Depletion of mitochondrial inorganic polyphosphate (polyP) in mammalian cells causes metabolic shift from oxidative phosphorylation to glycolysis.

Inorganic polyphosphate (polyP) is a linear polymer composed of up to a few hundred orthophosphates linked together by high-energy phosphoanhydride bonds, identical to those found in ATP. In mammalian mitochondria, polyP has been implicated in multiple processes, including energy metabolism, ion channels function, and the regulation of calcium signaling. However, the specific mechanisms of all these effects of polyP within the organelle remain poorly understood. The central goal of this study was to investigate how mitochondrial polyP participates in the regulation of the mammalian cellular energy metabolism. To accomplish this, we created HEK293 cells depleted of mitochondrial polyP, through the stable expression of the polyP hydrolyzing enzyme (scPPX). We found that these cells have significantly reduced rates of oxidative phosphorylation (OXPHOS), while their rates of glycolysis were elevated. Consistent with this, metabolomics assays confirmed increased levels of metabolites involved in glycolysis in these cells, compared with the wild-type samples. At the same time, key respiratory parameters of the isolated mitochondria were unchanged, suggesting that respiratory chain activity is not affected by the lack of mitochondrial polyP. However, we detected that mitochondria from cells that lack mitochondrial polyP are more fragmented when compared with those from wild-type cells. Based on these results, we propose that mitochondrial polyP plays an important role as a regulator of the metabolic switch between OXPHOS and glycolysis.


INTRODUCTION
Inorganic polyphosphate (polyP) is an ancient molecule that has been well-conserved throughout evolution and is present in all studied organisms (1). It is composed of tens to hundreds of units of orthophosphate linked together by high-energy phosphoanhydride bonds. In mammalian cells and tissues, polyP is a ubiquitous molecule, present in many different intracellular locations, including mitochondria (1). In fact, in these organisms, mitochondria appear to be one of the preferred locations of polyP (1)(2)(3).
PolyP, which bounds are isoenergetic to those found in ATP (4), has been proposed to play a crucial role in cellular energy storage, allowing for the stored energy to be easily available under conditions of cellular stress (5, 6). In fact, the role of this polymer as a key energy metabolite has already been demonstrated in a wide-variety of organisms and models (2,3,(7)(8)(9)(10)(11)(12)(13)(14). Consistent with its high energy feature and its preferred mitochondrial localization, polyP was recently proposed of being produced and hydrolyzed by the mammalian mitochondrial ATP synthase (15). It has been further hypothesized that polyP might regulate the mitochondrial energy metabolism in mammalian cells, either through its direct participation in the ATP production or through the regulation of the rates of key enzymatic activities (6,11). As a result, in mammalian models, amorphous polyP has been shown to have a protective effect when cells experience decreased levels of ATP as a result of the presence of stressors, such as amyloid peptides (12).
PolyP and other polyphosphate compounds have also been proposed to play a role in energy metabolism in other organisms. Specifically, it has been shown that inositol pyrophosphate represses mitochondrial functionality and enhances glycolysis in yeast (16). The authors hypothesized that the ability of inositol pyrophosphate to regulate the cellular energetic metabolism is the result of the interaction between inositol pyrophosphates, ATP, and polyP. Moreover, accumulation of polyP has also found to play a key role in the energy metabolism of Dictyostelium discouideum, which show a 2.5-fold decrease in their ATP levels when the polyphosphate synthesizing enzyme Downloaded from http://portlandpress.com/biochemj/article-pdf/doi/10.1042/BCJ20200975/907303/bcj-2020-0975.pdf by guest on 20 April 2021 Biochemical Journal. This is an Accepted Manuscript. You are encouraged to use the Version of Record that, when published, will replace this version. The most up-to-date-version is available at https://doi.org/10.1042/BCJ20200975 PPK1 is deleted (17). In addition to its potentially direct involvement into cellular energy metabolism, mitochondrial polyP has also been implied in the regulation of the mitochondrial permeability transition pore (mPTP) (18)(19)(20)(21) and in the buffering of mitochondrial calcium (3,7,14). Both mPTP and mitochondrial calcium homeostasis are closely related to the bioenergetic status of the cells.
Dysfunctional cellular bioenergetics is a crucial component in the etiopathology of many diseases and dysfunctions, ranging from diabetes to many neurodegenerative disorders, and cancer (22)(23)(24). Interestingly, polyP has been shown to protect cells against stressors commonly associated with dysfunctional bioenergetics, such as increased oxidative stress or the presence of amyloids (4,12,(25)(26)(27)(28)(29). However, it is unclear whether these effects are a direct consequence of polyP's role in regulating bioenergetics and/or are related to other functions of this multifaceted polymer.
The central aim of the current study was to elucidate the relationship between mitochondrial polyP levels and the status of the cellular energy metabolism in mammalian cells. To do this, we compared ATP levels and mitochondrial bioenergetics parameters in Wild-type (Wt) HEK 293 cells and cells lacking mitochondrial polyP (MitoPPX). Our studies demonstrated that MitoPPX cells have decreased ATP levels, lower rates of OXPHOS and increased levels of glycolysis, compared to Wt cells. Yet mitochondria isolated from the two cell lines showed no significant functional differences. Subsequent morphological and biochemical analysis revealed that cells lacking mitochondrial polyP show significant mitochondrial fragmentation, compatible with increased mitochondrial fission, even if this point should be further investigated.
Based on these data, we propose that mitochondrial polyP is a previously unrecognized regulator of the metabolic balance between OXPHOS and glycolysis.

Reagents.
Dulbecco's Modified Eagle medium (DMEM), penicillin-streptomycin, Hank's Balanced Salt Solution (HBSS), G418, trypsin, and heat-inactivated fetal bovine serum (FBS)  These cells were generated following the protocol published in (7). Briefly, we transfected HEK 293 cells with the plasmid containing the sequence for the mitochondrial expression of the yeast exopolyphosphatase enzyme (PPX), GFP and the resistance to geneticin. We selected the cells positive for the expression of this construct by treating our samples with geneticin.

Seahorse assays.
Cells grown to the confluence of 80-90% in 75 cm 2 flasks were trypsinized and centrifuged. Thereafter, the supernatant was discarded, and the pellet was re- Once the Seahorse measurements were concluded, the medium was carefully removed and cells were incubated with 5 μM Calcein-AM in PBS, in a regular incubator, for 30 min. Thereafter, fluorescence and cell number were measured, using a Cell Insight CX7 LZR reader, with a 4X magnification objective lens (ThermoFisher Scientific, Waltham, Massachusetts, US). The data obtained from the Seahorse measurements were normalized with the number of cells calculated using this technology.

Image acquisition (Seahorse experiments).
Before and after conducting the Seahorse experiments, cells were imaged using an EVOS FL Auto Cell Imaging System, with a 20x magnification objective lens (ThermoFisher Scientific, Waltham, Massachusetts, US). Specifically, the cells on the Seahorse plates were placed on the microscope just before the hypoxia incubation and just after the Seahorse assay. Images were taken from cells from the exact same wells before and after the experiments.

Fluorescence and confocal microscopy
Cells were plated on 25-mm borosilicate poly-L-lysine-coated glass coverslips at a 50% confluence. 24 h later, cells were washed twice using sterile HBSS and mounted on Sykes-Moore microscopy chambers (BellCo, Vineland, New Jersey, US). Sterile HBSS was added to the glasses on the chambers and cells were imaged on a Nikon Fluorescence microscope, using a 20x magnification objective lens (Chiyoda, Tokyo, Japan); or on a Leica SP8 Confocal microscope, using a 40x oil-immersion magnification objective lens (Wetzlar, Germany).

ATP luminescence measurement.
ATP was measured per the instructions that were provided by the manufacturer. Briefly, 10,000 cells per well were plated on a flat bottom, white, 96-well plate. We prepared Subsequently, the substrate solution, also provided in the kit, was added and the cells were incubated under the same conditions as in the previous incubation. Cells were then incubated for another 10 min in the dark, in order to dark-adapt the plate. Lastly, luminescence was measured, at 500 ms, from the bottom.

Mitochondrial length measurement.
Mitochondria from Wt and MitoPPX cells were loaded with TMRM, following the same protocol published in (30). Subsequently, cells were mounted on microscopy chambers and images were acquired through the protocol previously described above.
Mitochondrial length was then measured, using ImageJ software (National Institutes of Health, Bethesda, Maryland, US) and following the protocol published in (31).

Western Blotting.
Western Blotting assays were conducted as previously described in (32,33) through the use of the specific antibodies for the proteins of interest. Briefly, cells were scraped on ice and lysed. Protein was assayed using the Pierce BCA Protein Assay kit. Thereafter, equal amounts of protein were loaded on 12% polyacrylamide gels and electrophoresis was conducted at 100 V. The proteins were then transferred from the gel to a PVDF membrane, blocked and incubated with a 1:1000 dilution of the specific antibody overnight. The day after, the membrane was washed three times with TBS-Tween, incubated with the specific secondary antibodies, washed three times and the signal was visualized using Pierce ECL Western Blotting Substrate. Subsequently, membranes were stripped, blocked and Western Blotting for actin (loading control) was conducted, following the same protocol.

Mitochondrial isolation and high resolution respirometry.
Wt and MitoPPX cells were grown until 90% confluence, after which they were trypsinized and spun down. The cellular pellet was then resuspended in 25 mL of isolation buffer, containing 200 mM mannitol, 50 mM sucrose, 5 mM dipotassium phosphate, and 0.1% v/v BSA, at pH=7.15. Homogenization was done using an Omni International Handled homogenizer (Kennesaw, Georgia, US) at 18.000 rpm for 20 sec.
Mitochondria were then recovered by gradient centrifugation in isolation buffer, at 4 ⁰C.
Mitochondrial protein was quantified with the Pierce BCA Protein Assay kit and BSA standards, using an Olis DM-245 spectrofluorometer (Bogart, Georgia, US). The mitochondrial stock solution was then diluted to a working concentration of 10 mg/mL.
Oxygen consumption data was obtained at 37 ⁰C, using Oxygraph 2k (Oroboros Instruments, Graz, Austria). Each respiratory chamber was loaded with respiration

PolyP degradation measurement.
Wt and MitoPPX cells were grown under regular conditions on Petri Dishes. When the confluence was around 90%, they were washed three times in ice-cold PBS, scraped on ice using the same PBS and collected into a 1 mL tube. Cellular suspensions were then centrifuged at 1,000 rpm for 5 min, at 4 ⁰C. Thereafter, supernatants were discarded and the pellets, containing the cells were re-suspended in 500 µL of lysis buffer. The composition of this buffer and the protocol for the assay were modified from (2).
Specifically, our solution contained 20 mM TRIS-HCl, 10% glycerol, 1% Triton X100 and 1 mM PMSF. Cellular suspensions were then sonicated four times for 25 sec, at 100% amplitude, with 45 sec of cooling on ice between sonication cycles. BCA Protein Assay was conducted on the samples and appropriate amounts of the buffer were added to the samples, in order to have the same concentration of protein in Wt and MitoPPX cells. Subsequently, the incubation buffer, which contained 0.1% BSA (w/v), 10% glycerol, 20 mM TRIS-HCl (pH=7.5) and 50 mM potassium chloride, was prepared and kept in an ice bucket. Samples were diluted 1:1 in this buffer.
A solution containing 40 µM DAPI was prepared in incubation buffer and kept in the dark. DAPI has been proven to be a useful probe to analyze the presence of polyP in biological samples (34). Medium chain polyP was diluted to 0.5 M phosphate (51 mg/mL

Statistical analysis of the experimental data.
Statistical significance of differences between groups was determined by the Student's test or the two-tailed Student's test. The level of statistical significance was set at α = 0.05 (* p ≤ 0.05, **p ≤ 0.01, ***p ≤ 0.001). For the statistical analysis and graphical representation, Origins Lab software (Northampton, Massachusetts, US) was used.

MitoPPX cells show decreased levels of oxidative phosphorylation
To assess the metabolic consequences of mitochondrial polyP depletion, we first compared OXPHOS-linked ATP production in Wt and MitoPPX cells. We used HEK 293 cells for these studies since their glucose metabolism has been extensively characterized (36). Specifically, it has been shown that 15% of the consumed glucose is channeled through the pentose phosphate pathway (PPP) whereas 77% of metabolized glucose enters glycolysis. About 22% of the pyruvate generated in this process is then metabolized into acetyl-CoA for energy production and lipid biosynthesis (36).
These cells express GFP-labeled yeast exopolyphosphatase (PPX) selectively in the  Supplementary Fig. 1B). Based on these results, we conclude that MitoPPX cells express functional yeast PPX specifically in the mitochondria, leading to decreased levels of mitochondrial polyP.
To assess the metabolic consequences of mitochondrial polyP depletion, we monitored ATP production in Wt and MitoPPX cells, using the Seahorse technology (37,38).
Specifically, we employed the MitoStress Kit to assay the Oxygen Consumption Rate (OCR) in both cell lines. This kit includes the following components: oligomycin, which serves as an inhibitor of the ATP synthase; FCCP, an uncoupling agent that collapses the proton gradient; and rot/AA, which inhibits Complex I and III of the Electron Transfer Chain (ETC) and hence shuts down mitochondrial respiration. We focused our studies initially on select parameters of mitochondrial respiration. Specifically, we measured ATP-linked respiration by determining the difference between basal respiration (the basal signal) and the signal after addition of oligomycin, as well as the spare capacity by determining the difference between basal respiration and maximal respiration upon addition of FCCP.
The observed oligomycin response in Wt and MitoPPX cells indicated that ATP production was dramatically reduced in MitoPPX cells, compared to Wt cells (Fig. 1A-B). Furthermore, and in striking contrast to Wt cells, MitoPPX cells exhibited a significantly altered response to FCCP addition. In Wt cells, addition of FCCP elicited the expected stimulation of mitochondrial respiration, with a maximal stimulation being achieved at 0.5 μM of FCCP. In stark contrast, addition of the same concentration of FCCP to MitoPPX cells reduced respiration from approximately 200 pmolO2/min/cells to 50 pmol O2/min/cells. (Fig. 1A-B). Lower concentrations (i.e. 0.25 µM) of FCCP were still able to stimulate respiration in MitoPPX cells, indicating that the respiratory chain was generally functional in MitoPPX cells (Fig. 1C). However, MitoPPX cells appear to be more susceptible to the membrane depolarization that is triggered by higher concentrations of FCCP (Supplementary Fig. 3). Microscopy images that were taken before and after the Seahorse experiments indicated that similar number of cells were present under both conditions and that no major cell detachment occurred during the treatment (Supplementary Fig. 4A). These results confirmed that the observed differences were not caused by differences in cell number.

MitoPPX cells show increased rates of glycolysis
The decrease in ATP generation can be either an indication of the overall slower energy metabolism or of a shift of ATP production towards the less effective glycolysis. To explore these possibilities, we measured glycolytic rates in Wt and MitoPPX cells utilizing the Glycolytic Rate Assay kit. This assay measures the protons extruded into the extracellular media as a consequence of the breakdown of glucose into lactate during glycolysis (ECAR, Extracellular Acidification Rate). Since the Krebs cycle also acidifies the media through the generation of CO2, we inhibited oxygen consumption during the assay using rot/AA. By using this approach, the rate of proton efflux from respiration can be calculated and subtracted from the total proton efflux rate, yielding in the glycolytic proton efflux rate (glycoPER). As control, we administered 2-deoxy-Dglucose (2-DG), which serves as a glucose analog that competitively inhibits glycolysis (39).
As shown in Fig. 2A-B, MitoPPX cells showed a 1.5-fold increase in ECAR and an almost 2-fold increase in glycoPER compared to Wt cells. Inhibition of mitochondrial OXPHOS by the addition of rot/AA revealed the same maximal rates of glycolysis, and addition of 2-DG triggered the same abrupt signal decrease in both Wt and MItoPPX cells (Fig. 2A). These experiments suggest that cells lacking mitochondrial polyP undergo a metabolic shift from OXPHOS to glycolysis. As before, we confirmed that both Wt and MitoPPX samples show a similar cell density (Supplementary Fig. 4B). Notably, increased glycolysis was not able to completely compensate for the loss of aerobic ATP production. Indeed, measurements of total endogenous levels of ATP using a luminescence assay revealed decreased levels of ATP in the MitoPPX cells, compared with the Wt cells (Fig. 2C).

Metabolomics data indicate changes in glycolysis, and energy and nucleotide metabolism
To independently assess the functional consequences of mitochondrial polyP depletion on cellular bioenergetics, we conducted metabolomic analysis in our Wt and MitoPPX samples. We used a LC-MS/MS-based metabolomics approach targeting a list of 310 metabolites, many of them related to cellular bioenergetics. To conduct these experiments, we prepared six biological duplicates of each cell type (Wt and MitoPPX).
Out of the 310 metabolites targeted, we consistently detected 175 in our cell samples.
Of these, 45 had p-values < 0.05, and 15 remained significant after FDR correction using a Benjamini-Hochberg approach with q < 0.05 (see Supplementary Table 1).
The metabolomics analysis showed a large and highly significant reduction (>16 fold) in sugar monophosphates in MitoPPX cells, compared with the results in the Wt samples (Table 1). Moreover, the ratio (glucose/lactate) in MitoPPX cells did not change significantly, compared with the same ratio in the Wt samples (p=0.023, Supplementary Table 1). Furthermore, we observed significant differences in the abundance of the energy-related coenzymes, NADP and NADPH, as well as of ADP and AMP. All of these compounds were significantly increased in MitoPPX cells, when compared to Wt samples. In fact, AMP was one of the most highly altered metabolites in MitoPPX cells compared to Wt samples. The levels of two other energy-related metabolites, succinylcarnitine and isovalerylcarnitine, were also significantly increased in MitoPPX cells compared to Wt cells. In addition to these changes, we found that MitoPPX cells our Seahorse measurements, and were indicative of increased rates of glycolysis and glycolysis-specific metabolites in MitoPPX cells compared to Wt cells (Fig. 3). Notably, as stated, in addition to changes in glycolysis, our data also indicate alterations in purine, pyrimidine, glutathione and sugar metabolism in cells lacking mitochondrial polyP.

Isolated MitoPPX and Wt mitochondria have similar activity
The shift in energy metabolism towards glycolysis that we observed in MitoPPX cells might be caused by a loss in mitochondrial function. To test this possibility, we measured the rates of respiration in isolated mitochondria, using resolution respirometry. This technique, which is based on the amperometric measurement of O2, allows us to assay OCR in isolated mitochondria, providing minimal O2 leak and maximum sensitivity to O2 Hutter,[40][41][42].
To our surprise, we found that Wt and MitoPPX cells behave very similarly in this assay. Figures 4A-B, the leak respiration and ADP-coupled respiration rates from Wt and MitoPPX cells were not significantly different. Since ADP-coupled respiration, which drives OXPHOS of ADP to ATP, is mediated by proton pumps across the inner mitochondrial membrane (41), this result indicates that both mitochondrial inner membrane and the ETC are fully functional in MitoPPX cells. The same is true for leak respiration, which is defined as the electron flow, coupled to proton pumping to compensate for proton leak (41). Accordingly, when we calculated the Respiratory Control Ratios for Wt and MitoPPX isolated mitochondria, we did not find significant differences between the two cell types. Analysis of the steady state levels of selected protein components of the ETC complexes did not reveal any significant differences between Wt and MitoPPX cells, consistent with our results showing that mitochondria from MitoPPX cells are fully functional ex vivo (Fig. 4C). These results suggest however that the differences between Wt and MitoPPX cells that we observe in regards to the in vivo ATP production are more likely due to changes in the regulation of the energy metabolic pathways than due to structural or functional defects in the ETC. We cannot eliminate the possibility that mitochondria isolated from MitoPPX cells will also present Downloaded from http://portlandpress.com/biochemj/article-pdf/doi/10.1042/BCJ20200975/907303/bcj-2020-0975.pdf by guest on 20 April 2021 increased rates of ATP hydrolysis. This could be also affecting the levels of mitochondrial respiration in our samples.

MitoPPX cells show increased fragmentation
It has recently been proposed that increased mitochondrial fission might trigger the metabolic shift from OXPHOS to glycolysis (43). Increased mitochondrial fission, which has been broadly reported in different pathologies, leads to higher levels of mitochondrial mass but with a defective ability to produce ATP. However, mitochondrial fission is also a physiological process, naturally present in cells and crucial to maintain the physiology or the organelle (44).
To investigate the effect of the depletion of mitochondrial polyP in the fission of the organelle, we measured mitochondrial length in Wt and MitoPPX cells by microscopy.
Our analysis revealed that mitochondria of MitoPPX cells are significantly shorter compared to mitochondria of Wt cells (Fig. 5A-B). Shorter and more abundant mitochondria have been widely associated with the over-activation of mitochondrial fission (45). To further assess the state of mitochondrial fission in our cells, we assayed the levels of Drp1, which is the main protein involved in mitochondrial fission in mammalian cells (46) Consistent with our microscopy data, we found that MitoPPX cells contain substantially increased levels of Drp1 (Fig. 5C). However, when we addressed the levels of phosphorylated Drp1 (serine 616), which is a clear marker of Drp1dependent mitochondrial fission (47), our data showed similar levels of the presence of this protein in both Wt and MitoPPX samples (Supplementary Fig.5B). Accordingly, no increased levels of Drp1 were found on mitochondria isolated from MitoPPX cells, when compared with mitochondria isolated from the Wt samples ( Supplementary Fig. 5A).
These results suggest that in our model mitochondrial fission might occur by a mechanism not directly linked to the Drp1 pathway. Further, no differences were observed in the maximal respiration rate between our Wt and MitoPPX mitochondria. Importantly, we did not detect any differences in the activation of the mitochondrialspecific authophagy (mitophagy) or the intrinsic pathway of apoptosis, two processes that can act downstream of Drp1-mediated mitochondrial fission. This result became evident when we assayed for Parkin, one of the main proteins involved in mitophagy (51) or Bax, a key regulator of the intrinsic pathway of apoptosis (52) (Fig. 5C). Our data suggest that the direct effect of polyP in Drp1 does not activate other pathways within the organelle. These data could further support that the activation of mitochondrial fission in our samples, even if present, is still within the physiological range, as well as that the mechanism involved on this activation might not be the Drp1-mediated classical pathway.

DISCUSSION
Here we present a comprehensive study of the effects of mitochondrial polyP depletion on the energy metabolism of mammalian cells. One of the key findings of our study is the observation that lack of mitochondrial polyP decreases OXPHOS, and concomitantly leads to an increase in glycolysis. Since mitochondria isolated from either Wt or MitoPPX cells showed very similar rates of OXPHOS and both cell types showed comparable expression levels of the key enzymes of the respiratory chain, we propose that mitochondrial polyP is not required for the process of OXPHOS per se, but it might play a regulatory role in energy metabolism.
The metabolomics analysis that we conducted shows a major upregulation in the production of a number of nucleotides, including several energy coenzymes in cells lacking mitochondrial polyP. This increase could be a consequence of the PPX activity.
Moreover, we found that the lactate to glucose ratio increases in MitoPPX cells, consistent with the increased glycolytic conversion of glucose to lactate. It remains to be investigated how lack of mitochondrial polyP triggers the observed changes in other It is possible that the observed metabolic shift from OXPHOS to glycolysis in MitoPPX cells is linked to the levels of mitochondrial fission, through a mechanism independent of Drp1. However, further studies will have to be conducted to unequivocally confirm the mechanistic relationship between these processes. The link between increased mitochondrial fission and a shift from OXPHOS to glycolysis has been demonstrated before in cancer models (43,53). However, in our experiments the morphological changes in mitochondria, which are compatible with increase fission, did not appear to be linked to the classical Drp1-mediated mitochondrial fission pathway. This suggests that other mechanisms might be in play. Based on these results, we now consider the intriguing possibility that mitochondrial polyP serves as a regulator of the mitochondrial fission machinery, maybe through a Drp1-independent mechanism. It is of note that the AMP-activated Protein Kinase (AMPK) enzyme is regulated by the inositol polyphosphate multikinase (IPMK), which is involved in the catalysis of inositol polyP and inositol pyrophosphate (54)(55)(56). Moreover, AMPK has been described as an energy sensor, involved in cellular bioenergetics, regulating mitochondrial fission, in response to bioenergetics stress (57, 58). Thus, decreased levels of polyP might result in decreased activity of IPMK, dysregulating AMPK and, hence stimulating mitochondria fission.
In summary, we report that enzymatic depletion of the mitochondrial polyP leads to the metabolic shift from OXPHOS to glycolysis which is linked to the increased mitochondrial fragmentation. Our data suggest that polyP is a previously unrecognized regulator of the cellular energy metabolism in mammalian cells.

ACKNOWLEDGMENTS
We kindly thank the Northwest Metabolomics Research Center, at University of Washington, for the metabolomics analysis. Moreover, we thank the High Throughput Biology Laboratory, at New York University-Langone Health, especially Sabina Cook, Rebecca Lee and Chi Yu,n for their help with planning, preparing and conducting the Seahorse experiments, as well as to Kevin Bittman for his input on interpreting the obtained data on those experiments. We also thank Dr. Toshikazu Shiba, from Kitasato University, Tokyo, Japan for kindly provide us with medium chain PolyP. Lastly, we thank Mr. Mitch Maleki, Esq., for editing this manuscript. This study was supported by the National Institutes of Health (1K99AG055701-01A1 and 4R00AG055701-03 to MES, GM115570-01A1 to EVP, 1S10OD021562 to DR, and GM122506 to UJ) and by the American Heart Association, Transformation Project Award (18TPA34230060 to EVP).

DECLARATION OF INTEREST
The authors declare no competing interests.

DATA AVAILABILITY STATEMENT
Further information and requests for resources and reagents should be directed to and will be fulfilled by the Lead Contact, Maria E. Solesio Torregrosa (m.solesio@rutgers.edu).