Nicotinamide-Containing Di- and Trinucleotides as Chemical Tools for Studies of NAD-Capped RNAs

Article abstract of DOI:10.1021/acs.orglett.8b03386

We report the chemical synthesis of a set of nicotinamide adenine dinucleotide (NAD) cap analogues containing chemical modifications that reduce their susceptibility to NAD-RNA-degrading enzymes. These analogues can be incorporated into transcripts in a similar way as NAD. Biochemical characterization of RNAs carrying these caps with DXO, NudC, and Nudt12 enzymes led to the identification of compounds that can be instrumental in unraveling so far unaddressed biological aspects of NAD-RNAs.

Full text of DOI:10.1021/acs.orglett.8b03386

Letter
pubs.acs.org/OrgLett
Cite This: Org. Lett. XXXX, XXX, XXX−XXX
Nicotinamide-Containing Di- and Trinucleotides as Chemical Tools
for Studies of NAD-Capped RNAs
Agnieszka Mlynarska-Cieslak,†,∥ Anais Depaix,†,∥ Ewa Grudzien-Nogalska,‡ Pawel J. Sikorski,§
Marcin Warminski,† Megerditch Kiledjian,‡ Jacek Jemielity,§ and Joanna Kowalska*,†
†Division of Biophysics, Institute of Experimental Physics, Faculty of Physics, University of Warsaw, Pasteura 5, 02-093 Warsaw,
Poland
‡Department of Cell Biology and Neuroscience, Rutgers University, Piscataway, New Jersey 08854, United States
§Centre of New Technologies, University of Warsaw, Banacha 2c, 02-097 Warsaw, Poland
S
* Supporting Information
ABSTRACT: We report the chemical synthesis of a set of
nicotinamide adenine dinucleotide (NAD) cap analogues containing
chemical modifications that reduce their susceptibility to NAD-RNA-
degrading enzymes. These analogues can be incorporated into
transcripts in a similar way as NAD. Biochemical characterization of
RNAs carrying these caps with DXO, NudC, and Nudt12 enzymes
led to the identification of compounds that can be instrumental in
unraveling so far unaddressed biological aspects of NAD-RNAs.
icotinamide adenine dinucleotide (NAD), discovered
in vitro and in vivo via pyrophosphate bond cleavage (Figure
Nover 100 years ago as a cofactor in cellular redox 1A).8,9
In contrast, in eukaryotes the presence of NAD at the 5′-end
reactions, has since been thoroughly studied and demonstrated
to be a highly important biomolecule with multiple biological
functions. Besides participation in redox biotransformations,
NAD serves as a substrate for protein post-translational
modifications (poly(ADP-ribosylation, deacetylation), for
signal transduction (as a precursor of cyclic ADP ribose),1
and, as was shown recently, as a 5′-end-modifying molecule in
some RNAs.2 NAD-capped RNAs (Figure 1A) were first found
in bacteria2 and thus are termed “bacterial caps” because of
some structural analogy to the 7-methylguanosine (m7G) cap
present at the 5′-end of eukaryotic mRNAs.3 More recently,
NAD-capped RNAs have also been discovered in other
organisms ranging from bacteria to eukaryotes,4,5 including
mammals. The latest studies indicated that up to 10% of
human mitochondrial transcripts are NAD-capped.6 The
biological roles of this new, fascinating RNA modification
remain elusive and likely differ among organisms. Nonetheless,
it has been established that in both bacteria and eukaryotes, the
5′-terminal NAD moiety influences RNA stability. Accord-
ingly, several deNADding enzymes that participate in RNA
degradation have been identified.7
appears to destabilize RNA and direct it for decay. The decay
is initiated by the removal of the whole 5′-terminal NAD
moiety by DXO nuclease, which cleaves the first phospho-
diester bond in RNA (Figure 1A).4 Nudt12 pyrophosphatase,
an eukaryotic homologue of NudC, also cleaves NAD-RNA in
vitro and contributes to NAD-RNA decay in human cells.10
Despite the increasing number of novel NAD-related analytical
and chemical biology tools,11 the biological significance of
NAD-capped-RNAs in different organisms remains unclear.
One of the most important questions to be addressed is
whether this modification has specific biological functions in
cells (e.g., to act as a signaling response to stress conditions) or
is merely a result of aberrant transcription initiation that is
rapidly eliminated by RNA quality control mechanisms.
We envisaged that investigations of the structure and
function of NAD-terminated RNAs and their specific binding
proteins can be expedited by the use of rationally designed
molecular tools enabling the synthesis of NAD-RNAs that are
less susceptible to enzymatic cleavage than natural NAD-
RNAs. Such tools would provide access to NAD-RNAs with
increased cellular stability suitable for structural and functional
studies, pull-down experiments, and many others.
In Escherichia coli, the presence of the 5′-NAD moiety
arguably increases the stability of RNA by decreasing its
susceptibility to the action of RppH,8 a bacterial pyrophos-
phatase that participates in dephosphorylation of RNA 5′-
triphosphate to 5′-monophosphate, thereby triggering 5′-
decay. However, NAD-capped RNAs are targeted by NudC,
a bacterial enzyme capable of specific 5′-deNADding of RNA
Here we report the synthesis of a set of novel NAD
analogues that can be incorporated into RNA by in vitro
Received: October 23, 2018
© XXXX American Chemical Society
A
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confer resistance to phosphodiesterases such as DXO,
stabilizing modifications were introduced within the first
phosphodiester bond (PD1) by utilizing trinucleotide NAD
analogues (compounds 3−6; Figure 1B). In addition to
replacing the first phosphodiester bond with the PS moiety
(two diastereomers of compound 4), we also explored 2′-O-
methylation of the first nucleoside in the RNA body
(compound 5), as it has been recently reported to decrease
the susceptibility to DXO for m7GpppN and GpppN-capped
RNAs.13 Finally, expecting to achieve resistance to both types
of deNADding enzymes, we combined the O-to-S substitution
at the α-phosphate (α-PS) with 2′-O-methylation (compounds
6a and 6b).
A key intermediate in the synthesis of all of the NAD
analogues was imidazole-activated nicotinamide mononucleo-
tide (8), obtained from nicotinamide riboside (NR) 5′-
monophosphate (7). The latter can be either synthesized de
novo by a multistep procedure,14 prepared enzymatically,15 or
obtained by hydrolysis of commercially available NAD in the
presence of aqueous ZrCl4.16 We chose the last of these
methods, as it appeared the most time- and cost-efficient for
gram-scale synthesis. The resulting mixture of AMP and
nicotinamide mononucleotide was separated by ion-exchange
chromatography (IEC) using DEAE Sephadex A25 to give
compound 7 as a triethylammonium salt in 30% yield (Scheme
1A). Reaction of 7 with imidazole in the presence of
triphenylphosphine and 2,2′-dithiodipyridine in DMSO
followed by precipitation with NaClO4 in acetone gave the
sodium salt of P-imidazolide 8 in 56% yield. Adenosine 5′-
phosphorothioate (9) was synthesized by thiophosphorylation
of adenosine with PSCl3 in the presence of 2,6-lutidine
followed by hydrolysis17 and purification by IEC. Finally, to
yield α-substituted NAD dinucleotide 1, compounds 8 and 9
were reacted in the presence of a 3-fold excess of ZnCl2 in
DMF as the solvent. RP-HPLC analysis of the reaction mixture
revealed the formation of two P-diastereomers of 1 (1a and
1b) in a ratio of ∼1:1.4 (Figure 2A) and nicotinamide
nucleotide dimer (NRMP)2 as a side product. The
diastereomers were labeled D1 (1a) and D2 (1b) according
to their elution order from the RP HPLC column. The
diastereometric product was isolated by IEC, after which the
diastereomers were separated by semireparative RP-HPLC.
Compound 2 was synthesized in an analogous coupling
reaction of 8 and adenosine 5′-phosphorothiolate (11), which
was obtained by S-alkylation of triethylammonium phosphor-
othioate by 5′-deoxy-5′-iodoadenosine.18
Figure 1. (A) Structure of NAD-linked RNA and activity of
deNADding enzymes identified to date. (B) Structure of NAD
analogues synthesized in this study. Abbreviations: NR, nicotinamide
riboside; A, adenosine; G, guanosine.
transcription (IVT). These analogues were designed to be
either partially or completely resistant to hydrolytic enzymes
that target NAD-RNAs. We further demonstrate efficient
incorporation of these NAD analogues into RNA by
polymerase T7. Finally, we examine the susceptibility of the
modified NAD-linked RNAs to several deNADding enzymes
(NudC, Nudt12, and DXO) and identify analogues that are
most suitable for future biological applications.
To achieve resistance to NUDIX-family pyrophosphatases
such as NudC and Nudt12, we designed dinucleotide NAD
analogues carrying an O-to-S substitution at the nonbridging
position of the pyrophosphate moiety or at the 5′-bridging
position of the phosphoester (Figure 1B). This type of
modification has been previously shown to be a viable strategy
for stabilizing m7G-capped RNAs against decapping enzymes
without disrupting their translational activity.12 To this end,
three dinucleotide NAD analogues were synthesized: two
stereoisomers of NAD carrying the α-phosphorothioate (α-PS)
moiety (compounds 1a and 1b) and one NAD analogue
carrying 5′-phosphorothiolate (PSL) (compound 2). To
Dinucleotide precursors required for the synthesis of
trinucleotide NAD analogues were obtained by phosphor-
amidite solid-phase synthesis (Figure S1) using iodine and
DDTT as oxidating and sulfurizing reagents, respectively. The
coupling reactions with 8 that led to trinucleotides were carried
out under conditions similar to those described above (Scheme
1B). However, these reactions proceeded notably slower,
requiring 2−3 days for maximal conversion. The yields of all
final compounds (1−6) were moderate to low (8−33%),
mainly because of the two-step purification required before
biochemical studies. The spectroscopic and high-resolution
mass spectrometry data for all of the analogues were consistent
with the presence of the oxidized form (NAD+) with trace
amounts of NADH (<5%). 31P NMR analysis confirmed the
presence of phosphorothioate and phosphorothiolate moieties,
as noted by the shifts of the resonance signals by approximately
+55 and +20 ppm, respectively, compared with the unmodified
B
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Scheme 1. Synthesis of (A) Dinucleotide and (B) Trinucleotide NAD Analogues
phosphate moieties (Figure 2B) and by the increase in the
2JP−P coupling constant from 19−20 to 27−28 Hz. The
absolute configurations of isomers D1 and D2 were not
assigned, but 31P and 1H NMR data confirmed that the
separation was successful (Table S1).
Next, we tested whether the newly synthesized NAD
analogues can be incorporated into RNA 5′ ends. Tran-
scription reactions were performed by T7 RNA polymerase
from a DNA template containing the T7 Aϕ2.5 promoter
sequence, defining A as the first transcribed nucleotide and G
as the second (Figure 3A).19 Transcripts were trimmed at the
3′ end by DNAzyme 10−23 to decrease the 3′-end
heterogeneity, followed by PAGE under conditions that
Figure 2. (A) Sample HPLC profiles from the synthesis of NAD
analogues 1a and 1b. (B) Comparison of 31P NMR spectra for
selected NAD analogues.
Figure 3. (A) Schematic representation of transcription initiation
leading to either uncapped or NAD-capped RNA. (B) RNAs (25 nt)
produced by IVT in the presence of different NAD analogues
analyzed by PAGE (SybrGold staining was used for the visualization).
C
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enabled separation of NAD-RNAs from uncapped (ATP-
initiated) RNAs (Figure 3B). Analogues 1a and 1b were
incorporated into RNA with efficiencies comparable to that for
unmodified NAD with good 5′-end homogeneity.
carrying analogue 2 was also susceptible to the action of DXO,
and despite noticeable 5′-end inhomogeneity at time point 0, it
behaved similarly to NAD-capped RNA. As expected, RNA
carrying trinucleotide 3 at the 5′ end was equally readily
degraded by DXO as RNA capped with unmodified NAD
(Figure 4B). RNA capped with trinucleotides 4a and 4b
containing a phosphorothioate moiety at the site of DXO-
mediated cleavage showed notably diminished susceptibility to
DXO. However, partial degradation was clearly visible at
higher enzyme concentration (50 nM), especially for stereo-
isomer D2. Similarly, 2′-O-methylated NAD-capped RNA
obtained with analogue 5 was notably less susceptible but still
partially degraded by DXO at the highest concentration. The
combination of 2′-O-Me and α-PS modifications turned out to
be additive, as RNAs capped with analogues 6a and 6b were
less susceptible to DXO than the corresponding RNAs carrying
NADs with only single modification (4a, 4b, or 5). As a result,
transcripts capped with 6a and 6b were the most resistant to
DXO activity, with almost negligible degradation even at the
highest DXO concentration, and thus were assigned as “DXO-
resistant” (Table 1).
The 5′-triphosphorylated fraction of these RNAs constituted
less than 40% of the total RNA. In the case of analogue 2, a
higher portion of uncapped RNA was observed, indicating that
the analogue was less efficiently recognized by T7 polymerase.
In contrast, the unmodified trinucleotide 3 was a very efficient
transcription initiator because uncapped RNA was at a
negligible level. Notably, the efficiency of trinucleotide NAD
analogue incorporation was superior to that of dinucleotide
NAD analogue incorporation19 and of the previously reported
method for chemical addition of nicotinamide riboside 5′-
monophosphate to 5′-phosphorylated RNA.20 Therefore, the
use of trinucleotide 3 may be the preferred method for NAD-
RNA synthesis when RNA with high 5′-end homogeneity is
required for biochemical experiments. Trinucleotides 4a, 4b, 5,
6a, and 6b were also efficiently incorporated into RNA,
indicating that the introduced modifications did not affect the
interaction with T7 polymerase.
Next, we assessed the susceptibility of RNAs capped with
various NAD analogues to previously described deNADding
enzymes, including two NUDIX-family pyrophosphatases
(NudC from E. coli and murine Nudt12)8−10 and a
phosphodiesterase (murine DXO).4 Recombinant proteins
were incubated with IVT 32P-labeled RNA, and the resulting
products were resolved using PAGE to assess the remaining
amount of NAD-capped RNA (Figures 4−6). RNA capped
with unmodified NAD was efficiently deNADded by DXO to
RNA 5′-monophosphate and further degraded by its 5′ → 3′
exonuclease activity, which was particularly well manifested at
the highest enzyme concentration (50 nM) (Figure 4A).
Table 1. Enzymatic Susceptibilities of Different NAD-
Capped RNAs
b
susceptibility to deNADding by
compound at the RNA
a
entry
5′ end
DXO
cleaved
partially resistant
partially resistant
cleaved
NudC
NUDt12
1
2
3
4
5
6
7
8
9
NAD
1a
1b
2
cleaved
resistant
resistant
resistant
cleaved
cleaved
resistant
resistant
resistant
cleaved
n.d.
n.d.
cleaved
resistant
resistant
3
cleaved
c
c
4a
4b
5
6a
6b
partially resistant
partially resistant
partially resistant
resistant
n.d.
c
c
n.d.
cleaved
resistant
resistant
10
resistant
a
b
Incorporated by IVT with polymerase T7; “cleaved” indicates that
the RNA susceptibility was similar to that of unmodified NAD-capped
RNA; “partially resistant” indicates that the susceptibility was reduced
but cleavage was observed at higher enzyme concentrations;
“resistant” indicates that cleavage was not observed at the
c
concentrations tested. The analogue was not investigated in this
assay.
Next, we assessed the susceptibility of modified RNAs to
pyrophosphate bond cleavage mediated by NudC and Nudt12.
RNAs uniformly labeled with 32P and carrying different NAD
analogues were exposed to increasing concentrations of NudC
(Figure 5) or Nudt12 (Figure 6).
For NudC, RNA capped with unmodified NAD, regardless
of the synthesis method (using NAD dinucleotide or
trinucleotide 3), was completely deNADded even at the
lowest enzyme concentration tested (50 nM). Similar
susceptibility to degradation was observed for analogue 5,
which was in agreement with our expectation that
modifications outside the pyrophosphate moiety would not
influence the susceptibility to pyrophosphatase activity. All of
the analogues carrying α-PS or PSL modifications within the
pyrophosphate were resistant to NudC at concentrations up to
250 nM (Table 1).
Figure 4. Susceptibility of RNAs capped with NAD analogues to
DXO. A schematic of NAD-capped RNA hydrolysis by DXO is shown
at the left. Reaction products of in vitro deNADding assays with
mouse DXO protein and the indicated 32P-labeled RNAs capped with
(A) NAD dinucleotide analogues or (B) NAD trinucleotide analogues
were resolved on 20% urea polyacrylamide gels.
As expected, RNAs capped with analogues 1a and 1b were
also efficiently hydrolyzed, although a higher concentration of
the enzyme was required. Interestingly, the transient/initial
deNADding product was not observed, suggesting a reduced
efficiency of deNADding with a subsequent rapid decay of the
deNADded product by the DXO 5′ → 3′ exonuclease activity
compared with RNA capped with unmodified NAD. RNA
D
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recognize the 5′-terminal NAD moiety in RNA as a molecular
hallmark.
ASSOCIATED CONTENT
* Supporting Information
■
S
The Supporting Information is available free of charge on the
ACS Publications website at DOI: 10.1021/acs.or-
glett.8b03386.
Experimental procedures and spectroscopic data (PDF)
Figure 5. Susceptibility of RNAs capped with (A) NAD dinucleotide
analogues or (B) NAD trinucleotide analogues to NudC. A schematic
of NAD-capped RNA hydrolysis by NudC is shown at the left.
Reaction products of NAD-capped RNA were resolved as in Figure 4.
AUTHOR INFORMATION
■
Corresponding Author
*E-mail: jkowalska@fuw.edu.pl
ORCID
Jacek Jemielity: 0000-0001-7633-788X
Joanna Kowalska: 0000-0002-9174-7999
Author Contributions
∥A.-M.C. and A.D. contributed equally.
Notes
The authors declare no competing financial interest.
ACKNOWLEDGMENTS
Figure 6. Susceptibility of RNAs capped with (A) NAD dinucleotide
analogues or (B) NAD trinucleotide analogues to Nudt12. A
schematic of NAD-capped RNA hydrolysis by NudC is shown at
the left. Reaction products of NAD-capped RNA were resolved as in
Figure 4.
■
We thank M. Baranowski (University of Warsaw) for recording
NMR spectra and B. E. Nickels (Rutgers University) for
providing recombinant NudC. This work was supported by
funding from the National Science Centre, Poland (2015/18/
E/ST5/00555 to J.K.), the Foundation for Polish Science
(TEAM/2016-2/13 to J.J.), and the National Institutes of
Health (Grant GM126488 to M.K.).
Mouse Nudt12 showed a similar substrate preference as
NudC, i.e., RNAs capped with NAD, analogue 3, and 5 were
susceptible to cleavage, whereas all of the other tested
analogues were resistant (Table 1).
REFERENCES
■
́
(1) Berger, F.; Ramírez-Hernandez, M. H.; Ziegler, M. The new life
of a centenarian: signalling functions of NAD(P). Trends Biochem. Sci.
2004, 29, 111−118.
In conclusion, we synthesized and biochemically charac-
terized eight novel NAD analogues that enabled the generation
of NAD-capped RNAs with reduced susceptibility to
enzymatic cleavage. These analogues can be incorporated
into RNA using a simple IVT reaction to produce modified
NAD-capped RNAs. The use of the trinucleotide NRppApG
(3) or its analogues (4−6) in IVT improved the efficiency of
RNA capping compared with initiation with NAD, which
makes the synthesized analogues particularly useful for
obtaining NAD-capped RNAs with superior 5′-end homoge-
neity. Studies of the susceptibility of RNAs with different NAD
caps to deNADding enzymes revealed compounds with
distinct characteristics. Among the eight new chemically
modified NAD analogues, we identified a compound that
conferred resistance to the pyrophosphatase activity of NudC/
Nudt12 on RNA but kept it susceptible to the phosphodiester-
ase activity of DXO (compound 2), a compound that was
susceptible to pyrophosphatases but conferred reduced
susceptibility to DXO (compound 5), and compounds that
conferred reduced (to various degrees) susceptibility to both
types of enzymes (compounds 1a, 1b, 6a, and 6b). Thus, the
chemical toolbox of our NAD analogues should be particularly
useful in biochemical studies of the contributions of different
deNADding enzymes to overall NAD-RNAs decay given that
in human cells different RNA sequences are targeted by DXO
and Nudt12.10 Other potential applications of the synthesized
analogues include studies of biological functions of NAD-
RNAs and identification of putative effector proteins that
(2) Chen, Y. G.; Kowtoniuk, W. E.; Agarwal, I.; Shen, Y.; Liu, D. R.
LC/MS analysis of cellular RNA reveals NAD-linked RNA. Nat.
Chem. Biol. 2009, 5, 879−881.
(3) (a) Luciano, D. J.; Belasco, J. G. NAD in RNA: unconventional
headgear. Trends Biochem. Sci. 2015, 40 (5), 245−247. (b) Jaschke,
A.; Hofer, K.; Nubel, G.; Frindert, J. Cap-like structures in bacterial
RNA and epitranscriptomic modification. Curr. Opin. Microbiol. 2016,
30, 44−49. (c) Cahova, H.; Winz, M. L.; Hofer, K.; Nubel, G.;
Jaschke, A. NAD captureSeq indicates NAD as a bacterial cap for a
subset of regulatory RNAs. Nature 2015, 519, 374−377.
(4) Jiao, X.; Doamekpor, S. K.; Bird, J. G.; Nickels, B. E.; Tong, L.;
Hart, R. P.; Kiledjian, M. 5′ End Nicotinamide Adenine Dinucleotide
Cap in Human Cells Promotes RNA Decay through DXO-Mediated
deNADding. Cell 2017, 168 (6), 1015−1027.
(5) (a) Walters, R. W.; Matheny, T.; Mizoue, L. S.; Rao, B. S.;
Muhlrad, D.; Parker, R. Identification of NAD+ capped mRNAs in
Saccharomyces cerevisiae. Proc. Natl. Acad. Sci. U. S. A. 2017, 114 (3),
480−485. (b) Frindert, J.; Zhang, Y. Q.; Nubel, G.; Kahloon, M.;
Kolmar, L.; Hotz-Wagenblatt, A.; Burhenne, J.; Haefeli, W. E.;
Jaschke, A. Identification, Biosynthesis, and Decapping of NAD-
Capped RNAs in B. subtilis. Cell Rep. 2018, 24, 1890−1901.
(6) Bird, J. G.; Basu, U.; Kuster, D.; Ramachandran, A.; Grudzien-
Nogalska, E.; Kiledjian, M.; Temiakov, D.; Patel, S. S.; Ebright, R. H.;
Nickels, B. E. Mitochondrial RNA capping: highly efficient 5′-RNA
capping with NAD+ and NADH by yeast and human mitochondrial
RNA polymerase. 2018, bioRχiv 381160. bioRχiv, The Preprint
Server for Biology. DOI: 10.1101/381160.
(7) Kiledjian, M. Eukaryotic RNA 5′-End NAD+ Capping and
DeNADding. Trends Cell Biol. 2018, 28 (6), 454−464.
E
DOI: 10.1021/acs.orglett.8b03386
Org. Lett. XXXX, XXX, XXX−XXX

Organic Letters
Letter
(8) Hofer, K.; Li, S. S.; Abele, F.; Frindert, J.; Schlotthauer, J.;
Grawenhoff, J.; Du, J. M.; Patel, D. J.; Jaschke, A. Structure and
function of the bacterial decapping enzyme NudC. Nat. Chem. Biol.
2016, 12, 730−734.
(9) Zhang, D. L.; Liu, Y. X.; Wang, Q.; Guan, Z. Y.; Wang, J.; Liu, J.;
Zou, T. T.; Yin, P. Structural basis of prokaryotic NAD-RNA
decapping by NudC. Cell Res. 2016, 26, 1062−1066.
(10) Grudzien-Nogalska, E.; Jiao, Y.; Wu, X.; Cui, H.; Hart, R. P.;
Tong, L.; Kiledjian, M. 2018, Biorxiv, https://doi.org/10.1101/
474478.
(11) (a) Winz, M. L.; Cahova, H.; Nubel, G.; Frindert, J.; Hofer, K.;
Jaschke, A. Capture and sequencing of NAD-capped RNA sequences
with NAD captureSeq. Nat. Protoc. 2017, 12, 122−149. (b) Halle, F.;
Fin, A.; Rovira, A. R.; Tor, Y. Emissive Synthetic Cofactors:
Enzymatic Interconversions of tzA Analogues of ATP, NAD+,
NADH, NADP+, and NADPH. Angew. Chem., Int. Ed. 2018, 57,
1087−1090. (c) Vvedenskaya, I. O.; Bird, J. G.; Zhang, Y. C.; Zhang,
Y.; Jiao, X. F.; Barvik, I.; Krasny, L.; Kiledjian, M.; Taylor, D. M.;
Ebright, R. H.; Nickels, B. E. CapZyme-Seq Comprehensively Defines
Promoter-Sequence Determinants for RNA 5′ Capping with NAD+.
Mol. Cell 2018, 70, 553−564. (d) Grudzien-Nogalska, E.; Bird, J. G.;
Nickels, B.; Kiledjian, M. NAD-capQ” detection and quantitation of
NAD caps. RNA 2018, 24, 1418−1425.
(12) (a) Grudzien-Nogalska, E.; Jemielity, J.; Kowalska, J.;
Darzynkiewicz, E.; Rhoads, R. E. Phosphorothioate cap analogs
stabilize mRNA and increase translational efficiency in mammalian
cells. RNA 2007, 13, 1745−1755. (b) Kowalska, J.; Lewdorowicz, M.;
Zuberek, J.; Grudzien-Nogalska, E.; Bojarska, E.; Stepinski, J.; Rhoads,
R. E.; Darzynkiewicz, E.; Davis, R. E.; Jemielity, J. Synthesis and
characterization of mRNA cap analogs containing phosphorothioate
substitutions that bind tightly to eIF4E and are resistant to the
decapping pyrophosphatase. RNA 2008, 14, 1119−1131. (c) Wojtc-
zak, B. A.; Sikorski, P. J.; Fac-Dabrowska, K.; Nowicka, A.; Warminski,
M.; Kubacka, D.; Nowak, E.; Nowotny, M.; Kowalska, J.; Jemielity, J.
5′-Phosphorothiolate Dinucleotide Cap Analogues: Reagents for
Messenger RNA Modification and Potent Small-Molecular Inhibitors
of Decapping Enzymes. J. Am. Chem. Soc. 2018, 140 (18), 5987−
5999.
(13) Picard-Jean, F.; Brand, C.; Tremblay-Letourneau, M.; Allaire,
A.; Beaudoin, M. C.; Boudreault, S.; Duval, C.; Rainville-Sirois, J.;
Robert, F.; Pelletier, J.; Geiss, B. J.; Bisaillon, M. PLoS One 2018, 13
(3), e0193804.
(14) Tanimori, S.; Ohta, T.; Kirihata, M. An efficient chemical
synthesis of nicotinamide riboside (NAR) and analogues. Bioorg. Med.
Chem. Lett. 2002, 12, 1135−1137.
(15) Preiss, J.; Handler, P. Enzymatic synthesis of nicotinamide
mononucleotide. J. Biol. Chem. 1957, 225, 759−770.
(16) Liu, R. H.; Visscher, J. A Novel Preparation of Nicotinamide
Mononucleotide. Nucleosides Nucleotides 1994, 13, 1215−1216.
(17) Moran, J. R.; Whitesides, G. M. A practical enzymic synthesis of
(SP)-adenosine 5′-O-(1-thiotriphosphate) ((SP)-ATP-α-S). J. Org.
Chem. 1984, 49, 704−706.
(18) Patel, B. K.; Eckstein, F. 5′-Deoxy-5′-thioribonucleoside-5′-
triphosphates. Tetrahedron Lett. 1997, 38, 1021−1024.
(19) Huang, F. Efficient incorporation of CoA, NAD and FAD into
RNA by in vitro transcription. Nucleic Acids Res. 2003, 31 (3), e8.
(20) Hofer, K.; Abele, F.; Schlotthauer, J.; Jaschke, A. Synthesis of
5′-NAD-Capped RNA. Bioconjugate Chem. 2016, 27, 874−877.
F
DOI: 10.1021/acs.orglett.8b03386
Org. Lett. XXXX, XXX, XXX−XXX