. The RNA in the interphase was purified using 75% ethanol, following standard methods, and quantified using a NanoDrop spectrophotometer. DNA was extracted from the organic phase by adding 100% ethanol, followed by isopropanol precipitation. The resulting pellets were washed with 0.3 M guanidine hydrochloride in 95% ethanol, resuspended in 0.1 M sodium citrate in 10% ethanol (pH 8.5), and washed with 75% ethanol. Finally, the DNA pellets were resuspended in 0.1 ml of 8 mM NaOH by pipetting. The pH was adjusted to 7.2 with HEPES, and the DNA was quantified using a NanoDrop spectrophotometer.

Measurement Of NADH/NAD⁺ Protocol

For HepG2 cells, 1×10⁶ cells treated with 1ug/ml dox for 24 hours were scraped in 1.5 ml tube. For NADH/NAD⁺ ratio analysis from spheroids, 18 spheroids were treated with 1ug/ml dox for 6 days were resuspended in 1mL 1XPBS. Scraped cells or pooled spheroids were centrifuged (13,500xg, 10 s, 4°C) and tresuspended in 250 ul lysis buffer. Fat bodies from male 10 larvae were dissected in 1X PBS and resuspended in lysis buffer provided with kit. To separate cytoplasmic and mitochondrial fractions by rapid subcellular fractionation, lysates were centrifuged (13,500xg, 10 s, 4°C) and the supernatant was collected for the cytosolic fraction, while the remaining pellet contained the mitochondria. Mitochondrial pellet was resuspended in 100 ul lysis buffer.

NADH to NAD+ ratios were measured using Amplite Fluorimetric NAD⁺/NADH ratio assay kit (AAT Bioquest, #15263) as directed by instructions provided. Briefly, 25uL of cytoplasmic or mitochondrial fractions were mixed with either NADH or NAD⁺ extraction solution. Samples were incubated at 37°C for 15 minutes. Later 25ul of either NAD⁺ or NADH extraction was added which was followed by incubation with 75ul mix of NADH sensor buffer and NAD⁺/NADH recycling enzyme for 1 hour at RT. Fluorescence intensity was recoded at 540 nm excitation and 590 nm emission.

Protein Extraction and Western Blotting

HepG2 cells were directly scraped into RIPA supplemented with protease and phosphatase inhibitors (Roche Molecular, #04906845001). For spheroids, 18 spheroids were pooled, washed with 1X PBS, and then incubated in RIPA buffer with the protease and phosphatase inhibitor cocktail. After 45 minutes on ice with vertexing every 15 minutes, lysates were centrifuged at 16,000xg for 15 minutes at 4°C to remove insoluble material.

Protein concentration was measured using the Bicinchoninic Acid (BCA) protein assay (Thermo Fisher Scientific, 23225). Samples were mixed with 4x sample loading buffer and heated at 95°C for 5 minutes. Protein samples (15 µg) were separated on SDS-polyacrylamide gel electrophoresis (SDS-PAGE) at 20 mA per gel, transferred onto a 0.45 µm nitrocellulose membrane (GE Healthcare) using the Mini Trans-Blot module (Bio-Rad) at 120 V for approximately 2 hours.

The membrane was blocked with 5% non-fat milk (Serva) in Tris-buffered saline with 0.05% Tween 20 (TBS-T) for 1 hour. It was then incubated overnight with primary antibodies diluted in TBS-T. The next day, the membrane was washed with TBS-T and incubated with fluorophore-conjugated secondary antibodies in TBS-T for 1 hour. Following additional washes with TBS-T, fluorescence was detected using the Odyssey CLx imaging system (LI-COR Biosciences) and analyzed using FIJI software. Antibodies used are listed in Appendix 1.

Steady-State Metabolomic Studies

10 million HepG2 cells with MPC+ and EV control expression were grown in 2D culture were treated with 1ug/ml dox in HPLM supplemented with 10% HPLM. After 24 hours, culture medium was collected and quenched with 1:4 volume of 100% methanol. Cells were washed and quenched with 1 ml of 80% methanol in water. Cell lysates were then subjected to three rapid freeze-thaw cycles and then spun at 16,000xg for 10 min at 4°C. The supernatants were evaporated using a SpeedVac concentrator. Each sample or treatment were with 4 to 5 replicates.

¹³C-glucose tracing for M+3 vs M+2 ratio of TCA cycle metabolism

10 million HepG2 cells with MPC+ and EV control expression were grown in 2D culture and were treated with 1ug/ml dox in HPLM. After 24 hours, culture medium was changed to ¹³C-glucose tracing media: glucose-free HPLM supplemented with 4.5g/L ¹³C glucose and 10% dialyzed FBS. 0 hour, 1 hour, 2 hours, and 4 hours later, cells were washed and quenched with 1 ml of 80% methanol in water. Cells were scrapped out in Methanol and lysates were then subjected to three rapid freeze-thaw cycles and then spun at 16,000xg for 10 min at 4°C. The supernatants were evaporated using a SpeedVac concentrator.

¹³C-Lactate Tracing for Gluconeogenesis Assay

10 million HepG2 cells with MPC+ and EV control expression were grown in 2D culture were treated with 1ug/ml dox in DMEM without glucose, without glutamine (Gibco, #A1443001) with 10% dialyzed FBS. After 16 hours, culture medium was replaced with 6 ml of 1ug/ml dox in DMEM without glucose, without glutamine and without FBS. 3 hours later, culture medium was replaced with 6 ml of 1ug/ml dox and 20mM lactate (Sigma, #490040) containing DMEM without glucose or glutamine or FBS. After 4 hours, 300 μl culture medium was collected and quenched with 1:4 volume of 100% methanol. Cells were washed and quenched with 1 ml of 80% methanol in water. Cell lysates were then subjected to three rapid freeze-thaw cycles and then spun at 16,000xg for 10 min at 4°C. The supernatants were evaporated using a SpeedVac concentrator. Each cell types had 4 to 5 replicates.

Gas Chromatography-Mass Spectrometry Derivatization

Dried metabolites were derivatized and prepared for Gas chromatography following standard methods. Dried samples were resuspended in 30 μl of anhydrous pyridine with methoxamine hydrochloride (10 mg/ml) and incubated at RT overnight. Next day, the samples were heated at 70°C for 15 min and centrifuged at 16,000g for 10 min. The supernatant was transferred to a pre-prepared gas chromatography–mass spectrometry autoinjector vial with 70 μl of N-(tert-butyldimethylsilyl)-N-methyltrifluoroacetamide (MTBSTFA) derivatization reagent. The samples were incubated at 70°C for 1 hour, after which aliquots of 1 μl were injected for analysis. Samples were analyzed using either an Agilent 6890 or 7890 gas chromatograph coupled to an Agilent 5973N or 5975C Mass Selective Detector, respectively. The observed distributions of mass isotopologues of glucose, pyruvate, citrate, succinate, aspartate, glutamate, malate, fumarate, phosphoenolpyruvate were corrected for natural abundance.

Liquid chromatography-mass spectrometry

Following standard methods, dried metabolites were resuspended in 100 μl of 0.03% formic acid in analytical-grade water, vortexed, and centrifuged to remove insoluble material. 20 μl of supernatant was collected and injected to AB SCIEX QTRAP 5500 liquid chromatography/triple quadrupole mass spectrometer (Applied Biosystems SCIEX). Chromatogram review and peak area integration were performed using MultiQuant (version 2.1, Applied Biosystems SCIEX). The peak area for acetyl CoA, ADP, ATP, NAD⁺, NADH was normalized against the total ion count of that sample.

Oxygen consumption rate

Oxygen consumption rates (OCR) were measured using an XFe96 Extracellular Flux Analyzer (Agilent) according to the manufacturer’s instructions. Cells were plated at a density of 60,000 cells per well in Seahorse microplates (Agilent) and allowed to adhere for 6 hours. MPC was induced with 1 μg/ml doxycycline overnight. Afterward, the cell culture media was removed and replaced with Seahorse assay media, which consisted of DMEM supplemented with 4 mM glutamine. OCR was assessed under basal conditions and following sequential injections of 1 mM glucose, oligomycin (2 μM), FCCP (0.5 μM), and a mix of rotenone plus antimycin A (2 μM each). Immediately after the measurements, cells were lysed in RIPA buffer, and the protein concentration was used to normalize the OCR data.

Primary Hepatocyte Cultures and Analysis

Rat primary hepatocytes (Lonza, #RWCP01) were thawed and plated in a 24-well plate at a density of 0.16 million cells per well in Hepatocyte Plating Medium (Lonza, #MP100). Four hours post-plating, the medium was replaced with Hepatocyte Basal Medium (HBM) supplemented with BCM SingleQuots (Lonza, #CC-4182), including Bovine Pituitary Extract (BPE), Insulin, Hydrocortisone, Gentamicin/Amphotericin-B (GA), Transferrin, and human Epidermal Growth Factor (hEGF).

Twenty-four hours after plating, the hepatocytes were transfected with 1 μg of either pT3.GFP or pT3.MPC2Flag-P2AT2A-MPC1HA using Lipofectamine 3000 (Invitrogen, #L3000001). Forty-eight hours post-transfection, cells were lysed in 1X RIPA buffer, and Western blot analysis was performed to confirm the overexpression of MPC1 and MPC2.

Forty-eight hours post-transfection, hepatocytes were fixed in 4% paraformaldehyde in 1X PBS. Following several washes with 0.1% PBT, cells were incubated in 5% BSA and stained with anti-MPC1 antibody overnight at 4°C. MPC1 was detected using a secondary anti-rabbit Alexa Fluor antibody. Cells were also stained with Phalloidin Red. Coverslips were mounted in DAPI-supplemented Vectashield, and images were captured using an LSM 880 confocal microscope. Cell area marked by Phalloidin Red was quantified using differential interference contrast filter and analyzed with FIJI software.

For protein synthesis measurement, hepatocytes were incubated with 20 μg/ml puromycin for 30 minutes. Puromycin-tagged peptides were visualized by immunostaining with rabbit anti-puromycin antibody (1:500) and mouse anti-Flag M2 antibody (1:1000, Sigma, F1800), followed by the appropriate secondary antibodies. Images were captured with the LSM 880 confocal microscope. Puromycin intensity was measured using FIJI, and the percent change in puromycin intensity in Flag-positive cells was compared to Flag-negative cells.

Gluconeogenesis was assessed using lactate and pyruvate as substrates. Forty-eight hours post-transfection, hepatocytes were incubated in low-glucose DMEM with 1 mM sodium pyruvate and 4 mM glutamine without FBS. After 16 hours, hepatocytes were treated with 200 ng glucagon in no-glucose DMEM without FBS for 3 hours. Cells were then cultured in media with 20 mM lactate and 2 mM pyruvate, with and without 10 μM UK5099, for 2 and 4 hours. Glucose levels in the media were measured using the Amplex Red Glucose Assay Kit (Invitrogen, #A22189) according to the manufacturer’s instructions. Glucose production per cell per hour was plotted on a graph. The compartmentalized NADH to NAD⁺ ratio in hepatocytes was quantified 48 hours post-transfection using the Amplite Fluorimetric NAD⁺/NADH Ratio Assay Kit as described previously.

Fly genotypes

Figure 1

Supplementary Figure 1

Supplementary Figure 2

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Funding

American Heart Association (24POST1201210)

Ashish G Toshniwal

National Institutes of Health (K00CA212445)

Alex J. Bott

National Institutes of Health (1K99HL168312-01)

Ahmad A Cluntun

National Institutes of Health (R01CA228346)

Carl Thummel

Howard Hughes Medical Institute and National Institutes of Health (R01CA228346)

Jared Rutter

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank the University of Utah core facilities, especially James Cox, PhD, at the Metabolomics core, Li Ying, PhD, at the Metabolic Phenotyping core, James Marvin, PhD, at the Flow Cytometry Core, Brian Dalley, PhD, at the High-Throughput Genomics Core, and the DNA/Peptide Synthesis Core. We thank members of the Rutter lab for discussion and feedback on the manuscript. This study was supported by 24POST1201210 to AGT; K00CA212445 to AJB; 1K99HL168312-01 To AAC, R01 CA228346 to CT and JR. JR is an Investigator of the Howard Hughes Medical Institute.