Solution Chemistry of Dihydroxyacetone and Synthesis of Monomeric Dihydroxyacetone

Abstract

Dihydroxyacetone (DHA) is a major byproduct of e-cigarette combustion and is the active ingredient in sunless tanning products. Mounting evidence points to its damaging effects on cellular functions. While developing a simple synthetic route to monomeric [¹³C₃]DHA for flux metabolic studies that compared DHA and glyceraldehyde (GA) metabolism, we uncovered that solid DHA ages upon storage and differences in the relative abundance of each of its isomer occur when reconstituted in an aqueous solution. While all three of the dimeric forms of DHA ultimately resolve to the ketone and hydrated forms of monomeric DHA once in water at room temperature, these species require hours rather than minutes to reach an equilibrium favoring the monomeric species. Consequently, when used in bolus or flux experiments, the relative abundance of each isomer and its effects at the time of application is dependent on the initial DHA isomeric composition and concentration, and time of equilibration in solution before use. Here, we make recommendations for the more consistent handling of DHA as we report conditions that ensure that DHA is present in its monomeric form while in solutions, conditions used in an isotopic tracing study that specifically compared monomeric DHA and GA metabolism in cells.

Introduction

Glyceraldehyde (GA) and dihydroxyacetone (DHA) are trioses that once phosphorylated to 3-phosphoglyceraldehyde (GAP) and 1-phospho, 3-hydroxyacetone (DHAP), respectively, are most commonly known intermediates of glycogenolysis and gluconeogenesis.¹ In addition to being an endogenous metabolite, DHA is also the active ingredient in sunless tanning products and a byproduct of electronic cigarette (e-cigarette) combustion,² both leading to elevated exposures to DHA through inhalation and absorption^2,3 In biological systems, these trioses can also act as nucleic acids’ and proteins’ glycating agents.^4,5 Once phosphorylated and isomerized, they also can become building blocks in several other intracellular pathways, including glycolipids and amino acid biosynthesis. In addition to the differences in their glycating properties, these two trioses are likely to enter biosynthetic pathways at different rates and thus present distinct metabolic and physiological profiles.^6,7 Mounting evidence now associates endogenous DHA’s reactivity with replication stress and cell death^8,9 via processes closely reminiscent of fructose-caused dysfunction.¹⁰ Like fructose, DHA promotes oxidative stress, mitochondrial dysfunction, and gene and protein expression changes.

Triose metabolism, and more specifically DHA metabolism, has long been under scrutiny for DHA’s ability to interfere with redox and biosynthetic balance in tissues.¹¹ More recently, metabolic imaging using [2-¹³C]DHA and NMR spectroscopy confirmed that DHA is rapidly distributed and absorbed into tissues and cells, where it is converted to glycolytic intermediates, amino acids, glycerol, and glucose 6-phosphate.¹²⁻¹⁷ While extremely comprehensive, sensitive, and informative, this method remains limited in technological convenience, as it requires access to high-field NMR instruments and akin expertise. Alternatively, facilities that perform liquid or gas chromatography coupled to mass spectrometry (LC-MS or GC-MS) are more widely available, as is the expertise, and thus provide greater opportunities for more varied experimental investigations (e.g., refs (18−21)). LC-MS has been repeatedly and successfully used to map the metabolism and biodistribution of a vast array of metabolites to help explain biological outcomes and metabolomics outputs.²²⁻²⁴ Accessibility and commercial availability of isotopically labeled standards for quantitative measurements have brought LC-MS to the forefront of metabolic analyses of cell, tissue, animal, and human samples. Additionally, molecule-specific isotopologues and isotopomers have been widely implemented as biochemical tracers of metabolic fluxes and biotransformation (e.g.,²⁵⁻²⁷). Their abundance measured by LC-MS, and sometimes LC-MS-MS, informs on the rates at which entities accumulate and are interconverted. These analyses offer information on the overall changes in the abundance of the labeled and nonlabeled metabolites upon exogenous and endogenous stimuli.²⁸

Such a tracing approach requires tailored materials that incorporate enough stable isotopes that enable traceability for flux experiments. A substantial mass shift achieved through chemical or enzymatic incorporation of isotopic atoms into a given molecule is necessary so that this molecule and its metabolites can be detected, assigned, and quantified simultaneously with the naturally occurring species. Furthermore, the isotopologues must retain isobaric properties of the nonlabeled species to be reliably detected under optimized LC-MS conditions.²⁹

To examine by LC-MS how DHA contributes to individual metabolic pathways, including its conversion to F6P/G6P, one can either consider ²H, ¹⁸O, or ¹³C modifications for this simple triose. However, isotopes of oxygen and hydrogen atoms are readily lost during metabolic conversions of sugars and, in some cases, are exchangeable with oxygen and hydrogen atoms present in water.³⁰ A ¹³C-labeled backbone remains traceable throughout the biotransformation and thus can inform the rates at which the labeled material enters a given biosynthetic pathway. Therefore, since the mass difference must be high, the ¹³C uniformly labeled DHA becomes the most obvious isotopologue of DHA.

While isotopic modifications of DHA have been documented¹³ and are commercially available, [¹³C₃]DHA is not so readily available. Its use in metabolic flux analyses has remained limited, even though this triose is likely to modulate cellular homeostasis and central metabolism. The challenges stem from two main parameters. First, the chemical manipulation of this sugar can lead to the generation of byproducts that are difficult to remove.^31,32 Second, the hydroxy-ketone scaffold is perfectly poised to form hemiketal dimers and hydrated species, which are inert to reaction conditions necessary for functionalization. Crucially, each chemical form of DHA, while in reversible equilibria with each other when in solution (Figure 1), can potentially be recognized by sugars’ specific transporters or cross-membranes via small solute channels.^33,34 As a result, in cell-based work exploring the effects of DHA exposure, the DHA payload taken up by cells could differ because of the differential abundance of each DHA form in solution. Relative differences in intracellular DHA levels could affect the rate at which phosphorylation of the monomeric forms of DHA to DHAP by the triose kinase/FMN cyclase (TKFC)³⁵ occurs and ultimately its impact on cellular metabolism and function. Therefore, the latter chemistry points to important differences in the DHA forms that might influence experimental outcomes and account for discrepancies.

Figure 1.

Equilibria between ketonic, hydrated, and dimeric hemiketal forms of DHA in water.

With the perspective of identifying an effective conversion of the commercially available D/L-[¹³C₃]GA (aqueous solution) to the monomeric [¹³C₃]DHA (>90%; aqueous solution), which could be directly applied to cells for DHA tracing studies, we characterized how the relative proportion of dimeric forms of solid DHA changes over time upon storage, and storage conditions since the time of manufacturing, and its influence on the rates at which DHA monomers form once solid DHA dissolves in water as a function of concentration. This knowledge is particularly important when preparing solutions of DHA from commercial sources for use in short-term DHA exposure studies.

Experimental Procedures

Chemistry

DL-glyceraldehyde (powder form) and D/L-(1, 2, 3–¹³C₃) glyceraldehyde (aqueous solution) were purchased from Sigma-Aldrich and BOC Sciences, respectively. Calcium phosphate tribasic or hydroxyapatite (powder form) was purchased from Beantown Chemical. All of the reagents were used without further purification. ¹H NMR and ¹³C NMR spectra were recorded on a Bruker Avance III HD 400 spectrometer (TopSpin 3.5 pl 6) instrument with nominal frequencies of 400.1 MHz for proton and 100.62 MHz for carbon. All of the NMR experiments were performed in H₂O/D₂O (9:1) at 298.2 K. Deuterium oxide (purchased from ACROS Organic) was used for locking and shimming. ¹H NMR spectra are reported in parts per million (ppm) relative to the residual internal reference (H₂O/HOD) or to trimethyl phosphate (TMP, δ 3.82 ppm, d). ¹³C NMR chemical shifts are reported in parts per million (ppm) relative to TMP.

Synthesis of Dihydroxyacetone

Glyceraldehyde (90 mg, 1.00 mmol) and hydroxyapatite (60 mg, 0.12 mmol) were added to water (2 mL) in a 7 mL vial. The reaction mixture was heated in an oil bath up to rt, 25, 50, and 80 °C and stirred for 1, 5, and 25 h. Then, the catalyst was filtered off and the filtrate was analyzed by NMR. Best conversions were obtained after 1 h heating at 80 °C. ¹H NMR (400 MHz, H₂O/D₂O): δ 4.41 (s, 4H; ketone), 3.57 (s, 4H, hydrate). ¹³C NMR (100 MHz, H₂O/D₂O): δ 212.0, 64.8 (ketone), 95.0, 63.6 (hydrate). The chemical formula of the ketone form of DHA: C₃H₆O₃; exact mass: 90.03169.

Synthesis of ¹³C-Label Dihydroxyacetone ([¹³C₃]DHA)

Hydroxyapatite (60 mg, 0.119 mmol) was added to an aqueous solution of D/L-[¹³C₃] glyceraldehyde (D/L-[¹³C₃]GA) (2.5 mL mg, 0.10 M) in a 7 mL vial. The reaction mixture was heated in an oil bath up to 80 °C and stirred for 1 h. Then, the catalyst was filtered off and the filtrate was analyzed by NMR. ¹H NMR (400 MHz, H₂O/D₂O): δ ketone 4.18 (dtd, J = 4, 12, 144 Hz; 4H), hydrate 3.58 (dd, J = 4, 144 Hz); >90% purity measured by ¹H NMR. ¹³C NMR (100 MHz, H₂O/D₂O): δ ketone: 212.02 (t, J = 42 Hz), 64.09 (d, J = 42 Hz); hydrate 95.0 (t, J = 49 Hz), 63.7 (d, J = 48 Hz). Chemical formula: ¹³C₃H₆O₃; exact mass: 93.04176.

Cell Culture

Human embryonic kidney cells containing the SV40 T-antigen (HEK293T) were purchased from ATCC (Manassas, VA). The cells were grown at 37 °C in a 5% CO₂ incubator in Dulbecco’s modified Eagle’s medium (DMEM, Hyclone, Logan, UT; 4.5 g/L glucose and l-glutamine) supplemented with 10% fetal bovine serum (FBS, Atlanta Biologicals, Flowery Branch, GA) and 1% sodium pyruvate (Gibco, Carlsbad, CA). Cell passages between 4 and 15 were used for all reported experiments. Cells were routinely tested for mycoplasma using the Lonza MycoAlert kit (Walkersville, MD) and found to be free of mycoplasma contamination.

Mass Spectrometry Analysis Sample Acquisition

HEK293T cells were seeded at a density of 3.5 × 10⁶ cells/well and incubated overnight (ON) at 37 °C in a 5% CO₂ incubator. The next day, cells were treated with a solution of filtered hydroxyapatite (control), 5 mM D/L-[¹³C₃]GA, or 5 mM [¹³C₃]DHA. The solution and isotopically labeled material were dissolved in the cell culture medium and applied to the cells for 15 min at 37 °C in a 5% CO₂ incubator. At the end of the exposure period, media was aspirated, cells were scraped, and pellets were resuspended in phosphate-buffered saline (PBS, Gibco, Carlsbad, CA) for cell counting. An equal number of cells (6 × 10⁶ cells/condition) were collected in 1.5 mL centrifuge tubes, pelleted, and immediately extracted without storing at −80 °C. Solvents (CHCl₃, MeOH, and H₂O) were degassed, and 200 μL of degassed MeOH was added in each 1.5 mL centrifuge tubes containing the cell pellet and sonicated three times (10 sec each) at 4 °C to rupture the cell wall. After sonication, the sample was incubated at 0 °C for 2 min on ice, followed by the addition of 200 μL MeOH, 400 μL CHCl₃, and 200 μL H₂O and vortexed for 30 sec. The resulting mixture was centrifuged at 13,000 rpm at 4 °C for 15 min. Aqueous and organic phases were separated by centrifugation and were collected separately. Four hundred microliters of the aqueous phase was transferred in a 1 mL centrifuge tube, flash-frozen in liquid nitrogen, and freeze-dried. Freeze-dried samples were reconstituted in 35 μL of 10 mM ammonium acetate, and 6 μL was used for each injection. LC-separation was achieved on an Agilent 1200 series liquid chromatography (LC) with an Agilent Zorbax 300SB (C18, 2.1 × 150 mm) column. Initially, the flow rate was set at 50 μL per minute and ramped up to 150 μL/min in 20 min and held there until the end of the run. Mobile phase A is 10 mM ammonium acetate in 0.1% formic acid (Fluke, NY), and mobile phase B is 0.1% formic acid in methanol (Thermo Fisher Scientific, Waltham, MA). Gradient conditions were 2% mobile phase B initially and increased to 95% in 20 min, held it for 5 min, brought back to 2% in 15 min, and equilibrated for 10 min. LC eluent was introduced to a Thermo LTQ Orbitrap XL Mass spectrometer (Thermo Scientific, San Jose, CA), equipped with electrospray ionization. The spray voltage was set at 3.5 kV, and the capillary voltage and temperature were set at 50 V and 275 °C, respectively. Data were acquired in negative ionization in a full scan mode, and the acquired data were processed in Xcalibur. Two technical replicates were run for each biological repeat. The graph was plotted as the average of the peak values relative to their respective control ± SEM of three biological repeats. Peak areas of isotopically labeled metabolites were corrected to natural abundance.³⁶

Results

In our initial investigation of DHA function in cell-based assays, we purchased solid DHA known as dimer DHA³⁷ to remain consistent with already established protocols. Based on some cell-based work, the laboratory perception was that solid DHA had a relatively short shelf-life and its ability to produce browning of cells by reaction with peptides degraded in a matter of weeks, even when stored cold. However, the rationale for such an opinion was lacking. In solution, DHA is thought to be present as a monomer, either as a ketone and a hydrate, rather than in its dimeric forms, a complex mixture of α/β hemiketals (Figure 1). It is anticipated that monomeric ketonic DHA is the active ingredient in such a mixture. Knowing its abundance in any given solution is therefore highly favorable. When compared to the literature³⁸ and upon inspection by ¹H NMR and ¹³C NMR of aqueous solutions prepared from the solid DHA at concentrations ranging between 10 and 400 mM on multiple occasions, we observed that the proportion of monomeric forms of DHA³⁹ declined steadily upon storage at −20 °C with an increased proportion of the dimeric forms and the more significant contribution of one specific dimeric isomer, especially at higher concentration.

More specifically, we examined by ¹H NMR the commercial dimeric DHA freshly purchased and the same material after an 8 week storage (total triose concentration in H₂O/D₂O; 370 mM). The analysis of the ¹H NMR spectra of the freshly acquired DHA showed that the monomeric forms constituted the main species,³² with 60% of the ketonic form and 14% for the hydrate form (Figure 2, panel A). The dimeric forms of DHA that can be the α/α, α/β, or β/β anomers accounted for 25% of the total DHA composition of monomeric and dimeric species. For the cold-stored DHA, the ¹H NMR spectrum showed that the ketone form was the minor species (20%). Moreover, the hydrated form was no longer observed (Figure 2, panel B), and the dimeric species represented 80% of the composition. These results showed that the commercial dimeric composition of DHA changed over time and, more importantly, presented different composition profiles once dissolved in water. The NMR analysis of a 4 week old sample DHA sold as a monomer purchased from another supplier showed even less monomeric forms (ESI 1). The proportion of the dimeric forms was particularly substantial and accounted for most of the materials present in solution when solid DHA had been stored for more than 4 weeks from its time of purchase (Figure 3).

Figure 2.

¹H NMR of fresh (panel A) and stored (panel B) commercial DHA (110 mM) purchased as a solid, known as DHA dimer. Panel A: commercial solid DHA dimer freshly opened once received from the supplier; panel B: commercial DHA dimer stored for >8 weeks at 4 °C as solid; NMR spectrum taken 5 min after the addition of D₂O. Labeled in red are proposed to be the hydrogens labeled in bold (a and b) in the figure panel A of the β,β and α,α anomeric dimers (enantiomers) based on ¹H and ¹³C NMR observations and J values, and labeled in black are proposed to be the hydrogens labeled in bold (a and b) of the α,β and β,α anomeric dimers (enantiomers).

Figure 3.

Contribution of the monomeric and dimeric forms of DHA in solution.

When the samples were analyzed by ¹³C NMR, the relative differences in composition confirmed the increase of the dimeric contribution to the composition of solid DHA occurring over time. However, we also observed that the relative intensity of the peaks representative of the monomeric forms (ketone and hydrate) was more substantial than what the ¹H NMR would have suggested. Indeed, we could observe the hydrated monomer in the ¹³C NMR spectrum of the stored DHA, although it was not observed in the ¹H NMR spectrum. We concluded that the length of the acquisition time for a ¹³C NMR spectrum compared to that of the ¹H NMR (∼1 h vs 1 min) provided time for the various forms of DHA to equilibrate when in solution.

Following this observation, we decided to monitor the relative abundance of the monomers and dimers of the 8 week old solid DHA dimer kept at rt while in solution. We observed that after 48 h, the monomeric forms of DHA represented more than 94% of DHA in solution and that this composition remained constant over days while kept at rt (ESI 2). Using a fresh batch of solid DHA, we also observed that the same equilibrium could be reached after 12 h at rt in water (Figure 4, panel A). We evaluated how fast the dimeric species present in a 110 mM DHA-H₂O/D₂O solution could convert to the monomeric species. Equilibrium was reached within 12 h (Figure 4, panel B). Further, in a serial dilution experiment, we investigated the effect of total DHA concentration on the isomeric distribution (Figure 5). At rt, we observed that dimeric forms are detectable at concentrations as low as 27 mM, while the ketone and hydrated forms were most abundant at lower concentrations (e.g., 3 mM).

Figure 4.

Equilibrium between the dimeric and monomeric forms of DHA in solution over time. Panel A: ¹H NMR monitoring; panel B: graphic representation of the DHA composition in water. Relative % proportion of the four forms of DHA as a function of time in solution.

Figure 5.

¹H NMR of a serial dilution of solid DHA solution. Dimeric forms of DHA are preferred at high mM concentration, while the monomeric forms of DHA are observed at low mM concentrations.

While it is acknowledged that lower storage temperatures (e.g., −80 °C) are likely to slow down the dimerization process in solid DHA, thawing and freezing will promote it. Therefore, the use of disposable DHA aliquots is highly recommended. Furthermore, unless confirmation of the dimeric to monomeric distribution is checked upon reception of the commercial DHA, it should not be assumed that DHA isomeric distribution is always the same, as this will vary with the length of storage and storage conditions at the manufacturer site.

These relative proportions of isomers in solution and isomerization rates are particularly relevant when preparing stock solutions of DHA for use in cell or animal studies. The relative abundance of each isomer at the time of exposure is dependent on the initial concentration of DHA (bolus; 80–200 mM^8,13,15 vs diluted applications; 1–10 mM⁹). While work to establish the nature of this equilibrium and the rates at which equilibrium is reached in cell culture media is underway, we sought to directly use DHA as a monomer in solution to avoid the potential need for serial dilutions of dimeric DHA solutions and equilibration in cultured media. Preparing DHA solutions kept at room temperatures in advance of use (ca. 12 h) on cells minimizes isomers’ distribution discrepancies between experiments.

Since we observed DHA has an IC_90DHA value of 5 mM in HEK293T,⁹ we wanted to conduct flux experiments comparing the conversion of monomeric DHA, the substrate of TKFC, to DHAP and its incorporation in metabolic pathways to that of GA in HEK293T cells using mass spectrometry (Figure 6). Therefore, we needed to generate the traceable [¹³C₃]DHA isotopologue in its monomeric form. The D/L-[¹³C₃]GA was commercially available, while the [2-¹³C]DHA was the only available ¹³C-isotopologue of DHA. The increased number of ¹³C isotopic carbons on D/L-GA means its tracing would offer more reliable measurements than the monolabeled DHA. An increased number of isotopic atoms introduced in a molecule ensure that the molecules incorporating isotopes and derived from the isotopically labeled tracer remain detectable at levels above the levels of natural isotopic abundance. For example, fructose 6-phosphate and glucose 6-phosphate (F6P/G6P; C₆H₁₃O₉P) have an exact mass of 260.02977 g/mol for an isotopic abundance of 100% when incorporating ¹H, ¹²C, and ¹⁶O. Along with this most abundant form of F6P/G6P are the isotopologues of F6P/G6P that contain one ¹³C atom. These species occur in 6.5%, with an exact mass of 261.03313 due to the natural abundance of ¹³C atom (1.07%) and the possibility of having any of the six ¹²C carbons of F6P/G6P replaced by a ¹³C atom (all respective isotopomers with identical molecular weight). Any labeled species metabolized to F6P/G6P will need to contribute to levels substantially above this 6.5% abundance to be measured in a statistically reliable manner if they only incorporate one isotope of 1 atomic unit (au) difference. The F6P/G6P masses that incorporate a higher number of isotopes or an isotope with higher atomic mass (e.g., ¹⁸O) are much less abundant (262.03xx natural abundance 2% and 263.03xx, natural abundance 0.1%) and therefore are more amenable for detection above the levels of natural abundance. We, therefore, decided to generate [¹³C₃]DHA from D/L-[¹³C₃]GA with as little chemical handling as possible while maintaining good yields and levels of purity with no requirements for product purification, using published literature. We examined the literature for simple one-pot conversions from glyceraldehyde rather than from glycerol.⁴⁰ In 2019, Usami reported the conversion of GA to DHA using hydroxyapatite in good yields,⁴¹ whereby the conversion was conducted at 80 °C for 25 h. Thus, we started with this approach.

Figure 6.

Synthetic preparation of monomeric [¹³C₃]DHA from D/L-[¹³C₃]GA, and metabolic tracing of D/L-[¹³C₃]GA and [¹³C₃]DHA in HEK293T cells upon 15 min exposure.

In contrast to DHA, commercial D/L-GA is mainly present in its hydrated form as a monomer (Figure 7, bottom NMR) and remains constant upon storage. Upon adding hydroxyapatite to a solution of D/L-GA and examining the reaction mixture by ¹H and ¹³C NMR after 25 h at 80 °C (ESI 3), we observed DHA formation; however, we also noticed the accumulation of byproducts other than the DHA monomeric or dimeric forms or GA contaminants. Following the optimization of the temperature and initial reduction of the reaction time to increase conversion while decreasing side-product formation, we monitored the reaction at 80 °C over a shorter time (e.g., 15, 30, and 45 min) and observed improved conversion of GA to DHA, with the reaction appearing near completion and with less byproduct after 45 min (Figure 7). These results indicate a rapid isomerization of GA to DHA at 80 °C (ESI 4), with a need to keep the reaction time short to minimize the formation of byproducts, including DHA dimers (Figure 7).

Figure 7.

Rapid conversion of GA to DHA with hydroxyapatite in water at 80 °C monitored by NMR.

Although yielding DHA monomers contaminated with some minor components other than the dimers, this chemistry appeared well suited for generating the [¹³C₃]DHA from D/L-[¹³C₃]GA. To ensure the complete conversion of D/L-[¹³C₃]GA to [¹³C₃]DHA, we conducted the reaction with a hydroxyapatite suspension at 80 °C for 1 h using an aqueous solution of isotopically ¹³C₃-labeled D/L-GA. ¹H and ¹³C NMR readily monitored the reaction. The ¹H and ¹³C NMR spectra show multiplicity consistent with the anticipated coupling constant between ¹³C carbons (>99% abundance) and ¹H hydrogens. Furthermore, ¹³C–¹³C coupling between the C1/3 and C2 carbons of DHA is observed in its ketonic or hydrated form. Complete conversion of D/L-[¹³C₃]GA to [¹³C₃]DHA was confirmed (Figure 8) after 1 h. Notably, after 1 h at 80 °C, [¹³C₃]DHA was mainly obtained as a monomer, with the α/α-β/β dimeric form of DHA being the predominant dimer with a relative abundance consistent with that observed in equilibrated DHA solutions (Figure 4). The DHA monomers accounted for ∼73% of the trioses, with the remainder accounted for by the dimers. Unfortunately, the complex coupling patterns rendered it impossible to accurately quantitate these species (ESI 5). Importantly, little byproducts were observed. Upon reaction completion, the catalyst was removed by filtration. To avoid the possibility of DHA dimerizing upon the removal of water under reduced pressure, the filtered crude reaction solution was used directly in the cell culture experiments to evaluate whether GA and DHA entered the glycolysis pathway and were converted to F6P/G6P to the same extent.

Figure 8.

¹H and ¹³C NMR spectra monitoring the conversion of D/L-[¹³C₃]GA to [¹³C₃]DHA at 80 °C in the presence of hydroxyapatite in water.

An equal volume of an aqueous solution of D/L-[¹³C₃]GA (5 mM), [¹³C₃]DHA (5 mM), or a solution of filtered hydroxyapatite (control) was applied to human embryonic kidney cells, HEK293T. Following a 15 min incubation period, the cells were harvested. The cell pellets were extracted for metabolomic evaluation of the D/L-[¹³C₃]GA vs [¹³C₃]DHA incorporation in the glycolytic and glycogenesis pathways. The conversion of each ¹³C-labeled triose into isotopically labeled lactic acid (LA) and F6P/G6P was measured by quantifying the relative increase in the proportion of each species for which the molecular weight had increased by 3 or 6 atomic units (MW+3 or MW+6), respectively. An MW shift by 3au in F6P/G6P indicates the incorporation of one unit of ¹³C₃-labeled GA or DHA, while a shift by 6au indicates that two GA or DHA contributed to the hexose formation. While both GA and DHA contribute to the G6P/F6P production, we consistently observed that only one triose unit was incorporated over 15 min of incubation. Further, we observed the formation of ¹³C₃-labeled DHAP/GAP when the cells were exposed to [¹³C₃]DHA, but we did not observe any ¹³C-labeled DHAP/GAP when the cells were exposed to D/L-[¹³C₃]GA (Figure 9).

Figure 9.

Relative proportion of labeled (+3au) and nonlabeled isotopologues of the lactic acid (LA), glyceraldehyde 3-phosphate/dihydroxyacetone phosphate (GAP/DHAP), and glucose 6-phosphate/fructose 6-phosphate (G6P/F6P) pools in HEK293T cell extracts following a 15 min incubation with either water, D/L-[¹³C₃]GA, or [¹³C₃]DHA.

Furthermore, we also observed that both GA and DHA contribute to the formation of LA. As for the generation of G6P/F6P, the [¹³C₃]LA synthesis from the [¹³C₃]DHA unit is delayed compared to that from the D/L-[¹³C₃]GA precursor (Figure 9). Crucially, when measuring the fold changes in LA, GAP/DHAP, and G6P/F6P following the 15 min cell exposure used here, we observed that DHA promoted the sharp production of LA, while GA promoted a more substantial increase in F6P/G6P than DHA (Figure 10). However, both trioses increase the overall levels of the two hexoses. Surprisingly, the total levels of DHAP and GAP decreased upon exposure to DHA and GA (Figure 10).

Figure 10.

Fold changes in the abundance of total lactate, glyceraldehyde 3-phosphate/dihydroxyacetone phosphate (GAP/DHAP), and glucose 6-phosphate/fructose 6-phosphate (G6P/F6P) upon exposure to 5 mM D/L-[¹³C₃]GA or [¹³C₃]DHA.

Discussion

Using NMR spectroscopy, we have characterized how the dimeric and monomeric forms of DHA equilibrate once in aqueous solutions and demonstrated that with sufficient time the dimeric forms of DHA, including the most predominant dimeric forms of DHA (the enantiomers α/α and β/β), are converted to the hydrate and ketonic form of DHA. Temperature, concentration, length of incubation, and solution type affect this equilibrium. This should be considered when exploring DHA biology since it is assumed that it is in its ketone form that DHA enters metabolic pathways in cells. Short-term experiments will be most affected by this slow equilibrium.

We successfully generated [¹³C₃]DHA from D/L-[¹³C₃]GA using a simple method that mainly leads to monomeric DHA forms that can then be used for short-time exposure experiments on cells. Crucially, we now know that a dilute solution of DHA will favor monomeric forms of DHA and demonstrated substrates of TKFC. In diluted solutions, the monomeric forms remain stable for days at rt. Such stock solutions can then be used over several experiments with little deviation as to the chemical form of the DHA applied. This observation is valid for labeled and nonlabeled DHA. Using this labeled DHA solution, we demonstrated that the monomeric forms of DHA are taken up by cells and rapidly phosphorylated.

Furthermore, we observed the formation of ¹³C-labeled DHAP/GAP in cells exposed to [¹³C₃]DHA but not to D/L-[¹³C₃]GA. These data demonstrate the rapid kinetics of GAPDH and suggest that the MW+3 observed for GAP/DHAP is due to [¹³C₃]DHA generated when the cells are exposed to [¹³C₃]DHA (Figure 8) and confirm that the triosephosphate isomerase (TPI) is a rate-limiting enzyme in the conversion of DHAP to GAP in cells. The unexpected decrease in overall triose levels upon exposure to DHA or GA could be explained by activating metabolic pathways that seek to redress the increased levels of exogenous trioses. Defects in the enzymatic activity of TPI or TKFC have been associated with metabolic disorders in humans^42,43 and could lead to the accumulation of nefarious intermediates. These results further support the need for careful regulation of triose levels within the cell.

Overall, using D/L-[¹³C₃]GA and [¹³C₃]DHA, we have shown that DHA is converted to triose phosphates and incorporated into hexose phosphates at rates that differ from GA. Further, we showed that exogenous GA and DHA affect lactate, triose phosphates, and hexose phosphate levels differently but substantially. DHA and GA contribute to multiple biosynthetic pathways once converted to their phosphate parent, including the substantial formation of hexose phosphates, G6P and F6P. DHA appears to have a delayed metabolism compared to GA. Therefore, this warrants further investigation since these differences are likely to provide some insights into the pathology of DHA exposure.

Unlike ¹³C NMR-enabled metabolomics and flux monitoring, either using standard NMR or hyperpolarized spectroscopy, mass spectrometry, and use of isotopologues allow for the evaluation of not only the fate of the tracers used in the experiment but also that of their impact on other metabolites that do not necessarily incorporate a traceable label but that nonetheless get modified by exposure. In this respect, the use of mass spectrometry in the flux experiment of ¹³C-labeled trioses offers a bigger picture of the effect of DHA exposure on cellular metabolism while providing means for establishing the fate of this specific metabolite. By combining the comprehensive information gathered by metabolomic NMR spectrometry experiments to mass spectrometry to help generate compounds’ panels for targeted analyses, one will establish the distribution and impact of accumulated DHA metabolites following DHA exposure on cells and in tissues.

Finally, it appears that the generation of diluted stock solutions of DHA (<25 mM) could offer a more consistent use of DHA in its monomeric species rather than storing solid DHA and generating DHA solution shortly before use, especially at high concentration. Crucially, this now raises the possibility that aerosolized (spray-tanning products) and vaporized (e-cigarettes) DHA might reach their targets with a different composition of monomers and dimers, which could affect their toxicity differently.

Conclusions

We have demonstrated that the commercial dimer DHA slowly equilibrates to its monomeric ketone and hydrate form in aqueous solution at room temperature and optimized the synthesis of monomeric DHA from GA by a simple process. This process minimizes dimeric species and other degradation products and allows for the evaluation of the monomeric species, independent of the relative distribution of DHA isomers present in the commercial source at the time of use. This synthesis can be applied in any setting and allows the rapid conversion of D/L-[¹³C₃]GA to [¹³C₃]DHA. These isotopologues were then used to demonstrate how GA and DHA affect the central carbohydrate metabolism.

Glossary

Abbreviations

DHA

dihydroxyacetone

GA

glyceraldehyde

DHAP

dihydroxyacetone phosphate

GAP

3-phosphoglyceraldehyde

F6P

fructose 6-phosphate

G6P

glucose 6-phosphate

LA

lactic acid/lactate

TPI

triosephosphate isomerase

Supporting Information Available

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acs.chemrestox.1c00403.

• ¹H and ¹³C NMR showing the equilibration of old and fresh solid DHA in water after 24 h (ESI 1); monitoring of the stability of DHA in water over 1 month and 24 days by ¹H NMR (ESI 2); monitoring of the synthesis of DHA from GA according to Usami conditions (1–25 h at 80 °C) (ESI 3); conversion rates of GA to DHA up to 1 h (ESI 4); and ¹H NMR spectrum and conversion yields of D/L[¹³C₃]GA to [¹³C₃]DHA after 60 min (ESI 5) (PDF)

Author Contributions

L.B.: data acquisition and curation, formal analyses, investigation, methodology, resources, and writing (review and editing). M.S./A.H.: investigation, resources, and writing (review and editing). N.R.G.: conceptualization, funding acquisition, resources, supervision, and writing (review and editing). M.E.M.: conceptualization, data curation, formal analyses, funding acquisition, resources, supervision, and writing (original draft).

NIH funding NIEHS grant R01 ES032450 supported this project.

The authors declare no competing financial interest.