All chemicals were from Sigma Aldrich (St. Louis, MO, US) unless otherwise stated. A BCA protein assay (ThermoFisher, Waltham, MA) was used for protein quantification.

Neonatal rat ventricular myocytes (NRVMs) were isolated from the ventricles of 1- to 2-day–old Sprague Dawley rats (Charles River, Wilmington, MA US) by enzymatic digestions as previously described (31). Cells were plated for 16 hr in MEM-Hanks Salts medium with 5% fetal bovine calf serum including 1 g/L of glucose, 0.2X Penicillin G, and 2 ug/mL Vitamin B12. After 16 hr, Cells were then washed with Minimum Essential Medium (MEM)-Hanks Salts medium with L-glutamine (Gibco) containing transferrin (2 ug/mL), bovine serum albumin (0.1%), insulin (2 ug/mL), bromodeoxyuridine (10 ug/mL), HEPES (0.2M), Penicillin G (0.2X) and Vitamin B12 (2 ug/mL) with 0 mM, 5, 10, or 20 mM L-lactate treatments added. Cells were harvested or measurements taken after 48 hr of lactate treatment. All animal protocols are in accordance with Public Health Service Animal Welfare Assurance, ID A3269-01, and approved by the University of Colorado, Denver—Animal Care and Use Committee.

VMs were plated on a gelatin coated Seahorse 96-well plate at a density of 50,000 cells/well. Lactate was added after 24 h and mitochondrial function assessed by the Seahorse after 48 hr of lactate treatment in medium containing 1 mM of pyruvate, 2 mM of glutamate and 10 mM of glucose according to manufacturer suggestions. The Mito Stress Test kit was used with a XFe96 Seahorse Analyzer (Agilent, Santa Clara, CA US). Three separate NRVM preparations were used to collect Seahorse data. After the Seahorse assay, differences in live cell density for each well were normalized by incubating with the dye CyQuant (ThermoFisher, Waltham, MA, US) to image live cells, incubating for an hr and imaging with a fluorescent plate reader (iD5, Molecular Devices) according to manufacturer directions.

Amplex™ UltraRed Reagent (Life technologies) was used to measure superoxide plus hydrogen peroxide in cells. Briefly, NRVMs were plated on black 96-well microplates. Forty-eight hours post lactate treatment, cells were rinsed one time with phosphate-buffered saline (PBS). A solution containing Amplex Red dye (final concentration: 50 microM), hydrogen peroxide (0.0015% final) and superoxide dismutase (SOD, final concentration: 5 units/mL) was added to each well to a final volume of 200 microL and incubated in the dark at 37°C for 30 mins. Sample fluorescence was measured using a fluorescence platereader (iD5, Molecular Devices) at excitation and emission wavelengths of 540 and 600 nm, respectively. The cells were washed twice with 100 microL PBS and cell number was assessed using CyQuant cell proliferation assay kit according to the manufacturer's instructions (ThermoFisher). Sample fluorescence was measured using the same plate reader at excitation and emission wavelengths of 580 and 527 nm, respectively. Each condition was examined in 15 wells per plate in 4 different NRVM preparations leaving the edges free of cells. Data shown is from a single representative experiment.

Carnitine Palmitoyltranferase (CPT) I and CPT II activities were quantified in NRVMs using a ¹⁴C carnitine-based radioactivity assay previously described in detail elsewhere (32). The assay measures CPT I activity by permeabilizing the plasma membrane and measuring the production of palmitoylcarnitine from palmitoyl-CoA. The activity of CPT II was measured by permeabilizing the mitochondrial inner membrane and adding malonyl CoA to inhibit CPT I.

Cardiolipin was quantified in NRVMs using previously published methods with liquid chromatography coupled to electrospray ionization mass spectrometry in an API 4000 mass spectrometer (Sciex, Framingham, MA) (33). After 48 hr of lactate treatment, cells were washed and scraped off plates using PBS. Lipids were extracted according to previously published methods with 1 mmol tetramyristoyl-cardiolipin as an internal standard (Avanti Polar Lipids, Alabaster, AL, US) (33, 34). Cardiolipin species were expressed as a percentage of the following cardiolipin species having a mass/charge (m/z) ratio of 1422, 1424, 1426, 1448. 1450. 1452, 1454, 1456, 1472, 1474, 1476, 1496, 1498, 1500. These percentages were summed for the species having side chains of 70 carbons (m/z 1422–1426), 72 carbons (m/z 1448–1456), 74 carbons (m/z 1472–1476) and 76 carbons (m/z 1496–1500). The fatty acyl side chains of the dominant form of these species are listed in Table 1.

Media for 0, 5, 10, and 20 mM lactate was made and 1 ml saved at −80°C until further processing for measurement of initial extracellular lactate. The remainder of the media was added to three 60 mm plates per condition. After 48 h of incubation with lactate treatments, 1 mL of spent growth media was collected from each of three NRVM plates per condition and frozen at −80°C until further processing for measurement of 48 hr extracellular lactate. Following aspiration of all spent media, cells were washed three times with cold PBS, 300 uL of cold PBS was added to each plate, and NRVMs were scraped and frozen at −80°C until further processing for measurement of intracellular lactate. Upon thawing, cells were kept on ice throughout the entire protocol. Cell suspensions were sonicated for 8 secs at 50% amplitude with a pencil-type sonicator (model 450, Branson, Danbury, CT) and 100 uL of each sample aliquoted into a clean microcentrifuge tube and protein concentrations determined using the Quick Start™ Bradford Colorimetric Protein Assay (#5000201, Bio-Rad, Hercules, CA) as per manufacturer's protocol. The remaining lysed-cell suspensions were spun at maximum speed in a microcentrifuge at 4°C for 15 mins (model PrismR, Labnet International, Edison, NJ). Following centrifugation, supernatants were transferred into new microcentrifuge tubes. Spent and initial growth media and intracellular (lysed-cell supernatant) lactate concentrations were determined via the L-lactate Colorimetric Assay Kit I from Eton Biosciences (#120001400A, San Diego, CA) as per the manufacturer's protocol. Colorimetry as measured via a Synergy 2 plate reader (BioTek, Winooski, VT) was employed to determine protein and lactate concentrations. Intracellular and spent media lactate concentrations were normalized to respective protein concentrations prior to further calculations/data analysis.

All data with the exception of reactive oxygen species measurements and intracellular/extracellular lactate measurements are graphed from multiple preparations on different days of primary cultured NRVMs and therefore for each preparation day, the data were normalized to untreated cells having a value of 1. Data analysis used Prism version 8.0 (GraphPad Software, La Jolla California USA). Treatment effects were analyzed using a 1-way ANOVA with corrections for multiple comparisons; the level of statistical confidence was set at p < 0.05. Data sets were tested for Gaussian distribution with D'Agostino & Pearson omnibus or the Shapiro-Wilk normality test. All data in this study conformed to a Gaussian distribution.

Chronic lactate exposure results in the uptake of L-lactate in NRVMs and a resulting decrease in the media concentration of lactate (Figure 1A). Intracellular lactate in the NRVMs increases with increasing concentration of lactate treatment (Figure 1B). On one hand, NRVMs take lactate added to the media for ATP production. Further, in NRVMs, cytosolic glycolysis and lactate account for ~44 and ~25% respectively for ATP synthesis (27). Fetal bovine calf serum (FBS) contains 1 g/L of glucose needed for NRVMs survival. Due to their high glycolytic and lactate metabolism, it is expected that over time, intracellular lactate is increased. Moreover, it is known that lactate transport kinetics obey a gradient pattern through monocarboxylate transporters (MCT1/4) (35). Therefore, if there is a higher lactate concentration extracellularly compared to the intracellular one, the exportation of intracellular lactate could be compromise leading to intracellular lactate accumulation as could be the case of the 20 mM exposure.

After 48 hr chronic exposure of NRVMs to 0, 5, 10, or 20 mM L-lactate, CPT activities decreased with increasing lactate concentration (Figure 2). There was a trend (p = 0.07) for a decrease of CPT I with 20 mM lactate exposure (Figure 2A). The effect was more pronounced for CPT II where 48 hr of lactate exposure of 10 and 20 mM significantly decreased CPT II activity (p < 0.001 p < 0.01, respectively, Figure 2B).

Chronic lactate exposure alters the cardiolipin molecular species profile in NRVMs in a concentration-related manner. Lactate causes a percentage increase in cardiolipin having a total side chain length of 72 carbons which are comprised of oleate and linoleate side chains p < 0.01 and p < 0.001 for 10 and 20mM respectively) (Figure 3B), but decreases in cardiolipins having either smaller (70 carbon) or longer (74 or 76 carbon side chains) (p < 0.05-p < 0.0001) (Figures 3A,C,D). Interestingly, there is no significant alterations in the ratio of L₄CL (m/z 1448 having four linoleate side chains) to either O₄CL (m/z 1456 having four oleates) or LO₃CL (m/z 1454). This indicates that the side chain composition is not changing within the 72 carbon grouping (data not shown).

Chronic lactate exposure causes an increase in the level of mitochondrial reactive oxygen species (both superoxide and hydrogen peroxide) production in NRVMs (p < 0.5) as shown in Figure 4.

Through Seahorse system analysis, NRVMs with chronic exposure to lactate had decreased oxygen consumption rate (OCR) due to decreased ATP production (p < 0.05) (Figure 5A) and decreased maximal respiration (p < 0.05) (Figure 5B). Representative Seahorse Analyzer trace for one cell preparation day is shown in Figure 5C.

As a model system noted for the cellular oxidation of fatty acids, a major component of metabolic flexibility, we employed a neonatal rat ventricular myocyte (NRVM) model. Beyond autocrine and endocrine roles of lactate on regulation of lipolysis via covalent binding to GPR-81 (HCAR-1) (40), effects of lactatemia on depressing lipid oxidation as well as lipolysis have long been known (37). Furthermore, an impaired ability to clear pyruvate and lactate by oxidation (the enzymes of gluconeogenesis are not expressed in cardiomyocytes) leads to accumulation of malonyl-CoA, which is an inhibitor of CPT1 and mitochondrial fatty acid uptake and oxidation as previously shown by Saddik and colleagues (30). In the present study we show that lactate inhibits the activities of both CPT I and CPT II, especially the latter. Such effects would limit cellular lipid oxidation by downregulating mitochondrial uptake of carnitine-fatty acid derivatives. Those metabolic limitations are involved in the pathogenesis of insulin resistance and T2D (41, 42) and heart failure (43). We believe that our results showing effects of chronic lactate exposure on CPT I and II may explain in whole, or in part the relationship between chronic lactate exposure and metabolic inflexibility in vivo.

In this study, results also show a unique perturbation of cardiolipin, the phospholipid that is highly integrated into the electron transport chain and other enzymes of fatty acid oxidation. These changes involve the enrichment of cardiolipins having oleate (18:1n9) or lineoleate (18:2n6) side chains, and a dramatic decrease in those cardiolipins having either smaller (palmitoleate, 16:1) or larger (arachidonate, 20:4n6 or docosahexaenoate, 22:6n3) fatty acyl side chains. This may be a direct effect of the inhibition of fatty acid transport via CPT since oleate and lineoleate are the most abundant unsaturated fatty acids and may already exist within the mitochondria to be put onto cardiolipin. Alterations to the cardiolipin molecular species profile of this sort have not been described before and further investigation is needed to determine the effect of these cardiolipin changes on mitochondrial function. However, the pattern of size shift in cardiolipin to four 18 carbon side chains is also seen in rat heart tissue during the developmental transition from the neonatal to the adult heart (Genevieve C. Sparagna and Kathryn C. Chatfield unpublished). What we do know from our studies with Barth Syndrome, where the cardiolipin profile is greatly altered, is that CPT function does not appear to be dependent on cardiolipin profile (22), making the cardiolipin alterations in this study not responsible for CPT activity decreases.

Lactate exposure increases mitochondrial ROS generation, which may be a direct effect of the alteration of cardiolipin profile in the electron transport chain. There are multiple studies associating increased ROS with mitochondrial and cellular damage (28, 29, 44) and evidence supporting the involvement of ROS in the pathogenesis of T2D (45).

Lastly, while acute lactate exposure results in tightly coupled mitochondrial lactate oxidation (10, 12), prolonged exposure of NRVMs to lactate negatively affects mitochondrial respiratory control and coupling efficiency as well as the uncoupled maximal respiration rate. The mechanism by which chronic lactate exposure affects mitochondrial respiratory capacity and coupling efficiency may be associated with ROS production (Figure 4) or alterations in the cardiolipin profile (Figure 3). The response of NRVMs to chronic hyper-lactate exposure in culture may serve as a model of what happens in vivo where deficits in mitochondrial respiratory capacity may be behind the pathogenesis of multiple diseases including insulin-resistance, T2D and even cancer as mentioned vide supra. Decreases in mitochondrial function affected by acute and chronic lactate exposure may serve to decrease muscle mitochondrial fatty acid transport and oxidation in vivo resulting in metabolic inflexibility.

Further, decrease in maximal respiration might be attributed to cardiolipins modifications. The work done by Chatfield and Sparagna on mice and human heart show that in Barth Syndrome when cardiolipin is dramatically altered, there is decreased fatty acid utilization, increased ROS and decreased mitochondrial function (20) and (Genevieve C. Sparagna and Kathryn C. Chatfield unpublished). Thus, cardiolipin alterations can also contribute alone to changes in cellular bioenergetics. Moreover, herein we show that lactate plays an important role in regulating cardiolipin configuration.

For nearly a century lactate has been considered as a “waste product” of anaerobic metabolism. However, we now know that lactate is formed under fully aerobic conditions, is a major energy source (1, 46–48), the major gluconeogenic precursor (46), a signaling molecule, a “lactormone” (37, 38) responsible for diverse actions such as gene expression (13, 49), and possibly a master regulator of carcinogenesis (50, 51). In the process of shuttling between sites of production and removal, lactate exerts profound effects on fat and carbohydrate (CHO) metabolism. Lactate is a preferred fuel over glucose in the beating heart (7, 52) working muscle (5, 46) and brain (9). Regarding fat metabolism, it is known that lactate binds G-protein coupled receptor (GPR81) that inhibits lipolysis in adipocytes (40, 53).

Lactate is oxidized in vivo in the mitochondrial reticulum via the mitochondrial lactate oxidation complex (mLOC) comprising a MCT1, its chaperone (CD147/Basigin), a mitochondrial LDH (mLDH), and cyclooxygenase enzyme (COX) (39, 54). It is well known that mitochondrial dysfunction is a hallmark in metabolic diseases like type 2 diabetes and cardiometabolic disease (CMD) characterized by poor fatty acid, carbohydrate oxidation and metabolic inflexibility (55–60). Thus, because lactate is a mitochondrial substrate, lactate oxidation is highly dependent on proper mitochondrial function as is demonstrated in individuals after exercise training (3, 61, 62). As early as 1962, Issekutz and colleagues observed the relationships between lactatemia and fatty acid oxidation (63). Recently, we have shown high inverse correlations (r > 0.9) between blood lactate levels and fatty acid oxidation rates. The inverse correlations hold on a wide range so human subjects ranging from those with metabolic syndrome to elite athletes (64).

Finally, the mechanisms by which chronic lactate exposure could elicit biological responses shown herein are not known and worth exploring in further studies. We believe that while acute, short-term lactate exposure, as occurs in physical exercise, is beneficial to skeletal muscle and other organ systems, chronic lactate exposure associated with lack of clearance could lead to metabolic dysregulation, and probably to disease. For example, Hashimoto et al. in the Brooks Lab showed that acute lactate exposure regulates 630 genes in mononucleated myotubes and striated L6 cells (13). Moreover, after studying MCF-7 breast cancer cells in culture we recently made a case that lactate accumulation from chronic aerobic lactate production (i.e., the Warburg Effect) in the absence of disposal, was an oncometabolite in cultured MCF-7 breast cancer cells (51). Although it may seem a paradox that acute lactate exposure, like in the case of exercise, could have beneficial effects, chronic lactate exposure, like in the case of cancer could be “detrimental.” The mechanisms behind acute vs. chronic lactate exposure must be further explored.

In summary, results of our study add to the current and emerging effects of lactate on acute and long-term regulation of metabolic rate and energy substrate partitioning. While hardly definitive or representative of metabolic regulation in humans and other mammals in vivo, a reasonable interpretation of the present results on cardiomyocytes is that they provide a partial explanation of the effect of lactatemia in downregulating mitochondrial lipid oxidation as might occur in heart failure and CMD (6, 43). On one hand, as reviewed above, lactate can be a preferred energy substrate when lactatemia is acute (37, 65, 66) including acute heart failure (67). However, on the other hand chronic intracellular and extracellular lactate exposure may negatively affect cellular bioenergetics by decreasing mitochondrial function, thus eliciting changes in fatty acid oxidation leading to metabolic inflexibility. Further research is needed to determine the mechanisms involved in decreased lactate clearance capacity that are involved in pathological conditions involving metabolic dysfunction in conditions and diseases associated with lactatemia.