Chemical and Biochemical Reactivity of the Reduced Forms of Nicotinamide Riboside

Article abstract of DOI:10.1021/acschembio.0c00757

All life forms require nicotinamide adenine dinucleotide, NAD+, and its reduced form NADH. They are redox partners in hundreds of cellular enzymatic reactions. Changes in the intracellular levels of total NAD (NAD+ + NADH) and the (NAD+/NADH) ratio can cause cellular dysfunction. When not present in protein complexes, NADH and its phosphorylated form NADPH degrade through intricate mechanisms. Replenishment of a declining total NAD pool can be achieved with biosynthetic precursors that include one of the reduced forms of nicotinamide riboside (NR+), NRH. NRH, like NADH and NADPH, is prone to degradation via oxidation, hydration, and isomerization and, as such, is an excellent model compound to rationalize the nonenzymatic metabolism of NAD(P)H in a biological context. Here, we report on the stability of NRH and its propensity to isomerize and irreversibly degrade. We also report the preparation of two of its naturally occurring isomers, their chemical stability, their reactivity toward NRH-processing enzymes, and their cell-specific cytotoxicity. Furthermore, we identify a mechanism by which NRH degradation causes covalent peptide modifications, a process that could expose a novel type of NADH-protein modifications and correlate NADH accumulation with protein aging. This work highlights the current limitations in detecting NADH's endogenous catabolites and in establishing the capacity for inducing cellular dysfunction.

Full text of DOI:10.1021/acschembio.0c00757

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Chemical and Biochemical Reactivity of the Reduced Forms of
Nicotinamide Riboside
Mikhail V. Makarov,† Faisal Hayat,† Briley Graves, Manoj Sonavane, Edward A. Salter,
Andrzej Wierzbicki, Natalie R. Gassman, and Marie E. Migaud*
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ABSTRACT: All life forms require nicotinamide adenine
dinucleotide, NAD+, and its reduced form NADH. They are
redox partners in hundreds of cellular enzymatic reactions.
Changes in the intracellular levels of total NAD (NAD+
+
NADH) and the (NAD+/NADH) ratio can cause cellular
dysfunction. When not present in protein complexes, NADH and
its phosphorylated form NADPH degrade through intricate
mechanisms. Replenishment of a declining total NAD pool can
be achieved with biosynthetic precursors that include one of the
reduced forms of nicotinamide riboside (NR+), NRH. NRH, like
NADH and NADPH, is prone to degradation via oxidation, hydration, and isomerization and, as such, is an excellent model
compound to rationalize the nonenzymatic metabolism of NAD(P)H in a biological context. Here, we report on the stability of
NRH and its propensity to isomerize and irreversibly degrade. We also report the preparation of two of its naturally occurring
isomers, their chemical stability, their reactivity toward NRH-processing enzymes, and their cell-specific cytotoxicity. Furthermore,
we identify a mechanism by which NRH degradation causes covalent peptide modifications, a process that could expose a novel type
of NADH-protein modifications and correlate NADH accumulation with “protein aging.” This work highlights the current
limitations in detecting NADH’s endogenous catabolites and in establishing the capacity for inducing cellular dysfunction.
a humid environment, 1,4-NADH powder isomerizes to the
1,2- and 1,6-NADH isomers.23,24 On average, a commercial
source of NADH will contain up to 4% of these isomers if
stored under nonanhydrous conditions for extended periods of
time. The identity of these isomers was confirmed when 1,2-
and 1,6-NAD(P)H were generated by reducing NAD(P)+
using NaBH4 in aqueous media.25 In solid-form storage, the
1,2- and 1,6-isomers of NAD(P)H have been proposed to form
through a bimolecular reaction in which an apparent double
bond rearrangement that lacks chemo-selectivity occurs via a
hydride transfer from NAD(P)H to NAD(P)+.23 A particularly
well-suited biochemical environment for this mismatched
hydride transfer and endogenous production of isomers is
the mitochondrial NAD(P)+ transhydrogenase (nicotinamide
nucleotide transhydrogenase; NNT), which is responsible for
the generation of NADPH and NAD+ from NADP+ and
NADH in the mitochondrion.26 However, this suggestion is
speculative as the mechanism of NNT remains under intense
INTRODUCTION
■
Nicotinamide adenine dinucleotide (NAD+), along with its
phosphorylated and reduced forms (NAD(P)(H)), is a vitamin
B3-containing dinucleotide (Figure 1). Together, the NAD-
(P)(H) cofactors catalyze most known enzymatic hydride
transfers.1−5 Unlike NAD+, NADH (Figure 1) is not a
substrate for nonredox NAD+ consuming enzymes, such as
sirtuins, adenosine diphosphoribosyl transferases (ARTs), and
glycohydrolases.6−9 Critically, it has been observed that in the
process of aging and several metabolic disease models (e.g.,
type II diabetes, cardiovascular diseases, cardiomyopathy, etc.),
the NADH to NAD+ ratio increases, with, on occasion, an
overall decrease of the (NADH + NAD+) pool.8,10−14
1,4-NAD(P)H, more commonly known as NAD(P)H, is
most conspicuous for its redox capacity in enzyme-catalyzed
reactions. The 1,4-nomenclature in 1,4-NAD(P)H indicates
the position of the double bonds in the dihydronicotinamide
moiety. While it is common knowledge that 1,4-NAD(P)H
gets readily oxidized when exposed to air in the presence of
moisture,15−17 the fact that NAD(P)H is also prone to
isomerization and hydroxylation (Figure 2) under these
conditions is less known. The products generated by this
class of chemistry are the 1,2- and 1,6-isomers of NAD(P)-
H18,19 and the 6-OH-NAD(P)H,20−22 a compound known in
the literature as NAD(P)HX (e.g., Figure 2; NADH(OH)). In
Received: September 24, 2020
Accepted: March 16, 2021
Published: March 30, 2021
© 2021 The Authors. Published by
American Chemical Society
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Figure 1. Ribosylated precursors to NAD(P)(H), the associated metabolic and NAD+ consuming enzymes, and redox equilibrium. AK, adenosine
kinase; NRK, nicotinamide riboside kinase; NMNAT, nicotinamide mononucleotide adenylyltransferase; NADK, NAD kinase; ARTs, adenosine
diphosphoribosyl transferases.
lated species back to NAD(P)+. Both enzymes are critical to
cellular homeostasis and lethal if absent.32,34 This speaks to the
physiological relevance of these NAD(P)H isomers.
Another critical point is that since these derivatives are
chemically formed and degraded within the cells’ compart-
ments where they are generated, the equilibrium between these
derivatives and 1,4-NAD(P)H is likely to be permanently
shifted away from the functional 1,4-NAD(P)H forms. As a
result and over time, the damage these derivatives
endogenously create could be much more severe than a simple
chemical ratio might indicate.35 Therefore, the complexity of
the solution chemistry of the reduced forms of NAD(P)+ and
derivatives is likely to explain why these derivatives have not
received their due attention in cell/tissues/organism-based
assays, even though physiologically critical repair pathways are
in place to prevent their accumulation.
Our inability to detect the presence of toxic NAD(P)H
derivatives and quantify their relative abundance limits our
efforts in ascertaining their physiological relevance and their
role in cellular homeostasis and disease. Moreover, character-
izing the correlation between abundance and function requires
the challenging task of manipulating their intracellular levels in
a physiologically meaningful manner. Being phosphorylated
species, isomers and hydroxylated derivatives of NAD(P)H
cannot be supplemented to cells directly with the expectation
that they will be intracellularized intact.
To address these limitations, we sought to understand better
the properties of 1,4-dihydronicotinamide riboside (Figure 1;
1,4-NRH, 1) as a model compound for NAD(P)H and explore
its chemical and biochemical space and that of its less common
metabolites, the 1,2-NRH (2) and 1,6-NRH (3) isomers and
the hydroxylated parents 4 (Figure 2). NRH 1 (Figure 1) has
been recently characterized as an effective precursor of
NAD+.36−38 While the conversion of exogenous NRH to
NMNH by adenosine kinase rather than its oxidation to
nicotinamide riboside by NRH:quinone oxidoreductase 2
Figure 2. Nonenzymatic routes to the derivatives of 1,4-NRH and
NADH.
scrutiny. Finally, direct hydroxylation at the 6-position of the
dihydropyridinyl ring of NAD(P)H generates NAD(P)H(OH)
and occurs preferentially under mildly acidic phosphate buffer
(pH 6.8) conditions.22,20
Together, these isomers and adducts are potent inhibitors of
NAD(P)(H) dependent redox enzymes with Ki ranging from
micromolar to low nanomolar values,24,27,28 and they
contribute to the degradation processes of NAD(P)H.29 Still,
the actual intracellular abundance of 1,2- and 1,6-isomers of
NAD(P)H remains unknown, as is that of NAD(P)H(OH).
One could assume that given the relatively low abundance of
the 1,2- and 1,6-NAD(P)H isomers compared to the 1,4-
parents, their physiological significance should be dismissed.
Yet, these isomers are so detrimental to cellular homeostasis
that their removal evokes specific conserved repair path-
ways.18−20,30−32 Two different classes of enzymes using either
FAD (renalase) or FMN (pyridoxamine-phosphate oxidase
and pyridoxamine-phosphate oxidase-related proteins) as a
cofactor remove damaged forms of NAD(P)H as they oxidize
1,2- and 1,6-NAD(P)H to NAD(P)+.30,33 Similarly, the
epimerase/dehydratase repair pathways convert the hydroxy-
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Scheme 1. Synthetic Sequence to 1,4-NRH and Its 1,2- and 1,6-Isomers
(NQO2) is thought to be the first step toward its conversion
to NAD,36,39 the biosynthetic mechanisms explaining this
process remain under scrutiny.40−42 Critically, the physio-
logical prevalence of intracellular NRH on the total NAD pool
maintenance remains to be explored.
Here, we report on the stability of NRH and its propensity
to isomerize and irreversibly degrade. We also report the
preparation of two of its naturally occurring isomers, their
chemical stability, their reactivity toward NRH-processing
enzymes, and their cytotoxicity when administered exoge-
nously. Finally, we identify a mechanism by which NRH
degradation causes peptidic conjugation via the aglycone unit
of NRH, a mechanism that differs from the well-studied ribose-
derived glycation processes.
riboside, NR+ (Figure 2, 5), and the formation of NR+’s
hydrolytic products, nicotinamide (Nam) and ribose. Rather,
1,4-NRH decomposes as a complex mixture of materials, which
varies upon the conditions under which this decomposition
takes place (Figure 2). Consequently, when used for biological
work, 1,4-NRH must be handled with care; otherwise, its use
brings discrepancies to the studies’ outcomes. Therefore, we
sought to better characterize the conditions under which NRH
decomposes and its decomposition products.
Production and Handling of 1,2- and 1,6-NRH. With
the view of better characterizing the 1,2- and the 1,6-isomers of
NRH, we then endeavored to synthesize these species from the
nicotinamide riboside triacetate (6). Unlike the dithionite-
catalyzed reduction of NR+, reduction of NR+ salts with
NaBH445 yields a mixture of 1,2; 1,4; and 1,6-NRH (Scheme 1;
2, 1, and 3) that is difficult to separate. The proportion of
isomers thus generated varies greatly as a function of solvents,
reagent equivalency, and temperature. Therefore, conditions
that offered the highest levels of 1,2- and 1,6-isomers were
sought. In the absence of a catalyst, the generation of 1,2-, 1,4-,
and 1,6-dihydropyridines from N-alkylated nicotinamide salts
is best achieved by using anhydrous DMF at 0 °C (Scheme
1).46 We applied this approach to the triacetate-protected NR-
OTf (6) to enable the purification of the reduction products by
silica or alumina chromatography. Following the NaBH4
reduction in DMF and an extensive extraction sequence, a
solid foam containing a mixture of the triacetylated form of
1,2-NRH (8, 1,2-NRH TA), 1,4-NRH (7, 1,4-NRH TA), and
1,6-NRH (9, 1,6-NRH TA) in a respective ratio of
1.00:0.44:1.00 (67% yield) was obtained. To isolate the 1,2-
and 1,6-isomers of NRH TA, the mixture was subjected to
silica gel column chromatography using EtOH/EtOAc as the
eluent system. While 1,2-triacetate NRH (8) eluted first, the
1,6-NRH triacetate (9) was extremely difficult to separate from
the 1,4-NRH triacetate (7). Following multiple purification
iterations, each isomer was isolated independently. However,
the purification process consistently led to a loss of material,
RESULTS
■
Production and Handling of 1,4-NRH. As previously
described, 1,4-NRH is readily generated from nicotinamide
and tetraacetate riboside.43 We applied mechanochemical
conditions to the Vorbruggen glycosylation of nicotinamide
̈
with the tetraacetate riboside to generate nicotinamide riboside
triacetate (Scheme 1, 6) as the synthetic intermediate to the
NRH series. 1,4-NRH 1 is generated by reducing intermediate
6 using sodium dithionite under aqueous conditions in a
biphasic system to allow the facile extraction of the
triacetylated intermediate 7. This step is followed by the
removal of the acetate moieties under methanolic potassium
carbonate conditions using the molar equivalency of methanol
under milling conditions.44 When in nondeoxygenated
aqueous solutions, 1,4-NRH (1) decomposes over time.
Therefore, NRH requires careful handling when in solution.
Importantly, 1,4-NRH oxidizes upon storage, even at −80 °C,
when stored in aqueous solutions. Freeze−thaws are
particularly discouraged. As per NADH, UV spectroscopy
measurements at 340 nm of 1,4-NRH solutions are particularly
well-suited to monitor whether the 1,4-NRH integrity is
maintained.29 However, the loss of 1,4-NRH is not due to
simple oxidation of NRH to its oxidized form, nicotinamide
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Figure 3. Decomposition of 1,4-NRH in aqueous solution over time. 1H NMR of a 1,4-NRH solution in phosphate buffer, 50 mM, pH 6.8 after 1,
3, 12, and 24 h at 37 °C.
Figure 4. Stability of 1,4-NRH in aqueous solutions. Panel A: 1H NMR of a 1,4-NRH solution in D2O after 24 h at 37 °C. Panel B: 1H NMR of a
1,4-NRH solution in phosphate buffer, 500 mM, pH 6.0 after 30 min at 37 °C.
with the best recovery achieved in 35% total yield. Attempts to
conduct this purification on alumina failed.
In summary, 1,2-NRH and 1,6-NRH triacetate are extremely
labile entities that readily react with either ethanol or water or
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with the solid support itself throughout the purification
process. While the trans-esterification process could be viewed
as a reasonable mechanism leading to the loss of materials, 1,4-
NRH triacetate 7 generated from the dithionite-method is
readily purified using this chromatography method, and the
recovery yields are good to excellent. The reactivity of 1,2-
NRH and 1,6-NRH triacetate (8 and 9) toward mild reagents
such as those employed in the purification process is indicative
of the difficulties that one encounters in seeking to isolate,
purify, or even detect derivatives of such nucleosides in cell
extracts. Yet, having generated and purified the triacetylated
form of 1,2-NRH and 1,6-NRH, careful deprotection was then
undertaken using NaOMe in methanol at low temperatures
(Scheme 1). Each fully deprotected isomer was then isolated,
but no further purification could be performed without
product decomposition. Characterization of each isomer was
conducted using COSY, HMQC, and MS and HRMS data,
while the LC-MS2 fragmentation pattern for each one of these
isomers was characterized.
Stability of 1,4-NRH in Culture Media. Having each
isomer at hand, we then examined the chemical stability of 1,4-
NRH when in solution in cell growth media containing a high
purity 10% FBS and established how resilient the reduced
nucleoside was to oxidation, isomerization, and hydrolysis
under cell-based assay conditions. As anticipated, 1,4-NRH
slowly converted to NR+ and Nam over a 48 h period when
incubated in standard nonheat-inactivated FBS-containing cell-
growth media at 37 °C and 5% CO2, standard cell-based assay
conditions. Importantly, in the absence of cells, 1,4-NRH was
removed from solution through protein interactions (no longer
detectable by 1H NMR), with only 80% of the initial 1,4-NRH
remaining in solution after 6 h. Longer incubation periods
resulted in the production of Nam through NR+ hydrolysis and
further reduction of the total NRH pool. This observation
indicates that when in the presence of cells, prolonged
supplementation with 1,4-NRH must be monitored to assign
physiological outcomes to the interaction of NRH with the
cells, rather than with the growth media and NRH byproducts.
Critically, when FBS is supplemented with 10 mM 1,4-NRH,
the 1H NMR spectrum indicates the formation of the 1,6-NRH
isomer, in proportions similar to those as observed in PBS. The
stability of 1,2-NRH and 1,6-NRH in the same growth media
was thus investigated. While 1,6-NRH proved relatively stable
in the growth media for up to 6 h, the 1,2-NRH isomer
decomposed in a matter of hours.
Stability of 1,4-NRH in Aqueous Media. The broader
stability of 1,4-NRH as a function of pH and temperature was
systematically investigated using 1H NMR spectroscopy
(Figures 3 and 4). Because NRH is used under conditions
where the temperature is maintained at 37 °C, either in cell-
based assays or animal studies, we discuss here the results
observed at 37 °C. First, the mild conditions developed by
Margolis to generate the hydroxylated forms of NAD(P)H
were adapted and applied to 1,4-NRH (50 mM KH2PO4, pH
6.8, 37 °C) and progression monitored by 1H NMR at 1, 3, 12,
Cytotoxicity of 1,2-, 1,4-, and 1,6-NRH. As 1,4-NRH is a
precursor to 1,4-NAD(P)(H),38 1,2- and 1,6-NRH are putative
precursors of 1,2- and 1,6-NAD(P)H, respectively. If this is the
case, the formation of such isomers of NAD(P)H would be
detrimental to cells.30 Therefore, to evaluate whether 1,2-
NRH, 1,4-NRH, and 1,6-NRH could modulate cellular
function, we evaluated their effect as cytotoxic agents using
the Human Embryonic Kidney (HEK293T) and Hepatoma
(HepG3) immortalized cell lines as cell-based models. The
cytotoxicity of 100 μM 1,2-NRH, 1,4-NRH, and 1,6-NRH was
determined by CellTiter-Fluor (Promega), which unlike the
standard and widely used MTT assay47 measures cell viability
independent of metabolic markers related to NAD(P)(H).
The cells were exposed to the NRH isomers for 6 h, consistent
with our stability studies. The 6 h media did not contain
1
and 24 h. The H NMR spectra (Figure 3) provided evidence
that 1,4-NRH oxidizes to NR+ in addition to showing the
formation of new species including one that matched the
hydrated form of NRH NRH(OH) (Figure 2, 4, and Figure 4)
with characteristic peaks at 7.35 and 5.64 ppm. The 1,6-NRH
isomer could also be detected with characteristic peaks
observed at 5.03, 6.10, and 7.24 ppm after just 3 h, consistent
with what is known for NADH multicomponent composi-
tions.23,24,44 Crucially, the carbohydrate and aliphatic regions
became a lot more complex. The aliphatic region (1−2.5 ppm)
reveals a −(CH2−CH2−CH)− sequence that can be attributed
to hydrated catabolites described in Figure 4. It must be noted
that only the 6-hydroxylated byproduct is shown in Figure 4,
even though the 2-hydroxylated byproducts can also be
envisaged as contributing to the mixture. Crucially, this
chemistry is not observed to occur that rapidly in water, as
when NRH was kept in water for 24 h at 37 °C, mainly NR+
formation was observed (NRH/NR+ = 454:1; Figure 4, panel
A). At lower pH, pH 6.0, and in 500 mM PBS buffer, the
decomposition of 1,4-NRH is even more rapid as evidenced by
1
residual NRH isomers (confirmed by H NMR), indicating
that each isomer was either transported into the growing cells
or was bound to media proteins or degraded and thus no
1
longer detectable by H NMR. The cell survival after 6 h
exposure to NRH isomers is shown in Figure 5. At this
concentration and over this 6 h incubation period, the 1,4- and
1
the H NMR after just 30 min of incubation (Figure 4, panel
B).
Reactivity of 1,4-NRH with NR+ in Aqueous Media.
We then investigated whether 1,4-NRH when in the presence
of the oxidized form NR+ could isomerize to 1,2-NRH (Figure
2; 2) and 1,6-NRH (Figure 2; 3). 1,4-NRH was coincubated
with NR+ in D2O for 100 h at room temperature. Under these
conditions, the 1,6-NRH isomer accounted for 1 in 20 mol of
(NRH+NR+), but the 1,2-NRH was not detectable (exper-
imental materials, ESI-NMR1). Of note, the hydrated species
were not observed; in particular, the aliphatic region remained
free of signals even after 100 h of incubation in water. The salt
composition and pH of the aqueous solution appear to be the
limiting factors leading to the hydroxylation of 1,4-NRH.
Figure 5. Cytotoxicity of NRH derivatives (100 μM) on HEK293T
and HepG3 cell lines. Values are reported as the mean percentage
survival
standard error of the mean (SEM) of three biological
replicates (values in the experimental section).
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the 1,6-NRH isomers reduced cell survival by at least 10%, and
this change was statistically significant. 1,6-NRH was found to
be the most cytotoxic isomer (Figure 5), and HEK293T cells
are more sensitive to this isomer than HepG3 cells. 1,2-NRH
was found to be less cytotoxic to these cells. This difference
could be attributed to the reduced stability of 1,2-NRH in the
media.
indication that 1,2-NRH decomposed materials were taken
out of the soluble phase. Such materials are likely to be protein
adducts.
Critically, FAD alone was able to facilitate the oxidation of
1,4-NRH (Figure 6) and 1,6-NRH to NR+ (Figure S3).
However, in HEPES buffer containing BSA and FAD at pH 7.2
in the presence of the cosubstrate menadione, the rate of
oxidation of 1,4-NRH to NR+ was greatly increased by NQO2
(Sigma-Aldrich; Figure 6). Under the selected conditions for
measuring NQO2 activity, menadione was the rate-limiting
substrate, being present at 1/2 the concentration of the NRHs.
The residual 1,4-NRH present in 1,2-NRH and 1,4-NRH
samples was readily oxidized to NR+ when NQO2 was present
(Figure S4). Compared to 1,4-NRH, the oxidation of 1,6-NRH
by NQO2 was significantly slower (Figure 7), and this
oxidation could be accounted for by the presence of free FAD.
Crucially, 1,6-NRH appeared to be a relatively stable isomer of
1,4-NRH under the reaction conditions.
Metabolism of 1,2-, 1,4-, and 1,6-NRH to NR+. Renalase
and pyridoxamine-phosphate oxidase are riboflavin-dependent
oxidases, using FAD and FMN, respectively. NAD(P)H
quinone dehydrogenase (NQO1) is one of the most
ubiquitous riboflavin-dependent oxidases and one of the two
major quinone reductases in mammalian systems. Its
homologue NQO2, known as ribosyl dihydronicotinamide
dehydrogenase, quinone, is the only enzyme known to oxidize
1,4-NRH to NR+ in the presence of a co-oxidant like O2 or
menadione (Figure 1). NQO2 is expressed in both HEK293T
and HepG3 cells, and its levels remain constant following the 6
h exposure to 100 μM 1,4-NRH. Considering the role of
NQO2 as a detoxifying enzyme, we sought to establish
whether the 1,2- and 1,6-NRH isomers affected the NQO2
1
function, as a possible route to cell toxicity. Under H NMR-
enabled monitoring conditions, we examined whether NQO2
could oxidize all three nucleosides. First, we sought to establish
whether these isomers were stable to the enzymatic reaction
conditions. We rapidly realized that both BSA and FAD
significantly modified the relative amount to NRH derivatives
in solution over time (Figure S2), including 1,4-NRH (Figure
6). Under these conditions, we observed more than 20%
Figure 7. 1,2-NRH, 1,4-NRH, and 1,6-NRH are oxidized to NR by
NQO2 in the presence of FAD and menadione. 1,2-NRH and 1,6-
NRH are oxidized to NR+ more slowly than 1,4-NRH. Slow oxidation
of all three isomers is observed in the presence of FAD only, with 1,4-
NRH being the most reactive toward FAD.
Inhibition of NQO2 by 1,2- and 1,6-NRH. With this
perspective that the rate of oxidation of 1,2- and 1,6-NRH to
NR+ was slower than that of 1,4-NRH to NR+, we decided to
investigate whether 1,2-NRH and 1,6-NRH could be inhibitors
of NQO2. Co-incubation of 1,4-NRH and NQO2 in the
presence of molar equivalents of 1,6-NRH led to a decrease in
the rate of production of NR+ (Figure 8). The loss of NRH-
derived species was also less when 1,6-NRH was used (Figure
8, panel B). This observation is consistent with 1,6-NRH
behaving as a stable isomer of 1,4-NRH in solution.
Modeling of 1,2-NRH and 1,6-NRH in the Binding Site
of NQO2. We sought a rationale for the more pronounced
inhibitory effect of 1,6-NRH than of 1,2-NRH on NQO2.
Using the computational model of FAD-bound NQO2 based
upon PDB entry 2QX8,48 we have found dock poses for the
two isomers at the hydride-exchange site of FAD-bound
NQO2. Our quantum-based computational modeling indicates
that the ring system of 1,6-NRH stacks favorably with the
central ring of the flavin moiety and overlaps the expected
binding mode of 1,4-NRH (Figure S5). This inhibitory binding
position of 1,6-NRH is itself unlikely to lead to hydride transfer
with r(H6−NFAD) = 3.2 Å versus r(H4−NFAD) = 2.7 Å for
1,4-NRH. For the 1,2-NRH isomer (not shown), we find a
dock pose similar to that of 1,6-NRH; r(H2−NFAD) is
likewise too far to engender hydride transfer, and weaker
Figure 6. Panel A: % of loss over time. Panel B: Relative products’
distribution over time.
material loss over 24 h in the 1,6-NRH and 1,4-NRH
containing reaction solutions and up to 65% material loss over
that same period for the 1,2-NRH containing solution (Figure
S2). In the case of 1,2-NRH, the addition of the 1,2-NRH to
the buffer containing BSA and FAD promoted an almost
1
instantaneous change in the H NMR spectrum of 1,2-NRH,
which over time lost signal-to-noise resolution, a clear
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incubation process in the same buffer as that used for NQO2
reactions, but in the absence of NQO2, the protein content of
this reaction mixture was analyzed following simple tryptic
digest followed by LC-MS2 analyses to determine whether
glycation other than the one anticipated from a residual
riboside conjugation could be detected. The conjugation of a
ribosylated aldehydic intermediate such as the aldehyde
described in Figure 4 with a lysine residue was thought to
cause a C11H18N2O5 addition. We observed this increase in the
occurrence of adducts in several peptidic sequences, all rich in
lysine residues.
Furthermore, upon the loss of the ribose, the anticipated
mass shift stemming from a monoconjugation of a dialdehyde
(Figure 4) with a lysine residue of BSA would be (125 = −2 +
127)(−2H + C6H9NO2) with one positive charge loss. Many
peptidic sequences with this type of modification were also
detected. The remaining aldehyde can be readily oxidized to
the carboxylic acid upon sample handling, and a mass shift of
141 (−2H + C6H9NO3) can be rationalized. Critically, some of
these modifications are already present on the BSA before its
exposure to 1,4-NRH, indicating that BSA modifications by
NRH-like entities occur endogenously.
DISCUSSION
■
The universally agreed correlation between the maintenance of
NAD levels and (NAD+/NADH) ratio, and cellular homeo-
stasis, has led to a wide range of nutritional approaches seeking
to increase NAD bioavailability in humans to improve health,
ward off aging, and mitigate diseases. A range of products to
increase NAD bioavailability are now available (e.g., ref 49).
While changes in circulating NAD levels can be monitored, the
effects of a given supplementation on the NADH levels and
thus the NAD+/NADH ratio are often overlooked. A sustained
or even transient unregulated increase of NADH abundance
associates with reductive stress that leads to reactive oxygen
species generation and oxidative stress outcomes.38 The
possibility of increasing the levels of NADH as total NAD
levels rise should be of concern. Pyridoxamine-phosphate
oxidase and renalase are two mammalian enzymes that are
known to oxidize endogenous 1,2- and 1,6-NAD(P)H to
NAD(P)+.30,33 Crucially, renalase is known to oxidize 1,2- and
1,6-NAD(P)H preferentially over 1,2- and 1,6-NRH.19,50 It has
low expression levels in HEK293T but increases upon
oxidative stress in hepatocytes.51,52 Similarly, pyridoxamine-
phosphate oxidase is primarily expressed in the liver. The
toxicity observed for the two isomers of 1,4-NRH in HEK293T
cells could stem from their conversion to 1,2- and 1,6-NADH,
respectively, going unchallenged by low renalase and pyridox-
amine-phosphate oxidase expression, and the subsequent
inhibition of redox enzymes. Alternatively, these nucleosides
could compromise cellular function through mechanisms not
requiring conversion to dinucleotides, but rather are associated
with their ability to decompose to aldehydes and glycating
Figure 8. Inhibition of NQO2 catalyzed oxidation of 1,4-NRH to
NR+ by 1,2-NRH and 1,6-NRH: Panel A: 1,6-NRH reduces the rate
of 1,4-NRH oxidation by NQO2 more effectively than 1,2-NRH.
Panel B: The loss of NRH isomers present in solution occurs over
time under NQO2 incubation conditions in the presence of 1,6-NRH.
inhibitory activity is expected because 1,2-NRH’s amide abuts
Trp195 of NQO2.
NRH and Its Degradation Products React with
Peptidic Lysines and Modify Proteins. Protein modifica-
tions occur widely in cells and in vivo. For instance, protein
glycation by reductive sugars like glucose is common and
occurs via a reaction between the aldehydic form of glucose
and a free amine residue, either an N-terminus or a lysine side
chain, leading to a Schiff-base rearrangement. These
modifications are not reversible. This type of modification
occurs with other types of glycating species such as glyoxal etc.
The chemical commonality in these types of modifications is
the reactive aldehyde. NAD is also known to modify proteins
via parylation and modification of the peptidic sequence by the
introduction of the ribosyl moiety. Unlike glycation, this
process is often reversible and does not include the formation
of a Schiff base. Having observed a sustained loss (irreversible)
of the three NRH isomers in the presence of the buffer protein
BSA over time, we decided to explore whether conjugation to
proteins could be detected upon incubation of BSA with either
of these NRH isomers. Considering that 1,4-NRH is the most
abundant form of NRH and a representative of functional 1,4-
NADH, we selected it for this investigation. Following a 24 h
Table 1. Coverage and Relative Abundance of Selected Lysine Modified BSA Peptides
sequence
modifications
abundances in BSA
abundance in BSA treated with 1,4-NRH
LHEKTPVSEKVTK
LHEKTPVSEK
CTKPESERMPCTEDYLSLILNR
YQEAKDAFLGSFLYEYSR
NRH adduct C6H9NO2 [K4]
NRH adduct C6H9NO2 [K4]
NRH adduct C6H9NO3 [K3]
NRH adduct C11H18N2O5 [K5]
2.61 × 1008
32 803 514
BDL
1.1 × 1009
88 550 358
6.46 × 1008
16 282 244
8 007 322
610
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NRH + H2O solution was transferred to an NMR tube. A total of 50
μL of D2O was added to the NMR tube to yield a total volume of 500
μL. The NMR tube was vortexed to achieve a homogeneous solution
and then placed in a 37 °C hot water bath for 24 h, after which time a
1H NMR spectrum was taken.
derivatives (Figure 4) affecting HEK293T more than HepG3
cells.
With the present work, we explored the chemical stability of
1
the dihydropyridine moiety using H NMR spectroscopy to
evaluate the chemical changes that 1,4-NRH undergoes while
in solution. Other analytical methods, more prevalent in cell
biology and molecular biology, which include mass spectrom-
etry coupled to liquid chromatography, UV spectroscopy, as
well as fluorescence, do not allow the characterization of the
Figure 4, Panel B: 50 mg of 1,4-NRH was dissolved in 2.5 mL of
sodium phosphate buffer (500 mM, pH = 6.0). A total of 450 μL of
the 1,4-NRH + buffer solution was transferred to an NMR tube. A
total of 50 μL of D2O was added to the NMR tube to yield a total
volume of 500 μL. The NMR tube was vortexed to achieve a
homogeneous solution and then placed in a 37 °C hot water bath for
1
changes as they happen; H NMR spectroscopy provides this
1
30 min, after which time a H NMR spectrum was taken.
flexibility. Having explored the stability of 1,4-NRH and found
that it correlates quite closely to that of NAD(P)H with regard
to its reactivity to oxygen and ability to isomerize to 1,6-NRH,
we chemically synthesized and isolated the pure form of the
other isomers of 1,4-NRH and explored the integrity of the
heterocycle. We demonstrated here that 1,4-NRH’s isomers
inhibited NQO2 and that 1,4-NRH can also generate glycating
entities. We observed that the addition of water on 1,4-NRH
was rapid in phosphate-containing buffers, and that decom-
position led to intermediates which readily react forming Shiff
bases, those of solvent-exposed lysine residues of proteins of
the like of BSA. While 1,4-NRH might not be overly abundant,
its chemical reactivity toward protein residues could provide
some evidence that intracellular NADH might promote protein
aging, especially in organelles where it is the most abundant,
the mitochondrion. Such conjugation is particularly relevant to
cellular regulation for which nonreversible protein modifica-
tions could lead to protein loss of function and over time to the
aging of the proteome. Therefore, NQO2’s role in the
maintenance of cellular homeostasis might be less about the
detoxification of xenobiotics and more about the removal of
endogenous excess 1,4-NRH, generated from excess NADH.
Such a paradox highlights a new role for NQO2 activity, as
its enzymatic activity has dual functions: reduction of quinones
in a detoxification process and oxidation of NRH to NR+ to
maintain low NRH intracellular levels. This work provides
some insights as to how NAD(P)H:quinone oxidoreductase 1
(NQO1), the homologue of NQO2 which uses NADH and
NADPH as substrates, might be interacting with the respective
isomers of its dinucleotide substrates. Enzymes that reduce the
levels of unbound NADH, like NQO1, would prevent the
occurrence of these derivatives. While the cellular mechanisms
underpinning the observed outcomes remain to be inves-
tigated, the role of excess NADH and its ability to chemically
isomerize and generate endogenous 1,2- and 1−6-NADH
isomers as well as Schiff-base adducts could be a link between
metabolic dysfunction and intracellular NADH overabun-
dance.
Isomerism of NRH and NR+. 1,4-NRH was coincubated with
NR+ in D2O (final concentration 10 mM of each component). Under
these conditions, the 1,6-NRH isomer formed and could be detected
after 100 h. Products that can readily be identified include alpha and
beta-ribose, nicotinamide, NR+, and 1,6-NRH. The appearance of
products with aliphatic hydrogens consistent with structures proposed
by Margoli via the intermediacy of NR(OH)H, which can also
generate decomposition products, remain to be characterized.
Cell Culture. Human Embryonic Kidney (HEK293T) and
Hepatoma (HepG3) cells were purchased from American Type
Culture Collection (ATCC). Cells were grown at 37 °C in a 5% CO2
incubator in Dulbecco’s modified Eagle’s medium (DMEM; Hyclone)
supplemented with glutamine (Gibco), 10% fetal bovine serum (FBS;
Atlanta Biologicals Premium Select), and 1% sodium pyruvate
(Gibco). Cells were routinely tested and found to be free of
mycoplasma contamination using Lonza MycoAlert Assay.
Cytotoxicity Studies. The cytotoxicity of NRH and its derivatives
was determined by CellTiter-Fluor (Promega), which measures cell
viability independent of metabolic markers related to NAD(P)(H).
HEK293T cells were seeded at a density of 5000 cells/well in a 96-
well clear bottomed black plate coated with poly-L-lysine (Corning).
HepG3 cell lines were seeded at a density of 15 000 cells/well in an
uncoated 96-well clear bottomed black plate (Greiner Bio). Cell
plating density was optimized to ensure ∼75% confluency at the end
of the assay. The following day, 100 mM stock solutions of 1,4-NRH,
1,2-NRH, and 1,6-NRH were prepared in a phosphate buffered saline
solution without calcium or magnesium (PBS, VWR). Cells were
exposed to 100 μM of each compound in the medium for 6 h. After 6
h, supplements were washed once in PBS, and fresh medium was
replaced. The cells were placed back at 37 °C in a 5% CO2 incubator
and allowed to grow for another 66 h (total 72 h). At the end of 72 h,
100 μL of CellTiter-Fluor reagent was added to all wells and
incubated for at least 30 min at 37 °C. Fluorescence intensity was
measured using a Tecan Infinite M1000 Pro plate reader (380 nm Ex/
505 nm Em). The fluorescent signal is then expressed as a percentage
relative to the control to reflect cell survival. Values are reported as
the mean percentage survival standard error of the mean (SEM) of
three biological replicates.
Recombinant Human NQO2 Enzyme Activity. Recombinant
human NQO2 was obtained from Sigma and diluted in 50 mM
phosphate buffer. All one-dimensional proton NMR spectra were
obtained at 300 K on a Bruker Ascend 400 MHz ultrashielded
spectrometer (Bruker Biospin) operating at 400.13 MHz for protons
and used to determine enzyme activity. TopSpin 3.2 (Bruker BioSpin)
was used for all NMR spectral acquisition (ns = 128) and
preprocessing, and the automation of sample submission was
performed using ICON-NMR (Bruker BioSpin). All samples were
automatically shimmed, and their acquisition time was 10 min and 8 s.
All experiments were performed at 25 °C. The 1,2-NRH, 1,4-NRH,
and 1,6-NRH used were synthesized according to the procedure
described in the experimental supplementary file. The rates of the
NQO2 enzyme were determined by NMR spectroscopy by recording
the rate of oxidation of 1,2-NRH, 1,4-NRH, and 1,6-NRH to NR at t
= 0, 6, and 24 h (ns = 128) at 25 °C in the NMR tube, made up of
borosilicate glass (7 in. × 5 mm) containing a final volume of 5.10
mL: 450 μL of phosphate buffer (50.0 mM, pH 7.1), 10 μL of
recombinant NQO2 (1 mg dissolved in 500 μL HEPES buffer), 50.0
μL of NRH (10.0 mM in D2O). All the experiments were performed
METHODS
■
Full details of the synthetic and analytical experiments reported in this
manuscript can be found in the Supporting Information. The full
characterization spectra of the reported molecules can be also found
in the Supporting Information.
Stability of NRH in Water and in Phosphate Buffer. Figure 3:
50 mg of 1,4-NRH was dissolved in 2.5 mL of sodium phosphate
buffer (50 mM, pH = 6.8). A total of 450 μL of the 1,4-NRH + buffer
solution was transferred to an NMR tube. A total of 50 μL of D2O was
added to the NMR tube to yield a total volume of 500 μL. The NMR
tube was vortexed to achieve a homogeneous solution and then placed
1
in a 37 °C hot water bath for 1, 3, 12, and 24 h, at which time a H
NMR spectrum was taken.
Figure 4, Panel A: 30 mg of 1,4-NRH was dissolved in 1.5 mL of
deionized degassed water (pH= 7.3). A total of 450 μL of the 1,4-
611
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in a single day. For each independent experiment, freshly prepared
solutions of NRHs in D2O and 450 μL of phosphate buffer,
containing 125 mM NaCl, 5 μM FAD, BSA (1 mg mL−1), and 10 μL/
mL menadione (93 mM solution in 1 mL ethanol) were used.
Identification of Modified Lysine Residues in BSA. A total of
50 μL of 1 mg mL−1 bovine serum albumin (BSA) was incubated with
50 μL of 1,4-NRH and 400 μL of a phosphate buffered saline solution
at RT for 8 days. BSA was also incubated in the absence of NRH for
comparison at the same concentration. A total of 100 μL of each
solution was further diluted with 900 μL of 0.1% trifluoroacetic acid
(TFA) and loaded onto separate washed and equilibrated C2 columns
(Waters Sep-Pak Vac 1 cm3 C2 cartridge, WAT052710). After the
entire sample was loaded, the column was washed with 3 mL (3
column volumes) of 0.1% TFA, followed by an additional wash with 3
mL of 10% ACN in 0.1% TFA. The protein was eluted from the
column with 2 mL of 60% ACN in 0.1% TFA collected into one
fraction and brought to dryness in a speed-vac concentrator. The
dried sample was resuspended into 50 μL of 50 mM ammonium
bicarbonate/10 mM tris(2-carboxye thyl)phosphine hydrochloride
and digested for 7 h with 1 μL sequencing grade trypsin at 37 °C.
High resolution mass spectrometry was performed on the resulting
peptide solution using a Q-Exactive Plus mass spectrometer (Thermo
Fisher Scientific) coupled with a Thermo Easy-nLC 1000 liquid
chromatograph. A 0.5 μL injection volume was used for each sample,
loaded onto an ES800 C18 column equipped with a C18 guard
column. Solvent A consisted of 3% acetonitrile (ACN) in H2O with
0.2% formic acid (FA), and solvent B consisted of 3% water in ACN
with 0.2% FA. The flow rate was set to 300 nL per minute. To provide
chromatographic separation, a linear solvent gradient was used from
2% to 30% B over a period of 36 min, as well as a linear gradient from
30% B to 90% B for an additional 15 min. A 5 min wash at 90% B was
used to elute any remaining material from the column at the end of
each run. Blanks were also run between samples to minimize
carryover. The EASY-spray source was used, in positive ion mode, to
introduce the sample into the MS using electrospray ionization. A
spray voltage of 2500 and a capillary temperature of 250 °C was used.
One full scan from 400 to 2000 m/z was performed at 70,000
resolution, followed by 6 dd-ms2 scans at 17 500 resolution of the top
6 features identified in the full scan. Features with a charge of no less
than 2 and no more than 8 were selected for HCD fragmentation with
a 2.0 m/z isolation window, using a normalized collision energy of 27.
A dynamic exclusion of 15 s was utilized to maximize coverage. Total
run time for each sample analyzed was 60 min. The resulting raw files
were uploaded into Thermo Proteome Discoverer software v2.2 to
provide peptide identification. The sequest processing mode was used
to search a custom fasta file containing the sequence of bovine serum
albumin, with a precursor mass tolerance of 5 ppm and a fragment
mass tolerance of 0.05 Da. Semitryptic peptides were allowed with a
maximum of two missed cleavage sites.
The following custom dynamic modifications specific to lysine
residues were included in the search: (1) NRH Adduct
C11H18N2O5/+258.122 Da (K); (2) NRH/+256.106 Da (K); (3)
NRH Adduct C11H18N2O6/+274.117 Da (K); (4) NRH Adduct
C6H9NO2/+127.063 Da (K); (5) NRH Adduct C6H9NO3/
+143.058 Da (K).
A maximum of two equal modifications and four dynamic
modifications were allowed per peptide. The spectrum confidence
filter node was used to filter out any spectra that did not have high
confidence. The percolator node was used with a target FDR (strict)
set to 0.01 and 0.05 for relaxed, with validation based on q value.
Precursor quantification was performed using both unique and razor
peptides, with abundance based on area. No normalization or scaling
was applied.
Quantum-Based Computational Modeling NQO2. A trun-
cated model of the NQO2(FAD) enzyme (17 amino acid residues)
was built from PDB entry 2QX8 [48] (res 1.60 Å) using Gaussview
5.0 (Gaussian Inc., Wallingford, CT). The FAD molecule was
truncated at the first phosphate to produce a model system with a net
charge of −1. In the search for favorable docking poses of the three
NRH isomers on the face of FAD, eight classes of poses of each
isomer were considered through manual positioning: two positions for
the amide moiety × two positions for the glycosidic bond × two
positions for the ribose moiety (left versus right of FAD). Manual
docking was followed by semiempirical PM7 energy optimizations in
which only the NRH ligand was permitted to move. The resulting
lowest energy structures were selected for follow-up ONIOM hybrid
calculations (high-level:B3LYP[3,4]/6-31G(d):low-level:PM7) in
order to distinguish them by using an all-electron density functional
model (B3LYP) for the high-level region. The ligand positions were
reoptimized, ultimately using the ONIOM(wB97XD/6-31G-
(d):PM7) model in the presence of water using the default SCRF
model. The wB97XD model is a density functional method that
includes the effects of dispersion. Optimizations were deemed
complete when the forces met the default convergence criteria. All
calculations were carried out using Gaussian16.
CONCLUSION
■
Intracellular 1,4-NAD(P)H, just like 1,4-NRH in solution, not
only oxidizes but also generates isomers and hydroxylated
adducts. Accumulation of NAD(P)H in cells53,54 would lead to
increased formation of these isomeric and hydrated species,
and since these species possess irreversible chemical reactivities
to protein conjugating agents, the equilibrium is constantly
displaced toward their formation and thus could deplete the
NAD(P)(H) pools. The work highlights some of the processes
that such endogenously generated species are likely to disrupt.
Unfortunately, quantification of these NAD(P)H catabolites
from cell culture or tissues is not yet possible as they
“disappear” from the quantifiable environment. Additionally,
protein modifications by such species could be substantial and
be mistaken for enzyme-catalyzed ADP ribosylation, in a way
reminiscent of the chemically driven mitochondrial protein
acetylation.55 It is this nonenzymatic intrinsic biochemistry
that NRH helps expose. Critically, this work brings up possible
unwanted effects of boosting not only NAD+ but also NAD(P)
H in cells and in tissues, as these reduced forms could
potentially be detrimental to cellular homeostasis and promote
cellular dysfunction.
ASSOCIATED CONTENT
* Supporting Information
The Supporting Information is available free of charge at
https://pubs.acs.org/doi/10.1021/acschembio.0c00757.
■
sı
Fully synthetic experimental details; full characterization
1
spectra; H NMR, 13C NMR, 2D COSY, and HMQC,
HRMS, and comparative fragmentation patterns of the
1,2-NRH, 1,6-NRH, and 1,4-NRH compounds reported
in the Methods section (PDF)
AUTHOR INFORMATION
Corresponding Author
■
Marie E. Migaud − University of South Alabama, Department
of Pharmacology, Mobile, Alabama 36688-0002, United
States; orcid.org/0000-0002-9626-2405;
Email: mmigaud@southalabama.edu
Authors
Mikhail V. Makarov − University of South Alabama,
Department of Pharmacology, Mobile, Alabama 36688-
0002, United States
Faisal Hayat − University of South Alabama, Department of
Pharmacology, Mobile, Alabama 36688-0002, United States
Briley Graves − University of South Alabama, Department of
Pharmacology, Mobile, Alabama 36688-0002, United States
612
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Articles
(5) Stromland, O., Niere, M., Nikiforov, A. A., VanLinden, M. R.,
Heiland, I., and Ziegler, M. (2019) Keeping the balance in NAD
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Manoj Sonavane − University of South Alabama, Department
of Physiology and Cell Biology, Mobile, Alabama 36688-
0002, United States
Edward A. Salter − University of South Alabama, Department
of Chemistry, Mobile, Alabama 36688-0002, United States
Andrzej Wierzbicki − University of South Alabama,
Department of Chemistry, Mobile, Alabama 36688-0002,
United States
Natalie R. Gassman − University of South Alabama,
Department of Physiology and Cell Biology, Mobile, Alabama
36688-0002, United States; orcid.org/0000-0002-8488-
2332
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Complete contact information is available at:
https://pubs.acs.org/10.1021/acschembio.0c00757
Author Contributions
Mikhail V. Makarov: synthesis, data curation, methodology,
writing−review and editing. Faisal Hayat: enzyme kinetics,
synthesis, data curation, methodology, writing−review and
editing. Briley Graves: stability work, data curation, method-
ology, writing−review and editing. Manoj Sonavane: cell-based
work, data curation, methodology, writing−review and editing.
Edward A. Salter: computational analyses, data curation,
methodology, writing−review and editing. Andrzej Wierzbicki:
computational analyses, writing−review and editing. Natalie R.
Gassman: data curation, formal analysis, supervision, writing−
review and editing. Marie E. Migaud: conceptualization, data
curation, formal analysis, funding acquisition, supervision,
writing−review and editing.
Author Contributions
(17) Morrison, C. S., Armiger, W. B., Dodds, D. R., Dordick, J. S.,
and Koffas, M. A. G. (2018) Improved strategies for electrochemical
1,4-NAD(P)H2 regeneration: A new era of bioreactors for industrial
biocatalysis. Biotechnol. Adv. 36 (1), 120−131.
†Joint first authorship.
Funding
This work was supported by R21AT009908 from the National
Institutes of Health, National Center for Complementary and
Integrative Health. M.E.M. is partially supported by Elysium
Health. The funders had no role in the study design, data
collection and analysis, decision to publish, or preparation of
the manuscript.
(18) Beaupre, B. A., Hoag, M. R., Roman, J., Försterling, F. H., and
Moran, G. R. (2015) Metabolic function for human renalase:
oxidation of isomeric forms of β-NAD(P)H that are inhibitory to
primary metabolism. Biochemistry 54 (3), 795−806.
(19) Moran, G. R., and Hoag, M. R. (2017) The enzyme: Renalase.
Arch. Biochem. Biophys. 632, 66−76.
̌
(20) Kremer, L. S., Danhauser, K., Herebian, D., Petkovic Ramadza,
Notes
D., Piekutowska-Abramczuk, D., Seibt, A., Muller-Felber, W., Haack,
̈
The authors declare no competing financial interest.
T. B., Płoski, R., Lohmeier, K., Schneider, D., Klee, D., Rokicki, D.,
Mayatepek, E., Strom, T. M., Meitinger, T., Klopstock, T., Pronicka,
E., Mayr, J. A., Baric, I., Distelmaier, F., and Prokisch, H. (2016)
NAXE Mutations Disrupt the Cellular NAD(P)HX Repair System
and Cause a Lethal Neurometabolic Disorder of Early Childhood.
Am. J. Hum. Genet. 99 (4), 894−902.
ACKNOWLEDGMENTS
■
We thank The University of South Alabama Mass Spectrom-
etry Core Facility and the valuable technical contributions
from L. Schambeau and S. Tadi. We thank the Alabama
Supercomputer Authority for providing computational support
and resources.
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