Out of 10 steps in glycolysis, only one reaction- Glyceraldehyde 3-phosphate (G3P) to 1,3-bisphosphoglycerate (PGP), uses NAD+ and thereby producing NADH. Furthermore, this very step is solely responsible for net gain of 2ATP in glycolysis since here phosphorylation occurs without the expenditure of ATP.
Why is NAD+ used only in this step? Is it possible to predict by other information that this is the only step to require NAD+? Is it possible to make this reaction happen without the involvement of NAD(P)+? Is it possible to skip this step and directly produce 3-phosphoglycerate (Lets not worry even if we cannot produce ATP in this process). I am interested in this because, this step necessitates the ferementation for regeneration of NAD+ in anaerobic condition. E. coli produced acetate as a side effect, as expected during anaerobic situations, but also rather unexpectedly during aerobic and high-growth rate situation (acetate-swtich, overflow problem). Production of acetate is unwanted in many industrial applications.
NAD+ is important in this step, since it is co-factor for the glyceraldehyde-3-phosphate dehydrogenase (G3PDH), which acts as a acceptor for the hydrogen atom from the C1 (see below).
If you look at the reaction, the aldehyde from the C1 is oxidized to a carboxylic acid which in a second step is turned into a phosphoester. To do so, a cysteine from the active center of the G3PDH attacks the G3P and forms a hemiacetal. This is oxidized into a thioester, the hydride which is split off in this step is transfered to the NAD+ bound by the G3PDH. In the second step the thioester formed between the enzyme and the G3P is attacked by a molecule of inorganic phosphate to form 1,3-BPG. This step is only made possible by splitting the very energy rich thioester bond. So for this reaction to happen you need both steps, as the formation of an energy rich phosphate ester bond would otherwise not happen. See here for more details.
To go directly from G3P to 3-PG you would need an enzyme which oxidizes the aldehyde group in the G3P. And then you need again an acceptor for the hydrogen which is split off here.
- Although four ATP molecules are produced in the second half, the net gain of glycolysis is only two ATP because two ATP molecules are used in the first half of glycolysis.
- Enzymes that catalyze the reactions that produce ATP are rate-limiting steps of glycolysis and must be present in sufficient quantities for glycolysis to complete the production of four ATP, two NADH, and two pyruvate molecules for each glucose molecule that enters the pathway.
- Red blood cells require glycolysis as their sole source of ATP in order to survive, because they do not have mitochondria.
- Cancer cells and stem cells also use glycolysis as the main source of ATP (process known as aerobic glycolysis, or Warburg effect).
- pyruvate: any salt or ester of pyruvic acid the end product of glycolysis before entering the TCA cycle
NAD + metabolism and its roles in cellular processes during ageing
Nicotinamide adenine dinucleotide (NAD + ) is a coenzyme for redox reactions, making it central to energy metabolism. NAD + is also an essential cofactor for non-redox NAD + -dependent enzymes, including sirtuins, CD38 and poly(ADP-ribose) polymerases. NAD + can directly and indirectly influence many key cellular functions, including metabolic pathways, DNA repair, chromatin remodelling, cellular senescence and immune cell function. These cellular processes and functions are critical for maintaining tissue and metabolic homeostasis and for healthy ageing. Remarkably, ageing is accompanied by a gradual decline in tissue and cellular NAD + levels in multiple model organisms, including rodents and humans. This decline in NAD + levels is linked causally to numerous ageing-associated diseases, including cognitive decline, cancer, metabolic disease, sarcopenia and frailty. Many of these ageing-associated diseases can be slowed down and even reversed by restoring NAD + levels. Therefore, targeting NAD + metabolism has emerged as a potential therapeutic approach to ameliorate ageing-related disease, and extend the human healthspan and lifespan. However, much remains to be learnt about how NAD + influences human health and ageing biology. This includes a deeper understanding of the molecular mechanisms that regulate NAD + levels, how to effectively restore NAD + levels during ageing, whether doing so is safe and whether NAD + repletion will have beneficial effects in ageing humans.
Carbon monoxide was identified to inhibit platelet aggregation, but the mechanism involved has not been defined.
We observed that antiplatelet effect of carbon monoxide is accompanied by inhibition of mitochondrial respiration and inhibition of glycolysis at the level of GAPDH (glyceraldehyde 3-phosphate dehydrogenase).
In the presence of exogenous pyruvate inhibitory effects of carbon monoxide on platelet aggregation and glycolysis were lost, but restored after inhibition of cytosolic NAD + regeneration, suggesting a key role of NAD + depletion in the observed effects.
Antiplatelet effect of carbon monoxide is mediated by inhibition of both ATP-generation processes, mitochondrial respiration at the level of cytochrome c oxidase, and glycolysis, attributed to cytosolic NAD + depletion
See accompanying editorial on page 2344
Carbon monoxide (CO) delivered to an organism through inhalation is lethally toxic. Hence, for a long time, CO has been referred to as a silent killer. However, CO produced endogenously during haem degradation by haem oxygenase enzymes is an important endogenous signalling molecule implied in cellular responses to various stimuli. 1 In turn, CO supplied in relatively low quantities by CO-releasing molecules (CORMs) affords cytoprotective and anti-inflammatory effects. 1,2
Numerous reports provide evidence that CO, due to its antiplatelet and antithrombotic action, plays an important role in maintaining vascular homeostasis in vivo. For example, it was demonstrated that HO-1 (haem oxygenase 1)-derived CO protects against hepatic ischemia-reperfusion injury through the inhibition of platelet adhesion to the sinusoids. 3 Moreover, HO-1-deficiency induces an acceleration of arterial thrombosis, which occurs through various mechanisms (including platelet activation), while CO inhalation 4 or CO delivery by CORM-2 5 can rescue from prothrombotic phenotype of HO-1 deficiency. These reports confirm the importance of HO-1-derived CO in the regulation of vascular thromboresistance and suggest that CORMs may represent a novel class of antiplatelet agents. Indeed, CORM-A1, a prototypic CORM slowly releasing CO, 6 was shown to afford antiplatelet and antithrombotic activities in vivo without any hypotensive effect, while CORM-3, which releases CO instantly, displayed both antithrombotic and hypotensive effects. 7 Recently, various novel CORMs with tuneable CO-releasing properties have been synthetized, all of which were shown to display antiplatelet activity similar or even slightly more pronounced than CORM-A1, 8 confirming the significant antiplatelet effects of CORMs.
Despite the firm evidence on the antiplatelet effects of CO in in vivo and in vitro studies, the mechanisms involved have not been elucidated so far. Even though high concentrations of gaseous CO act in platelets via sGC (soluble guanyl cyclase), 9,10 the antiplatelet effect of CORM-3, in contrast to NO-donors, was not mediated by the activation of sGC. 11,12 Recently, the involvement of glycoprotein-mediated HS1 phosphorylation was suggested to be involved in CORM-2-induced suppression of platelet activation by LPS (lipopolysaccharide). 13 However, this mechanism does not explain the antiplatelet effect of CO reported for platelet aggregation induced by classical platelet agonists such as collagen or thrombin, 11,12 which entail a mechanistically distinct intraplatelet pathways as compared with LPS-induced platelet activation. 14
Platelet aggregation is an energy-demanding process. 15,16 In resting or activated states, platelets draw energy from oxidative phosphorylation and glycolysis. 17–19 However, besides a couple of works describing altered bioenergetics of platelets derived from patients with sickle cell disease, 20 asthma, 21 sepsis, 22 Parkinson disease 23 or diabetes mellitus, 24 still little is known about the role of metabolism in the regulation of platelets’ function. A number of reports, including ours, have demonstrated that CO modulates cellular bioenergetics in various cells types, 25–30 including endothelial cells. 31–33
Given the fact that platelet aggregation is an energy-demanding process, we hypothesized that CO may modulate platelet activity through the modulation of platelet bioenergetics involving oxidative phosphorylation and glycolysis. In the present work, we have characterized the bioenergetics of platelets in resting and activated states, and analyzed the effects of CORM-A1, on mitochondrial respiration, glycolysis, and metabolome in platelets. We have chosen CORM-A1 because of its well documented antiplatelet action in in vitro and in vivo models. 7,12 Here, we propose that the mechanism of antiplatelet effect of CO involves the inhibition of 2 major ATP-generating pathways in platelets—mitochondrial respiration and glycolysis, by the inhibition of cytochrome c oxidase and cytosolic nicotinamide adenine dinucleotide (NAD + ) depletion, respectively.
The authors declare that all supporting data are available within the article in the Data Supplement.
Isolation of Human Platelets
Venous blood was obtained from male volunteers at the University Hospital Blood Bank Centre. Volunteer donors had not taken any medicines for the preceding 2 weeks. Informed consent was given by a volunteer before the blood withdrawal and study conformed to the principles outlined in the World Medical Association Declaration of Helsinki. Blood obtained from at least 3 donors per one independent experiment was collected into vials containing sodium citrate (3.2%, 9:1v/v) as an anticoagulant agent. Blood was centrifuged (260g, 15 minutes) followed by a centrifugation/washing cycle using prostacyclin-containing PBS (PBS containing albumin [1 g/L] and glucose [1 g/L]), according to the previously described method. 34 Washed platelets (WP) were finally suspended in assay medium (Seahorse XF Base Medium Minimal DMEM supplemented with glucose [1 g/L] and glutamine [2 mmol/L], pH 7.4) at a density of 2×10 5 platelets/µL, unless otherwise stated. Contamination of neutrophils in WP was <1/10 6 platelets.
Platelet Aggregation Assay in Humans
Aggregation of blood platelets was assessed in WP using a dual channel aggregometer (CHRONO-LOG) according to the method previously described by Born. 35 WP (500 µL) were equilibrated for 2 minutes at 37°C with a continuous stirring at 800 rpm and then stimulated with collagen or thrombin to cause aggregation. At the beginning of each experiment, concentrations of thrombin (in the range of 0.1–0.5 U/mL) that induced sub-maximum aggregation response were determined. CORM-A1, metabolic inhibitors, and other tested compounds were added 2 minutes before stimulation of platelets with collagen or thrombin. Transmittance was read within 6 minutes after stimulation of platelets with an agonist. The concentrations of CO released from CORM-A1 were measured with myoglobin assay (Figure I in the Data Supplement), as described previously 36 with minor changes (Materials in the Data Supplement).
Analysis of Cellular Bioenergetics Using Extracellular Flux Technology
To measure mitochondrial function and glycolysis in isolated human platelets, a Seahorse XFe96 Analyzer was used (the details are in Materials in the Data Supplement). Briefly, freshly isolated platelets suspended in assay medium were introduced into the Seahorse XFe96-well plates (10×10 6 of platelets per well) followed by centrifugation (5 minutes, 700g) and incubation with bicarbonate-free low buffered assay medium (1 hour, 37°C) in air without CO2 before the beginning of the assay. Changes in oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were assessed over time by sequential injections of reagents in ports A, B, C, and D. Concentrations of oligomycin, FCCP (carbonyl cyanide 4-[trifluoromethoxy]phenylhydrazone), rotenone, and antimycin A were optimized (data not shown). Mitochondrial stress test enabled determination of the following key parameters of mitochondrial function: acute response, proton leak, maximal respiration, spare respiratory capacity, ATP production, and nonmitochondrial oxygen consumption (calculated as described in Materials in the Data Supplement). Additionally, we calculated acute response and oligomycin-induced changes in ECAR. We analyzed the effects of CORM-A1 on thrombin-induced changes in OCR (Δ [Thr-A1]mt. respiration) and in ECAR (Δ [Thr-A1]glycolysis), calculated based on the differences in OCR and ECAR before and after thrombin injection. Based on the experiments with oligomycin, we analyzed the ability of platelets pretreated with CORM-A1 and thrombin to further maximize their glycolysis in response to oligomycin, reflecting a spare glycolytic capacity (Δ [O-Thr]), while based on the experiments with FCCP/pyruvate, we analyzed the ability of platelets pretreated with CORM-A1 and thrombin to further maximize their mitochondrial respiration in response to FCCP, reflecting a spare respiratory capacity (Δ [FCCP-Thr]).
Measurements of lactate were performed by the enzymatic photometric methods using an automatic Pentra 400 (Horiba, Japan) biochemical analyzer according to the manufacturer’s instructions. Samples were suspended in assay buffer at a density of 3×10 4 platelets/μL and treated with tested compounds for 8 minutes at 37°C or preincubated with tested compounds for 2 minutes and then activated with thrombin for 6 minutes at 37°C, followed by centrifugation (1000g, 10 minutes) and measurement of the concentration of lactate in supernatant.
Analysis of Intraplatelet Metabolites
Detection of intraplatelet metabolites was performed according to the protocol described previously, 33,37 with minor changes. Briefly, WP (suspended in PBS containing 1 g/L glucose and 2 mmol/L glutamine, in a number of 500×10 6 per 0.5 mL per sample) were untreated or preincubated for 2 minutes with 300 μmol/L CORM-A1, followed by the addition of thrombin (0.1 U/mL) and further incubation for 6 minutes. The metabolism was then quenched by the addition of 0.5 mL of extraction solution (acetonitrile: methanol: water, 5:2:3, v/v/v) cooled to −80°C. Metabolites were extracted by sonication for 15 minutes on ice, centrifuged (15 000g, 15 minutes, 4°C) and lyophilized. Before analysis, the samples were reconstituted in water, injected into a liquid chromatography column, and analyzed on a QTRAP 5500 (Sciex, Framingham, MA) coupled with UFLC (ultra-fast liquid chromatography) Nexera (Shimadzu, Kyoto, Japan). Chromatography separation was achieved on an Acquity UPLC BEH C18 1.7 μm 3.0×100 mm analytical column (Waters, Milford, MA). Samples were analyzed twice in positive and negative ionization MRM (multiple reaction monitoring) mode. For the analysis in positive ionization using acetonitrile, 100 mmol/L ammonium formate (pH 5.0) 95:5 v/v and 5 mmol/L ammonium formate (pH 5.0) were used as a mobile phase using gradient elution. The total run time was 8 and 10 minutes for negative and positive ionization modes, respectively.
Analysis of GAPDH and PFK-1 Activities
GAPDH (glyceraldehyde 3-phosphate dehydrogenase) activity was analyzed in samples of WP (suspended in PBS containing 1 g/L glucose and 2 mmol/L glutamine, in a number of 300×10 6 per 0.5 mL per sample) untreated or incubated for 8 minutes with CORM-A1, processed as described by Schmidt and Dringen 38 and measured according to the protocol described by Bisswanger. 39 PFK-1 (phosphofructokinase 1) activity was analyzed followed by incubation of WP (500×10 6 per 0.5 mL per sample) with CORM-A1 for 8 minutes or 30 minutes, according to the protocol described previously. 40,41 Details are described in Materials in the Data Supplement.
Analysis of Metabolic ATP Concentration
Measurements were performed in samples treated for 8 minutes with tested compounds, in the presence of apyrase (1 U/mL) using ATPlite 1step Luminescence Assay System (PerkinElmer).
Statistical analysis was performed using ORIGINPRO 9.1 software (OriginLab Corporation, Northampton, MA) or Graphpad Prism software (GraphPad Software, La Jolla, CA). Results were expressed as means±SD. For statistical analysis, the data have been analyzed for normality, and 1-way ANOVA test was performed with Benferroni test with P values being provided in the legends (*P<0.05, # P<0.01, $ P<0.001, and & P<0.0001).
Effects of CORM-A1 on Mitochondrial Respiration and Glycolysis in Resting Platelets
In resting human WP, CORM-A1 induced a concentration-dependent decrease in OCR and biphasic changes in ECAR as analyzed by the Seahorse XFe technique (Figure 1). A slight inhibitory effect of CORM-A1 on mitochondrial respiration (Figure 1A) was already visible at the concentrations of 10 and 30 μmol/L, with stronger effects reaching ≈50% to 60% of basal OCR inhibition at concentrations of 100 to 300 μmol/L. The mitochondrial stress test (Figure 1C) revealed that CORM-A1 induced a decrease in ATP-linked respiration, maximal respiration, and spare respiratory capacity, which all may have resulted from the inhibition of cytochrome c oxidase by CO. CORM-A1 induced a slight increase in proton leak, but did not affect nonmitochondrial oxygen consumption. CORM-A1 decreased also intraplatelet ATP concentration as assessed by biochemical assay (Figure 1D).
Figure 1. Effects of CORM-A1 on mitochondrial respiration and glycolysis in resting human platelets as monitored by Seahorse XFe96 Analyzer. Oxygen consumption rate (OCR A) and extracellular acidification rate (ECAR B) measurements of human washed platelets (WP) treated with PBS (control), CORM-A1 (A1 10–300 μmol/L) or inactive CORM-A1 (iA1 300 μmol/L) followed by sequential addition of oligomycin (1 μg/mL), FCCP (carbonyl cyanide 4-[trifluoromethoxy]phenylhydrazone)/pyruvate (0.3 μmol/L/1 mmol/L) and rotenone/antimycin A (0.5/0.5 μmol/L). Bioenergetic parameters of mitochondrial respiration and glycolysis (C) were calculated as described in Materials in the Data Supplement. Data represent the means±SD of 2 independent experiments n=4–6 replicates in each experiment. D, Intracellular ATP concentration measured in platelets treated with PBS or CORM-A1 together with apyrase (1 U/mL) to remove released extracellular ATP. Data represent the means±SD of 3–5 independent experiments *P<0.05 as compared with PBS group.
Biphasic effects of CORM-A1 on glycolysis comprised a slight increase in ECAR for the lower concentrations (10–100 μmol/L) and a decrease in ECAR for the higher concentrations (200 and 300 μmol/L) of CORM-A1 (Figure 1B). To confirm that all the described effects of CORM-A1 were induced by CO, inactive CORM-A1 was used, which neither induced changes in mitochondrial respiration nor in glycolysis. The levels of LDH (lactate dehydrogenase) released into the assay medium after CORM-A1 or lysis buffer treatment confirmed that CORM-A1 in the concentration range of 10 to 300 μmol/L did not affect viability of platelets (Figure II in the Data Supplement).
Effects of CORM-A1 on Mitochondrial Respiration and Glycolysis in Activated Platelets
To characterize the effects of CO on bioenergetics of activated platelets, mitochondrial stress test was performed in WP, which were pretreated with CORM-A1 and then activated with thrombin (0.1 U/mL). Thrombin induced a slight increase (by 11±2.2%) in OCR, which was further increased by FCCP (Figure 2A and 2E). In contrast to the mild activation of OCR, thrombin induced a substantial increase (by 116±0.7%) in ECAR, which was further increased by oligomycin (Figure 2D and 2E). In the presence of CORM-A1, the thrombin-induced increase in OCR was diminished, in particular for 100 or 300 μmol/L CORM-A1 as compared with control platelets (Δ [Thr-A1] in Figure 2E). When mitochondrial respiration was further activated with FCCP, CORM-A1 in a concentration-dependent manner in a wide range of concentrations (10–300 μmol/L) diminished the increase in OCR (Δ [FCCP-A1]). The effects of CORM-A1 on thrombin-activated glycolysis were more remarkable than those on thrombin-induced increase in OCR, due to stronger activation of glycolysis by thrombin. In the presence of 100 to 300 μmol/L CORM-A1, thrombin-activated glycolysis was substantially diminished. Only a slight increase in ECAR after thrombin (by around 30%) was observed for 100 μmol/L CORM-A1 (Δ [Thr-A1] in Figure 2), whereas 300 μmol/L CORM-A1 inhibited glycolysis so strongly that thrombin-induced increase in ECAR was abrogated (Δ [Thr-A1] in Figure 2E). CORM-A1 diminished also oligomycin-induced increase in glycolysis (Δ [O-A1]) in a concentration-dependent manner (10–300 μmol/L). To confirm that the observed changes in ECAR resulted from the changes in glycolytic flux, lactate extrusion by platelets was measured after thrombin stimulation with or without CORM-A1 (Figure 2F). Thrombin (0.1 U/mL) increased lactate release, whereas CORM-A1 in a concentration-dependent manner (100–1000 μmol/L) decreased lactate concentration in supernatant of resting or thrombin-stimulated platelets (Figure 2F).
Figure 2. Effects of CORM-A1 on mitochondrial respiration and glycolysis in parallel to aggregation in activated platelets. Oxygen consumption rate (OCR A and C) and extracellular acidification rate (ECAR B and D) measurements of human washed platelets (WP) treated with PBS (control) or CORM-A1 (A1 10–300 μmol/L) followed by the addition of thrombin (0.1 U/mL) and further sequential addition of FCCP (carbonyl cyanide 4-[trifluoromethoxy]phenylhydrazone)/pyruvate (0.3 μmol/L/1 mmol/L) and rotenone/antymycin A (R/A 0.5/0.5 μmol/L A and B) or oligomycin (1 μg/mL) and R/A (C and D). Bioenergetic parameters of mitochondrial respiration and glycolysis (E) were calculated as described in the Data Supplement. Data represent the means±SD of 3 independent experiments n=4–8 replicates in each experiment. F, Concentration of lactate extruded from platelets (WP) treated with CORM-A1 (0, 100, 300, 1000 µmol/L CONT) or CORM-A1 and thrombin (0.1 U/mL Thr) data represent the means±SD of 4 independent experiments n=2 replicates in each experiment. G, Aggregation of platelets (WP) treated with CORM-A1 (10, 30, 100, 300 µmol/L) and activated with thrombin (0.5 U/mL) as compared with control data represent the means±SD of 4 independent experiments n=2 replicates in each experiment. *P<0.05, #P<0.01, $P<0.001, &P<0.0001 as compared with control group.
The comparison of CORM-A1 effects on platelet bioenergetics and aggregation revealed that concentrations of CORM-A1, which afforded nearly complete inhibition of platelet aggregation (100–300 μmol/L, Figure 2G), also led to simultaneous and complete inhibition of glycolysis and mitochondrial respiration (100–300 μmol/L, Figure 2A through 2E). Bioenergetics of platelets treated with 10 or 30 μmol/L CORM-A1 was impaired to a lesser degree, what was associated with a weaker antiplatelet effect. Inactive CORM-A1 (300 μmol/L) did not affect platelet aggregation (99.8±2.61% of control, n=5) confirming the involvement of CO in CORM-A1–induced effects. Altogether, the concentration-dependent antiaggregatory effects of CORM-A1 were mirrored by the concentration-dependent inhibition of platelet bioenergetics.
Effects of Inhibitors of Mitochondrial Respiration and Glycolysis on Platelet Bioenergetics and Platelet Aggregation-Metabolic Plasticity of Platelets
To characterize the reliance of platelets on mitochondrial respiration versus glycolysis, the effects of oligomycin (inhibitor of ATP synthase), rotenone (inhibitor of complex I), antimycin A (inhibitor of complex III), UK-5099 (inhibitor of mitochondrial pyruvate carrier), 2-deoxy-D-glucose (2DG competitive glycolytic inhibitor), and 3PO (inhibitor of PFKFB3 [6-phosphofructo-2-kinase/fructose-2,6-bisphosphatase 3]) on OCR, ECAR, and platelet aggregation were investigated.
2DG profoundly inhibited glycolysis, whereas it barely affected mitochondrial respiration (Figure 3A and 3B). Only at the highest concentration of 100 mmol/L, 2DG significantly reduced OCR to 95±0.6% of basal (P<0.05). 3PO, another inhibitor of glycolysis, slightly reduced ECAR however, even though the effect was statistically significant, it was not concentration dependent and was associated with concomitant reduction in OCR. In turn, oligomycin at 0.5 to 1 μg/mL reduced almost completely OCR, which was accompanied by a substantial increase in ECAR. Antimycin A and rotenone almost completely reduced OCR and highly increased ECAR, whereas UK-5099 slightly reduced OCR and increased ECAR. Notably, the complete inhibition of oxidative phosphorylation by oligomycin, antimycin A, or rotenone was compensated by almost a 3-fold increase in ECAR.
Figure 3. Effects of single or combined inhibition of mitochondrial respiration and glycolysis by metabolic inhibitors on platelet aggregation. Oxygen consumption rate (OCR A) and extracellular acidification rate (ECAR B) measurements of human washed platelets (WP) untreated (CONT) or treated with DMSO (as a vehicle), 2-deoxy-D-glucose (2DG), 3PO (3-[3-pyridinyl]-1-[4-pyridinyl]-2E-propen-1-one), oligomycin (OLIG), antimycin A (AA), rotenone (ROT) or UK-5099, presented as % of basal 20 minutes after addition of the reagents. Data represent the means±SD of 3 independent experiments n=4–8 replicates in each experiment. C, Aggregation of platelets (WP) untreated (CONT) or treated with 2DG, 3PO, oligomycin, antimycin A, rotenone or UK-5099, and activated with thrombin (0.5 U/mL), presented as % of control data represent the means±SD of 4 independent experiments, n=1–3 replicates in each experiment. D, OCR and ECAR measurements of WP treated with combined inhibitors: 2DG and oligomycin presented as % of basal 20 min after addition of the reagents. E, Aggregation of WP treated with combined inhibitors: 2DG and oligomycin (10 μg/mL) activated with thrombin (0.5 U/mL) presented as % of control. Data represent the means±SD of 3 independent experiments. F, Intraplatelet ATP concentration in WP treated with combined inhibitors: 2DG and oligomycin, measured in the presence of apyrase (1 U/mL). Data represent the means±SD of 3 independent experiments. *P<0.05, #P<0.01, $P<0.001, &P<0.0001 as compared with control group.
As shown in Figure 3C, those inhibitors which reduced exclusively mitochondrial respiration (oligomycin, antimycin A, rotenone, UK-5099) did not substantially inhibit platelet aggregation (less than by 10%). 2DG reduced platelet aggregation to 80% of control however, at this concentration, 2DG also slightly reduced OCR. Among metabolic inhibitors, the most effective was 3PO, which at 30 μmol/L concentration reduced platelet aggregation to 68% of control (Figure 3C). However, as demonstrated in Figure 3A and 3B, 3PO reduced not only ECAR, but also OCR.
Effects of Simultaneous Inhibition of Mitochondrial Respiration and Glycolysis on Platelet Aggregation
Figure 3D demonstrates the bioenergetics of WP after addition of oligomycin and 2DG individually or in combination. Oligomycin at a concentration of 0.5 μg/mL, which in the absence of 2DG even slightly increased metabolic activity (OCR decreased to 20±4.5% of basal, but ECAR increased up to 221±20.8% of basal), in combination with 2DG, caused a profound inhibition of platelet bioenergetics that resulted in significant inhibition of platelet aggregation (Figure 3E). Indeed, oligomycin alone (even at a concentration of 10 µg/mL) inhibited platelet aggregation by only 8% (Figure 3E), 2DG alone (up to 100 mmol/L) by 21 %. Each of them alone only slightly decreased intraplatelet ATP concentration. However, combination of 2DG and oligomycin (10 µg/mL) induced a substantial fall in intraplatelet ATP concentration (Figure 3F) explaining a profound inhibition of platelet aggregation under combined 2DG and oligomycin treatment (Figure 3E).
Effect of CORM-A1 on Metabolic Pathways in Platelets—Metabolomic Analysis
To explore the effects of CORM-A1 on platelet bioenergetics, we quantified tricarboxylic acid cycle (TCA) and glycolysis metabolites by targeted metabolomic analysis in platelets pretreated with CORM-A1 and stimulated with thrombin. As presented in Figure 4, the concentrations of TCA cycle metabolites in platelets were not substantially changed at sixth minute after thrombin activation, whereas in the presence of CORM-A1, they were reduced (fumarate by 40% and malate by over 50%). Moreover, the CORM-A1–induced changes in TCA metabolites levels were accompanied by a reduction in NAD + and an increase in NADH concentrations that is in agreement with inhibitory effects of CO on mitochondrial respiration in platelets. Interestingly, a sum of reduced and oxidized NAD (NADH together with NAD + ) was significantly decreased, whereas a ratio of NADH/NAD + was only slightly (nonsignificantly) increased after CORM-A1. In turn, glycolytic metabolites measured at sixth minute after thrombin stimulation were also not changed significantly, whereas CORM-A1 induced a massive increase in concentrations of proximal glycolytic metabolites. The concentration of hexose-6-P was increased 2-fold, while fructose-1,6-bis-phosphate and triose-phosphate (dihydroxyacetone phosphate [DHAP] and glyceraldehyde 3-phosphate [GA3P]) levels were >10-fold higher comparing to control platelets not treated with CORM-A1. Surprisingly, CORM-A1 did not induce changes in the concentrations of distal glycolytic metabolites (3-phospho-glicerate, phosphoenolpyruvate, and pyruvate Figure 5). Noteworthy, the altered balance between proximal and distal glycolytic metabolites was associated with an increased diversion of glucose into pentose phosphate pathway (PPP). Indeed, CORM-A1 induced an increase in concentrations of PPP metabolites (ribose-5-phosphate, sedoheptulose-7-phosphate, and erythrose-4-phosphate) that was not associated with an increase in NADPH or NADPH/NADP + ratio (Figure 5).
Figure 4. Effects of CORM-A1 on tricarboxylic acid cycle (TCA) metabolites and nicotinamide adenine dinucleotide (NAD + ) content in platelets. Human washed platelets (WP) suspended in PBS containing glucose (1 g/L) and glutamine (2 mmol/L) were untreated (8 min C), treated with CORM-A1 (300 μmol/L 8 min A1), treated with thrombin (0.1 U/mL T) or treated with CORM-A1 and thrombin (2+6 min A1/T). Data show amounts of metabolic by-products in TCA cycle. Data are presented as means±SD (nmoL/10 9 platelets) of 4 independent experiments *P<0.05 vs control.
Figure 5. Effects of CORM-A1 on glycolysis and pentose phosphate pathway (PPP) metabolites in platelets. Human washed platelets (WP) suspended in PBS containing glucose (1 g/L) and glutamine (2 mmol/L) were untreated (8 min C), treated with CORM-A1 (300 μmol/L 8 min A1), treated with thrombin (0.1 U/mL T) or treated with CORM-A1 and thrombin (2+6 min A1/T). Data show amounts of metabolic by-products in glycolysis and PPP. Data are presented as means±SD (nmoL/10 9 platelets) of 4 independent experiments. DHAP indicates dihydroxyacetone phosphate F-1,6-BP, fructose-1,6-bis-phosphate GA3P, glyceraldehyde 3-phosphate PEP, phosphoenolpyruvate and TCA, tricarboxylic acid cycle. *P<0.05 vs control.
These results, showing an activation of PPP and accumulation of proximal glycolytic metabolites, strongly suggest that the inhibition of glycolysis by CORM-A1 was targeted at the level of GAPDH. However, an activity of GAPDH in platelet was not inhibited directly by CORM-A1 (Figure 5). Similarly, another rate-limiting step of proximal glycolysis—the reaction catalyzed by PFK-1—was not modulated by CORM-A1, as evidence by the lack of effects of 8-minute long incubation with CORM-A1 on PFK-1 activity in platelets (data not shown). Only prolonged 30-minute incubation allowed to observe an inhibition of this enzyme (Figure IA), that was, however, not relevant to the inhibition of glycolysis by short-term exposure of CORM-A1. Another possible explanation for the inhibition of glycolysis at the level of GAPDH by CORM-A1 might have been linked directly to NAD + depletion (Figure 4).
Reversal of Effects of Pyruvate on CORM-A1–Induced Inhibition of Platelet Aggregation and Glycolysis Role of LDH and NAD + Regeneration
As LDH is a major enzymatic source for NAD + regeneration, we tested whether pyruvate could reverse antiplatelet effects of CORM-A1. As shown in Figure 6A, in the presence of pyruvate (1 mmol/L) the inhibitory effect of CORM-A1 on platelet aggregation was lost (Figure 6A) with concomitant increase in lactate release (Figure 6B) and ECAR (Figure 6C), in contrast to the absence of pyruvate (Figure 1). These data confirmed a high activity of LDH, catalyzing the conversion of exogenous pyruvate into lactate, accompanied by the regeneration of cytosolic NAD + from NADH. Importantly, in the presence of 1 mmol/L pyruvate, the accumulation of proximal glycolysis intermediates (fructose-1,6-bis-phosphate and DHAP/GA3P Figure 6D through 6F) was abrogated, and glycolysis flux was efficient again, as also evidenced by the lack of accumulation of PPP intermediates (ribose-5-phosphate, sedoheptulose-7-phosphate, erythrose-4-phosphate Figure 6H through 6K). These results suggest that pyruvate–induced LDH–dependent regeneration of NAD + reversed glycolysis inhibition at the level of GAPDH. Furthermore, these results demonstrate that antiaggregatory effect of CORM-A1 was dependent on NAD + availability, which was modulated by LDH activity. To substantiate the finding on the role of LDH and pyruvate-dependent regeneration of NAD + in the reversal of the CORM-A1–induced effects, we demonstrated that in the presence of GSK2837808A (LDH inhibitor 5 μmol/L) the effect of pyruvate (100 μmol/L) was lost, and 300 μmol/L CORM-A1 inhibited platelet aggregation again (Figure 6P). Noteworthy, GSK2837808A in the absence of pyruvate, when added alone, only slightly reduced platelet aggregation, but in combination with CORM-A1 displayed significant antiplatelet effect (Figure 6P). Furthermore, combination of GSK2837808A with antimycin A resulted in a clear-cut antiaggregatory effect, even though neither of them given alone affected platelet aggregation (Figure 6Q). To test a putative functional role of NAD + derived from malate-aspartate shuttle in platelets, we combined CORM-A1 or antimycin A with aminooxyacetic acid (an inhibitor of malate-aspartate shuttle), but in contrast to the combination of CORM-A1 with GSK2837808A, we did not observe enhancement of antiaggregatory effects of CORM-A1 (data not shown). Dimethyl malonate, which intracellularly is converted to malonate (an inhibitor of SDH [succinate dehydrogenase]), did not show significant antiaggregatory effects alone or in various combinations, excluding a possible role of SDH and malate-aspartate shuttle to regenerate NAD + in CORM-A1–treated platelets.
Figure 6. Effects of pyruvate on antiplatelet activity of CORM-A1 in platelets. Human washed platelets (WP) were suspended in assay medium (DMEM) supplemented with glucose (1 g/L) and glutamine (2 mmol/L) with or without pyruvate (1 mmol/L). A, Aggregation of WP treated with CORM-A1 (300 µmol/L) and activated with thrombin (0.1 U/mL) as compared with control data represent the means±SD of 4 independent experiments n=1–2 replicates in each experiment. B, Concentration of lactate extruded from platelets (WP) treated with CORM-A1 (0, 100, 300, 1000 µmol/L) data represent the means±SD of 3 independent experiments. C, Extracellular acidification rate (ECAR) measurements of WP treated with PBS (control) or CORM-A1 (A1 100, 300, 1000 μmol/L) in DMEM supplemented with glucose (1 g/L), glutamine (2 mmol/L), and pyruvate (1 mmol/L) Seahorse XFe96 Analyzer. Data represent means±SD from a representative experiment, n=3–6 technical replicates. D–O, Human WP suspended in PBS containing glucose (1 g/L) and glutamine (2 mmol/L) without (C, A1) or with pyruvate (1 mmol/L P, A1/P) were untreated or treated with CORM-A1 (300 µmol/L), followed by activation with thrombin (0.1 U/mL 6 min) data demonstrate concentrations of selected metabolic by-products of glycolysis, PPP or mitochondrial respiration are presented as means±SD (nmol/10 9 platelets) of 3 independent experiments, n=2 replicates in each experiment. P, Aggregation of WP treated with pyruvate (100 µmol/L), CORM-A1 (300 µmol/L) and GSK2837808A (GSK 5 µmol/L). Q, Aggregation of WP control or preincubated with dimethyl malonate (DMM 15 min) followed by treatment with antimycin A (AA 10 µmol/L) and GSK2837808A (GSK 3 µmol/L). Data represent the means±SD of 4 independent experiments n=1–3 replicates in each experiment. DHAP indicates dihydroxyacetone phosphate F-1,6-BP, fructose-1,6-bis-phosphate and NAD + , nicotinamide adenine dinucleotide. *P<0.05, #P<0.01, $P<0.001, &P<0.0001.
To further confirm, the key role of cytosolic NAD + depletion in the antiplatelet effects of CORM-A1, we examined the effects of CORM-A1 on aggregation of platelets pretreated with 78c (0.3–20 μmol/L), an inhibitor of CD38—one of the major NAD + consuming enzymes. The inhibition of CD38 by 1 µmol/L 78c blunted antiaggregatory effect of CORM-A1 by 13%, further supporting the key role of NAD + depletion in antiplatelet effects of CORM-A1.
Platelet aggregation is an energy-demanding process, 15,16 and recent evidence suggest a high substrate plasticity of metabolism in platelets. 17,19,42,43 In the present work, to our knowledge for the first time, we provide evidence that CO affords antiplatelet effects by simultaneous and efficient inhibition of 2 major ATP–generating pathways: mitochondrial respiration, attributed to the inhibition of cytochrome c oxidase, and glycolysis, ascribed to cytosolic NAD + depletion. Interestingly, an inhibition of glycolysis or mitochondrial respiration individually, using classical metabolic inhibitors, did not inhibit platelet aggregation because these 2 pathways compensate each other, as also suggested previously. 19 Combined inhibition of glycolysis and mitochondrial respiration with oligomycin and 2DG, respectively, afforded pronounced platelet inhibition. In this context, our results showing that CORM-A1–induced a simultaneous inhibition of glycolysis and mitochondrial respiration underscores the ability of CO released from CORM-A1 to overcome metabolic plasticity of platelets. CORM-A1–derived CO effectively inhibits platelet aggregation by compromising at the same time 2 major pathways of platelet bioenergetics by distinct mechanisms culminating in NAD + and ATP depletion. Given the fact that the sGC is a target for CO gas in platelets, 9 but not for CORM-A1 11,12 we suspect that bioenergetic effects of CORM-A1 may not necessarily be shared by gaseous CO.
The effects of CO on mitochondrial respiration and glycolysis, although clearly observed for resting platelets, were more pronounced in platelets metabolically activated by thrombin, oligomycin (activating platelet glycolysis Figure 3A and 3B), or FCCP (activating maximal mitochondrial respiration Figure 1C). Platelets in the settings of prothrombotic activation increase energetic demands 15,16,19 and, apparently in such settings, platelets also became more susceptible to the inhibitory effects of CO. Altogether, the concentration-dependent inhibition of platelet aggregation by CORM-A1 was accompanied by the concentration-dependent inhibition of platelet bioenergetics both ensued in the same range of CORM-A1 concentrations and pertained to 2 major pathways of ATP generation, overcoming substrate plasticity of platelets.
Indeed, metabolic plasticity enables platelet aggregation even after inhibition of a single metabolic pathway, 16,19 because platelets use diverse metabolic fuels—not only glucose, free fatty acids and glutamine, but also glycogen, citrate (added to the blood during collection), albumin, and acetate. 17,19,42,44 Recently, Ravi et al 19 demonstrated that the activation of platelets with thrombin results in metabolic reprogramming, leading to increased aerobic glycolysis, fatty acid oxidation and glutaminolysis, which all compensate each other. We confirmed platelet metabolic plasticity in our experimental set-up (Figure 3), demonstrating that selective inhibition of mitochondrial respiration (by oligomycin, antimycin A, rotenone) did not or only slightly (less than by 10%) inhibited platelet aggregation. Single inhibitors of mitochondrial respiration were almost ineffective in inhibition of platelet aggregation (Figure 3C), but inhibition of mitochondrial respiration was compensated by an almost 3-fold increase in glycolysis (Figure 3A and 3B), which probably met platelet aggregation energy requirements. In turn, glycolysis inhibition by 100 mmol/L 2DG slightly (by 21%) inhibited platelet aggregation, implying that glycolysis might be more important for platelet activity. It was suggested that in resting platelets ATP is produced mainly (65%) in glycolysis and only 35% in oxidative phosphorylation. 18 Clearly, upon platelet activation, the contributions of these metabolic pathways may be different, depending on the substrates’ availability, underscoring platelets’ plasticity. Interestingly, 3PO is known as an inhibitor of glycolysis, 45 but recently it was demonstrated to target and inhibit also TCA cycle and mitochondrial respiratory chain. 46 In our model, 3PO appeared to inhibit not only glycolysis, but also oxygen consumption (Figure 3A and 3B), resulting in the inhibition of platelet aggregation in contrast to other metabolic inhibitors given individually. Combined inhibition of both glycolysis and oxidative phosphorylation with 2DG and oligomycin (Figure 3D and 3E) resulted in almost complete inhibition of metabolism and concomitant substantial reduction of platelet aggregation. Given extraordinary platelet metabolic plasticity, high efficacy of CO released from CORM-A1 in the inhibition of platelet aggregation rely on the inhibition of both oxidative phosphorylation and glycolysis and subsequent efficient blocking of ATP production necessary for platelet aggregation.
As regards inhibition of oxidative phosphorylation, we demonstrated that in resting platelets, CO released from CORM-A1 inhibited mitochondrial respiration in a concentration-dependent manner. A CORM-A1–induced fall in concentrations of fumarate and malate (Figure 4) confirms reduced TCA turnover in the presence of CO, while the imbalance in NAD + and NADH concentrations, seen as increased NADH/NAD + ratio, confirms the inhibition of oxidative phosphorylation by CO. Altogether, metabolomic results, together with the profile of CORM-A1–induced effects on OCR, provide evidence that CORM-A1 inhibited mitochondrial respiration in platelets, most probably by binding to cytochrome c oxidase, which is a well-known target for CO action. 47 Moreover, CORM-A1–induced falls in fumarate and malate (Figure 4) may also suggest succinate dehydrogenase as another possible target for CO in mitochondria. However it seems to be of less significance for platelets’ mitochondrial respiration, as evidenced by the lack of antiplatelet effects of dimethyl malonate (Figure 6Q), a cell permeable precursor of succinate dehydrogenase inhibitor. 48 Surprisingly, mitochondria in platelets were more susceptible to the inhibitory effects of CO, as compared with microglia 30 or endothelial cells, 31 where only higher concentrations of CO inhibited mitochondrial respiration. These results suggest an important regulatory role of mitochondrial ATP turnover in platelets.
Interestingly, CO modulated platelet glycolysis in a biphasic way—activated by lower and inhibited by higher concentrations of CORM-A1 (Figure 1). These results suggest that the observed activation of glycolysis at lower concentrations of CORM-A1 was a compensatory response to the inhibition of mitochondrial respiration. However, CORM-A1 at higher concentrations induced inhibition of mitochondrial respiration and glycolysis, the latter possibly, by separate cytosolic target for CO. In the presence of CORM-A1, we noted a strong increase in proximal metabolites of glycolysis (hexose-6-phosphate, fructose-1,6-bis-phosphate, DHAP and GA3P) and PPP, contrary to the metabolites of distal glycolysis (3-phospho-glicerate, phosphoenolpyruvate and pyruvate), which were unchanged (Figure 5). These results suggest that glycolysis inhibition by CO might be targeted at GAPDH, which contains a haem moiety, and CO–haem interaction may inhibit GAPDH. 49 However, the activity of GAPDH in platelets was not inhibited by CORM-A1 (Figure 5). Another reason for the inhibition of GAPDH activity and subsequent inhibition of glycolysis (Figure 5) could be a substantial drop in NAD + (Figure 4), primarily linked to the inhibition mitochondrial respiration by CO, and a subsequent increase in NADH/NAD + ratio. However, due to NAD + compartmentalization (mitochondria contain up to 70% of total NAD content 50 ), it is necessary to consider a separate cytosolic NAD + pool involved in glycolysis, which may be regenerated from the metabolism of pyruvate by LDH, or through the action of glycerol-3-phosphate shuttle or malate-aspartate shuttle. 51 In the present work, we exclude the importance of malate-aspartate shuttle in NAD + regeneration in platelets (lack of the effects of aminooxyacetic acid added alone or in combination with other compounds on platelet aggregation data not shown), but we demonstrate that the addition of pyruvate: (1) induced LDH–dependent regeneration of NAD + accompanying lactate extrusion, (2) reversed glycolysis inhibition at the level of GAPDH, and most importantly, (3) abrogated antiaggregatory effects of CORM-A1 (Figure 6). Indeed, in the presence of pyruvate, NADH/NAD + ratio was normalized, and accumulation of proximal glycolysis and PPP intermediates were reversed (Figure 6D through 6F and 6H through 6K). Furthermore, inhibition of LDH–dependent regeneration of NAD + by GSK2837808A resulted in the elimination of the effect of pyruvate (100 µmol/L) on CORM-A1–inhibited platelet aggregation. Inhibition of LDH with simultaneous inhibition of oxidative phosphorylation (Figure 6Q) reduced platelet aggregation. These results confirmed the key role of LDH–dependent NAD + regeneration for glycolytic flux, the inhibition of which together with the inhibition of mitochondrial respiration resulted in the inhibition of platelet aggregation. Accordingly, NAD + depletion and subsequent inhibition of glycolysis together with mitochondrial respiration are 2 major targets for CO–induced downregulation of platelet bioenergetics. In the present work, we indicate the importance of NAD + in platelet glycolysis and aggregation, and suggest that CO, apart from inhibiting cytochrome c oxidase in mitochondria, has another cytosolic target(s) decreasing availability of NAD + , that was, however, not ultimately identified here.
Generally, among the enzymes involved in NAD + turnover, there are NAD + –regenerating enzymes (as LDH, malate-aspartate shuttle, glycerol-3-phosphate shuttle), NAD + –salvage enzymes (as NAD + synthase), and NAD + –consuming enzymes (sirtuins, poly [ADP-ribose] polymerases, cADP-ribose synthases). 51 Little is known about the NAD + turnover in platelets, however, it was shown that platelets contain an enzymatic machinery to metabolize nicotinamide riboside into NAD. 52 We directed our attention towards CD38 (cADP-ribose synthase), which generates one molecule of cADP-ribose for every 100 molecules of NAD + hydrolyzed, and plays a role of major regulator of cellular NAD + availability. 53 In fact, we found that 1 μmol/L 78c (inhibitor of CD38) induced a slight, but significant reduction of CORM-A1–induced inhibition of platelet aggregation, supporting the view that modulation of availability of NAD + by CD38 or other enzymes of NAD + turnover may influence antiplatelet effects of CORM-A1. However, a direct target for CO responsible for NAD + –deficiency still needs to be identified. It was demonstrated in pancreatic islets that CO activates CD38 through the activation of sGC. 54 Nevertheless, in platelets CO released from CORMs does not act through sGC (see Refs 11,12 and Figure IV in the Data Supplement) making CO/cGMP/CD38 pathway unlikely to operate in platelets.
Previously, the effects of CO on glycolysis were demonstrated among others in endothelial cells, whereby CO inhibited glycolysis and activated PPP. 31–33 Some authors claim that CO-inhibition of glycolysis is attributed to the inhibition of CBS (cystathionine-β-synthase), which through the reduction of methylation of PFK-1/fructose bisphosphatase type-3 (PFKFB3 an activator of glycolytic flux) inhibited glycolysis on the level of PFK-1 and redirected glucose towards PPP. 55 As shown in Figure 5, the concentration of fructose-1,6-phosphate as well as the combined concentration of DHAP and GA3P were increased in response to CORM-A1, whereas the concentrations of 3-P-glycerate, phosphoenolpyruvate and pyruvate were unchanged (1,3-bisphosphoglycerate was not analyzed). At first glance, the data could exclude the involvement of reduced activation of PFKFB3 in platelets in response to CO because increased, but not decreased concentration of fructose-1,6-phosphate was measured. Prolonged 30-minute incubation of platelets with CORM-A1 allowed to observe an inhibition of PFK-1 activity, however, not strong enough to avoid accumulation of fructose-1,6-bis-phosphate, DHAP and GA3P (Figure III in the Data Supplement). These data suggest that CBS inhibition, and a consequent PFKFB3 demethylation leading to PFK-1 inhibition, was not responsible for inhibition of glycolysis in platelets treated with CORM-A1, that was in turn ascribed here to NAD + depletion.
Further studies are needed for a detailed explanation of biochemical mechanisms of CO action in platelets on NAD + metabolome and elucidation whether haem-dependent or independent mechanisms are involved. Hemoproteins, such as voltage-gated K + channels 56 and epithelial Na + channels, 57 seem unlikely as targets for CO in platelets, as their link with NAD + – consuming processes is not obvious. In turn, large-conductance calcium-regulated K + channel, known to display a relatively high affinity to CO, is not abundant in platelets. 56
In summary, the results of the present work, to our knowledge for the first time, provide evidence that CO affords antiaggregatory effects by inhibition of mitochondrial respiration and glycolysis, attributed to the inhibition of cytochrome c oxidase and depletion of cytosolic NAD + , respectively. The results of our study not only uncover platelet-specific action of CO on bioenergetics, but also point out that metabolism-targeted antiplatelet therapeutic strategies may prove useful for inhibiting excessive platelet activation in various diseases.
This section discusses the process of fermentation. Due to the heavy emphasis in this course on central carbon metabolism, the discussion of fermentation understandably focuses on the fermentation of pyruvate. Nevertheless, some of the core principles that we cover in this section apply equally well to the fermentation of many other small molecules.
The "purpose" of fermentation
The oxidation of a variety of small organic compounds is a process that is utilized by many organisms to garner energy for cellular maintenance and growth. The oxidation of glucose via glycolysis is one such pathway. Several key steps in the oxidation of glucose to pyruvate involve the reduction of the electron/energy shuttle NAD + to NADH. You were already asked to figure out what options the cell might reasonably have to reoxidize the NADH to NAD + in order to avoid consuming the available pools of NAD + and to thus avoid stopping glycolysis. Put differently, during glycolysis, cells can generate large amounts of NADH and slowly exhaust their supplies of NAD + . If glycolysis is to continue, the cell must find a way to regenerate NAD + , either by synthesis or by some form of recycling.
In the absence of any other process&mdashthat is, if we consider glycolysis alone&mdashit is not immediately obvious what the cell might do. One choice is to try putting the electrons that were once stripped off of the glucose derivatives right back onto the downstream product, pyruvate, or one of its derivatives. We can generalize the process by describing it as the returning of electrons to the molecule that they were once removed, usually to restore pools of an oxidizing agent. This, in short, is fermentation. As we will discuss in a different section, the process of respiration can also regenerate the pools of NAD + from NADH. Cells lacking respiratory chains or in conditions where using the respiratory chain is unfavorable may choose fermentation as an alternative mechanism for garnering energy from small molecules.
An example: lactic acid fermentation
An everyday example of a fermentation reaction is the reduction of pyruvate to lactate by the lactic acid fermentation reaction. This reaction should be familiar to you: it occurs in our muscles when we exert ourselves during exercise. When we exert ourselves, our muscles require large amounts of ATP to perform the work we are demanding of them. As the ATP is consumed, the muscle cells are unable to keep up with the demand for respiration, O2 becomes limiting, and NADH accumulates. Cells need to get rid of the excess and regenerate NAD + , so pyruvate serves as an electron acceptor, generating lactate and oxidizing NADH to NAD + . Many bacteria use this pathway as a way to complete the NADH/NAD + cycle. You may be familiar with this process from products like sauerkraut and yogurt. The chemical reaction of lactic acid fermentation is the following:
Pyruvate + NADH &harr lactic acid + NAD +
Figure 1. Lactic acid fermentation converts pyruvate (a slightly oxidized carbon compound) to lactic acid. In the process, NADH is oxidized to form NAD + . Attribution: Marc T. Facciotti (original work)
Energy story for the fermentation of pyruvate to lactate
An example (if a bit lengthy) energy story for lactic acid fermentation is the following:
The reactants are pyruvate, NADH, and a proton. The products are lactate and NAD + . The process of fermentation results in the reduction of pyruvate to form lactic acid and the oxidation of NADH to form NAD + . Electrons from NADH and a proton are used to reduce pyruvate into lactate. If we examine a table of standard reduction potential, we see under standard conditions that a transfer of electrons from NADH to pyruvate to form lactate is exergonic and thus thermodynamically spontaneous. The reduction and oxidation steps of the reaction are coupled and catalyzed by the enzyme lactate dehydrogenase.
A second example: alcohol fermentation
Another familiar fermentation process is alcohol fermentation, which produces ethanol, an alcohol. The alcohol fermentation reaction is the following:
Figure 2. Ethanol fermentation is a two-step process. Pyruvate (pyruvic acid) is first converted into carbon dioxide and acetaldehyde. The second step converts acetaldehyde to ethanol and oxidizes NADH to NAD + . Attribution: Marc T. Facciotti (original work)
In the first reaction, a carboxyl group is removed from pyruvic acid, releasing carbon dioxide as a gas (some of you may be familiar with this as a key component of various beverages). The second reaction removes electrons from NADH, forming NAD + and producing ethanol (another familiar compound&mdashusually in the same beverage) from the acetaldehyde, which accepts the electrons.
Fermentation pathways are numerous
While the lactic acid fermentation and alcohol fermentation pathways described above are examples, there are many more reactions (too numerous to go over) that Nature has evolved to complete the NADH/NAD + cycle. It is important that you understand the general concepts behind these reactions. In general, cells try to maintain a balance or constant ratio between NADH and NAD + when this ratio becomes unbalanced, the cell compensates by modulating other reactions to compensate. The only requirement for a fermentation reaction is that it uses a small organic compound as an electron acceptor for NADH and regenerates NAD + . Other familiar fermentation reactions include ethanol fermentation (as in beer and bread), propionic fermentation (it's what makes the holes in Swiss cheese), and malolactic fermentation (it's what gives Chardonnay its more mellow flavor&mdashthe more conversion of malate to lactate, the softer the wine). In Figure 3, you can see a large variety of fermentation reactions that various bacteria use to reoxidize NADH to NAD + . All of these reactions start with pyruvate or a derivative of pyruvate metabolism, such as oxaloacetate or formate. Pyruvate is produced from the oxidation of sugars (glucose or ribose) or other small, reduced organic molecules. It should also be noted that other compounds can be used as fermentation substrates besides pyruvate and its derivatives. These include methane fermentation, sulfide fermentation, or the fermentation of nitrogenous compounds such as amino acids. You are not expected to memorize all of these pathways. You are, however, expected to recognize a pathway that returns electrons to products of the compounds that were originally oxidized to recycle the NAD + /NADH pool and to associate that process with fermentation.
Figure 3. This figure shows various fermentation pathways using pyruvate as the initial substrate. In the figure, pyruvate is reduced to a variety of products via different and sometimes multistep (dashed arrows represent possible multistep processes) reactions. All details are deliberately not shown. The key point is to appreciate that fermentation is a broad term not solely associated with the conversion of pyruvate to lactic acid or ethanol. Source: Marc T. Facciotti (original work)
A note on the link between substrate-level phosphorylation and fermentation
Fermentation occurs in the absence of molecular oxygen (O2). It is an anaerobic process. Notice there is no O2 in any of the fermentation reactions shown above. Many of these reactions are quite ancient, hypothesized to be some of the first energy-generating metabolic reactions to evolve. This makes sense if we consider the following:
- The early atmosphere was highly reduced, with little molecular oxygen readily available.
- Small, highly reduced organic molecules were relatively available, arising from a variety of chemical reactions.
- These types of reactions, pathways, and enzymes are found in many different types of organisms, including bacteria, archaea, and eukaryotes, suggesting these are very ancient reactions.
- The process evolved long before O2 was found in the environment.
- The substrates, highly reduced, small organic molecules, like glucose, were readily available.
- The end products of many fermentation reactions are small organic acids, produced by the oxidation of the initial substrate.
- The process is coupled to substrate-level phosphorylation reactions. That is, small, reduced organic molecules are oxidized, and ATP is generated by first a red/ox reaction followed by the substrate-level phosphorylation.
- This suggests that substrate-level phosphorylation and fermentation reactions coevolved.
Consequences of fermentation
Imagine a world where fermentation is the primary mode for extracting energy from small molecules. As populations thrive, they reproduce and consume the abundance of small, reduced organic molecules in the environment, producing acids. One consequence is the acidification (decrease in pH) of the environment, including the internal cellular environment. This can be disruptive, since changes in pH can have a profound influence on the function and interactions among various biomolecules. Therefore, mechanisms needed to evolve that could remove the various acids. Fortunately, in an environment rich in reduced compounds, substrate-level phosphorylation and fermentation can produce large quantities of ATP.
It is hypothesized that this scenario was the beginning of the evolution of the F0F1-ATPase, a molecular machine that hydrolyzes ATP and translocates protons across the membrane (we'll see this again in the next section). With the F0F1-ATPase, the ATP produced from fermentation could now allow for the cell to maintain pH homeostasis by coupling the free energy of hydrolysis of ATP to the transport of protons out of the cell. The downside is that cells are now pumping all of these protons into the environment, which will now start to acidify.
Online Biology Tutor
This tutorial presents fermentation at a level appropriate for most undergraduate biology classes and the MCAT exam.
Fermentation allows glycolysis to continue in the absence of oxygen, yielding a small amount of ATP. During fermentation 2 electrons a proton are transferred from each NADH product of glycolysis to pyruvate or a derivative acceptor molecule to regenerate NAD+. NAD+ is used to continue glycolysis.
Byproducts of fermentation include lactate, butate and ethanol.
1. Fermentation is an energy yielding process because it allows glycolysis to continue in the absence of oxygen. Since a net amount of 2 ATP is produced by glycolysis, a net amount of 2 ATP is also produced by fermentation.
2. NAD+ is required to complete glycolysis. NAD+ must be present to accept hydrogen before any ATP can be produced.
3. During glycolysis two molecules of NAD+ are reduced to NADH.
4. During lactic acid fermentation pyruvate is reduced to lactic acid and NADH is oxidized to regenerate NAD+.
5. In lactic acid fermentation hydrogen atoms (along with two electrons) are added directly on to pyruvate converting it to lactic acid. Pyruvate is the terminal electron acceptor and lactic acid is the product.
TERMS TO KNOW
adenosine triphosphate (ATP)
lactic acid fermentation
nicotinamide adenine dinucleotide (NAD+ / NADH)
terminal electron acceptor
RELATED TOPICS (Note: These links will go active as videos appear online.)
The conversion of NAD+ to NADH, and vice versa, are essential reactions in creating ATP during what’s called cellular respiration. The food you consume goes through three phases to become energy: glycolysis, the Krebs Cycle, and the electron transport chain.
In glycolysis and the Krebs cycle, NADH molecules are formed from NAD+. Meanwhile, in the electron transport chain, all of the NADH molecules are subsequently split into NAD+, producing H+ and a couple of electrons, too. The H+ are used to power a sort-of "pump" that sits on the inner membrane of the mitochondria, creating lots of energy in the form of ATP. Once the H+ have cycled through the pump, they subsequently merge with the electrons and a molecule of oxygen to form water. All of the three phases of respiration generate ATP however, the greatest yield of ATP is during the electron transport chain.
The cell uses NAD+ and NADH in other reactions outside of ATP production, too. In liver cells, for instance, the enzymes alcohol dehydrogenase (ADH) and aldehyde dehydrogenase (ALDH) utilize NAD+ as an oxidizing agent in order to breakdown ethanol from alcoholic drinks into a less toxic compound called acetate. In each of the enzymatic reactions, NAD+ accepts two electrons and a H+ from ethanol to form NADH.
We would like to kindly thank Yoh Terada for his helpful advice during the creation of this review. KJM is the recipient of a Heart and Stroke Foundation of Canada research fellowship award. CC is an employee of the Nestlé Institute of Health Sciences S.A. JA is the Nestlé Chair in Energy Metabolism. Work in the laboratory is supported by the ಜole Polytechnique Fຝérale de Lausanne, the National Institutes of Health (R01AG043930), the Swiss National Science Foundation (31003A-124713), and Systems X (51RTP0-151019).
Consumption of NAD+ in glycolysis - Biology
The question stem is describing a mitochondrial disease, which commonly present with lactic acidosis. There is an increase in anaerobic forms of energy production (glycolysis). The mitochondria are faulty, so they can’t use the end product of glycolysis (pyruvate) in TCA. Instead pyruvate is shunted over and is used by LDH (lactate dehydrogenase) to generate pyruvate.
Aside: Recall that LDH uses NADH and generates NAD+. Deficiency of LDH can lead to loss of regeneration of NAD+ and inhibits glycolysis.
So this question was something I really struggled with. I didn't recognize that the presentation was of MERRF as someone stated below, and I know you don't need to know that to answer the question, but it would have been helpful. My biggest frustration was the wording of the biopsy results "abnormal accumulations of mitochondria." This annoyed me because the definition of ragged red fibers (which I'm assuming was their intention) is "accumulations of abnormal mitochondria." Those are two very different statements in my mind, lol. The first, to me, just means there's too much mitochondria, but the second means there's too much AND they aren't functioning properly. It's also just the fact of remembering all of the terms for ETC at the time of reading the question (i.e. I didn't think about the fact that ETC is also called cellular respiraton or just respiration).
I also didn't really understand fully what VO2max is = "VO2 max, also known as maximal oxygen uptake, is the measurement of the maximum amount of oxygen a person can utilize during intense exercise. and is based on the premise that the more oxygen consumed during exercise, the more the body will generate adenosine triphosphate (ATP) energy in cells. VO2 max is reached when your oxygen consumption remains at a steady state despite an increase in the workload. It is at this plateau that the [muscle] moves from aerobic metabolism to anaerobic metabolism" https://www.verywellfit.com/what-is-vo2-max-3120097.
Based purely on this definition, where VO2max = essentially the time at which aerobic switches to anaerobic respiration, my interpretation of too much mitochondria vs too much and bad mitochrondria didn't matter, because even when the mitochondria are functioning properly, they reach a point and switch to anaerobic, thus if there was too much normal mitochondria, this would occur faster because there would be more overall cellular respiration occuring, meaning the body would switch to anaerobic and utilize glycolisis to lactate for energy, stop utilizing the mitochondria, and thus VO2max would decrease.
HOWEVER. because this is a MERRF question, the order of events is a little different, despite the outcome being the same (at least that's how I understand it). So, I think the key to any mitochondrial disorder is remembering that the mutations are almost certainly going to affect an encoded protein and thus a deficiency of that protein. One article that I found said that the tRNA mutations (as in MERRF) cause: "disrupt mitochondrial protein synthesis, decreasing the activity of Complex I and to a lesser extent Complex IV. which decreases respiration and lowers proton pumping, dramatically decreasing the membrane potential and proton electrochemical potential gradient across the mitochondrial inner membrane. The proton electrochemical potential gradient is the driving force for ATP synthesis and decreasing it substantially lowers the maximal rate of ATP synthesis." https://febs.onlinelibrary.wiley.com/doi/full/10.1046/j.1432-1327.1999.00066.x
Based on my understanding of oxidative phosphorylation, O2 consumption (i.e. taking the electron from complex IV and putting it on 1/2 O2 to create H2O and H+ drives the proton gradient which drives ATP production. Thus: deficient respiratory oxidation (i.e. mtDNA mutations of the ETC enzymes) leads to lowered O2 consumption (so lowered VO2 max) which then leads to lowered ATP production, and thus defective mitochondria. Then, lowered mitochondrial function leads to decreased aerobic respiration shunting ATP production to anaerobic respiration, driven by glycolysis, and thus increasing lactate levels.
Hope this helps! This took me WAY TOO LONG to figure out, lol, but hopefully I never freaking forget it, lol.
We are grateful to Kubo T for her excellent technical assistance and Nakagawa lab members for helpful discussion.
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Keywords : NAD+, nampt, alpha-ketoglutarate, demethylation, adipocyte, preadipocyte, differentiation, metabolomics
Citation: Okabe K, Nawaz A, Nishida Y, Yaku K, Usui I, Tobe K and Nakagawa T (2020) NAD+ Metabolism Regulates Preadipocyte Differentiation by Enhancing α-Ketoglutarate-Mediated Histone H3K9 Demethylation at the PPARγ Promoter. Front. Cell Dev. Biol. 8:586179. doi: 10.3389/fcell.2020.586179
Received: 23 July 2020 Accepted: 03 November 2020
Published: 24 November 2020.
Maria Barile, University of Bari Aldo Moro, Italy
Brijesh Kumar Singh, Duke-NUS Medical School, Singapore
Tibor Kristian, University of Maryland, United States
Copyright © 2020 Okabe, Nawaz, Nishida, Yaku, Usui, Tobe and Nakagawa. This is an open-access article distributed under the terms of the Creative Commons Attribution License (CC BY). The use, distribution or reproduction in other forums is permitted, provided the original author(s) and the copyright owner(s) are credited and that the original publication in this journal is cited, in accordance with accepted academic practice. No use, distribution or reproduction is permitted which does not comply with these terms.