Design, synthesis, and incorporation of fluorous 5-methylcytosines into oligonucleotides

Article abstract of DOI:10.1021/jo2015399

A palladium-catalyzed Negishi coupling reaction has been developed to synthesize fluorous 5-methyl-cytosines. These fluorous nucleosides are incorporated into the oligonucleotides that correspond to part of the promoter region of Oct4, a master gene that undergoes dynamic DNA demethylation during cellular reprogramming. The separation of the fluorous oligonucleotides from its nonfluorous analogues has been achieved through solid-phase extraction over fluorous silica, suggesting its potential use in probing DNA demethylation (Figure presented).

Full text of DOI:10.1021/jo2015399

Note
pubs.acs.org/joc
Design, Synthesis, and Incorporation of Fluorous 5-Methylcytosines
into Oligonucleotides
Zhiquan Song and Qisheng Zhang*
Division of Chemical Biology and Medicinal Chemistry, The University of North Carolina at Chapel Hill, Chapel Hill, North Carolina
27599, United States
S
* Supporting Information
ABSTRACT: A palladium-catalyzed Negishi coupling reac-
tion has been developed to synthesize fluorous 5-methyl-
cytosines. These fluorous nucleosides are incorporated into the
oligonucleotides that correspond to part of the promoter
region of Oct4, a master gene that undergoes dynamic DNA
demethylation during cellular reprogramming. The separation
of the fluorous oligonucleotides from its nonfluorous
analogues has been achieved through solid-phase extraction over fluorous silica, suggesting its potential use in probing DNA
demethylation.
eoxyribonucleic acid (DNA) methylation and demethy-
lation of cytosine at the C5 position play important roles
nature of living cells makes it difficult to trace DNA
demethylation reactions endogenously.
D
in many fundamental biological processes including tran-
scription regulation, stem cell pluripotency, and cancer.1 In
early embryogenesis, dynamic change of DNA methylation and
demethylation is one key event in nuclear reprogramming and
genomic imprinting. Aberrant regulation of this process leads to
developmental defects and plays a role in carcinogenesis.
Consequently, understanding the mechanisms of DNA
methylation and demethylation is important in illustrating the
epigenetic and genetic regulation of normal and disease
development and could have potential therapeutic applications.
Various DNA methyltransferases have been identified, and
their catalysis has been illustrated in detail.2 In contrast, the
enzymes that are responsible for the reverse process, DNA
demethylation of 5-methylcytosine (5mC), have not been
identified. Consequently, the mechanism of active DNA
demethylation is not clear, and three possibilities have been
proposed (Figure 1A). The demethylation process could occur
through direct C−C bond cleavage in pathway A, the glycosate
cleavage or deamination followed by a base excision repair in
pathway B, or dinucleotide replacement in pathway C. Enzymes
that are responsible for glycosate cleavage for 3-methylcytosine
(3mC) and dinucleotide replacement have been identified in
plants,3 but such mechanisms are unlikely to be present in
mammalian cells where analogous glycosylases do not exist.
Furthermore, any mistake from genome-wide DNA repair in
one-cell embryos in mammalian systems is likely detrimental.
Recently, ten eleven translocation (Tet) proteins have been
demonstrated to convert 5mC to 5-hydroxymethylcytosine
(5hmC), 5-formylcytosine (5fC), and 5-carboxylcytosine
(5caC).4 However, it is still unclear how Tet proteins control
the relative ratio of these species and how they are further
converted to cytosine. To elucidate the demethylation events, it
will be helpful to analyze the biochemistry of the demethylation
reaction actually taking place in vivo. However, the complex
Small molecules with highly fluorinated domains tend to
preferentially partition into the fluorocarbon-enriched phase of
a biphase or triphase system, partially due to the fluorine−
fluorine interactions.5 This fact forms the foundation of the
recently established fluorous chemistry. The word “fluorous” is
coined to describe the phase formed from highly fluorinated
molecules in analogy to an aqueous phase.6 Small molecules
with highly fluorinated domains are called fluorous compounds
or molecules. Initially, highly fluorinated (fluorous) solvents are
used to separate fluorous from nonfluorous compounds
through fluorous−organic liquid−liquid extraction.7 Later,
fluorous-functionalized silica gel and the fluorous solid-phase
extraction (FSPE) technique are used to further improve the
efficiency of separation.8 Typically, the reaction mixture is
loaded on a column with fluorous silica. The organic products
are eluted with organic solvents and the fluorous products with
fluorous solvents. FSPE has since been employed for the
recycling and reuse of catalysts,9 removal of reaction
intermediates,10 and fluorous mixture synthesis of libraries of
compounds.11 Recently, fluorinated peptides were demonstra-
ted to be efficiently separated from nonfluorinated peptides
through FSPE.12 The fluorous protecting groups have also
aided in the purification of oligosaccarides and oligonucleotides.
In addition, a single C8F17 group renders the easy separation of
up to 100-membered oligonucleotides from nonfluorinated
nucleotides or reagents in the mixture.13 These successes
prompt us to explore the applications of fluorous unnatural
DNAs in uncovering the mechanisms of DNA demethylation.
The concept of using fluorous 5mC derivatives to probe
DNA demethylaiton is illustrated in Figure 1B. Fluorinated 5-
Received: July 25, 2011
Published: November 14, 2011
© 2011 American Chemical Society
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Note
Figure 1. Fluorous approach to DNA demethylation. (A) Three possible mechanisms for DNA demethylation. (B) Schematic illustration of using
fluorous 5mC derivatives in DNA demethylation.
Scheme 1. Synthesis of Fluorous 5-Methylcytosine Derivatives 7a−c and 8a−c
methylcytosines 1 are synthesized and incorporated into a
sequence in a selected promoter region containing the CpG
unit of genes that are known to be regulated by DNA
methylation/demethylation. The resulting oligonucleotides 2
(XCFpGY) then are annealed with their complementary
oligonucleotides to form double-stranded DNAs (dsDNAs) 3
that will be introduced into cells. The resulting mixture then
will be applied to a fluorous chromatographic support to retain
the fluorinated compounds selectively. Elution with more
fluorophilic solvents affords the enriched fraction containing
products 4 derived directly from fluorinated unnatural DNAs.
Separation and analysis of these enriched products by HPLC-
MS will provide novel insights into the mechanisms of DNA
demethylation. Because the fluorous tags are chemically inert
and highly stable in tandem mass spectrometric analysis, the
DNA demethylation products of fluorous oligonucleotides will
be analyzed without the interference of other molecules in the
cells, thus representing a novel approach to understand the
biochemistry of DNA demethylation in vivo. For example, if the
demethylation reaction proceeds as in pathway A in Figure 1A,
then fluorinated small molecules will be enriched. Identification
of the enriched small molecules will provide insights into the
mechanisms of DNA demethylation. Similarly, if pathway B
dominates, then fluorinated cytosine will be recovered in the
fluorous fraction. In this work, we will report the results on
design, synthesis, and incorporation of fluorous 5-methylcyto-
sines into oligonucleotides.
To selectively enrich the products derived from DNA
demethylation, we envisioned that a fluorous tag (Rf) could be
attached to the 5-methyl group. The optimal size of the Rf
should be sufficient to separate the tagged components from
the untagged ones, while the DNA demethylation process is
minimally interfered. Accordingly, several 5mC analogues with
different Rf groups were synthesized. The modification at the
C5 position of cytosine typically starts with either 2′-
deoxyuridine or 5-iodocytosine. When 2′-deoxyuridine is used
as the starting material, multiple-step reactions14 that include
nucleophilic addition to aldehydes and conversion of
deoxyuridine to cyctosine are applied. The nucleophilic
addition reaction requires harsh conditions and generally
provides low yields.14 In addition, the hydroxyl group in the
addition product has to be removed. Consequently, this
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Note
Figure 2. Incorporation of fluorous 5-methylcytosines into oligonucleotides. (A) Structure of synthetic oligonucleotides. (B) Representative MS and
LC chromatogram of 9c.
Figure 3. Separation of fluorous from nonfluorous oligonucleotides. A mixture of 9d with 9a (A), 9b (B), or 9c (C) was separated through FSPE
into two fractions. Both fractions were analyzed by HPLC. Elution solvent for the first fraction was 10% MeCN in 0.1 M TEAA, while that for the
second fraction was 20% MeCN in 0.1 M TEAA (A), 25% MeCN in 0.1 M TEAA (B), and 30% MeCN in 0.1 M TEAA (C). The flow rate was 1
mL/min.
strategy is not efficient for synthesis of fluorous 5mC
derivatives. A more straightforward approach uses 5-iodocyto-
sine as the starting material. Sonagashira reaction or Heck
coupling with alkynes or alkenes has been used to synthesize
the corresponding modified cytosines. Although not reported,
further reduction of the coupling products with hydrogen
should provide 5-alkylated cytosines. However, the fluorous
terminal alkynes or alkenes are not readily available, making
additional synthetic efforts necessary. In contrast, fluorous
alkyliodides are commercially available and easy to handle.
Consequently, we decided to develop a palladium-catalyzed
coupling reaction between the 5-iodocytosine with organozinc
reagents to facilitate the synthesis of the modified 5-
methylcytosines.
0.6 M as determined by titration with iodine.16 Among several
ligands and metal complexes tested, the use of [1,1′-
bis(diphenylphosphino)ferrocene]dichloropalladium
(PdCl2(dppf)) and copper iodide (CuI) in the presence of 0.5
equiv of trifluoromethylbenzene (PhCF3) gave 41% yield of the
desired product when 6b was used (Scheme 1). These
conditions were also used for two other fluorous zinc reagents
6a and 6c to form the corresponding coupling products 7a and
7c, respectively. The triisopropylsilyl (TIPS) protective groups
in 7a−c were then removed by treatment with tetrabutylam-
monium fluoride (TBAF). In order to apply the standard
oligonucleotide synthesis protocol for the commercial nucleic
acid synthesizer,17 the 5′-hydroxyl group is protected as the 4,4′-
dimethoxytrityl (DMT) ether by reacting with DMT chloride,
and the 3′-hydroxyl group is converted to the phosphoramidite
monomer 8a−c by reacting with 2-cyanoethyl N,N-diisopropyl
chlorophosphine according to the literature protocol.17a,c
The organozinc reagents derived from the fluorinated iodide
6a−c were prepared according to the Knochel protocol.15 We
typically prepared fluorous zinc reagents in the concentration of
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Note
Oct4 is one master gene that regulates the differentiation and
pluripotency of embryonic stem cells.18 In the cellular
reprogramming in vivo and in vitro, the expression of Oct4 is
turned on and DNA demethylations of the reprogrammed cell
markers take place. For example, the CpG unit in the sequence
5′-GCAATCCGGTAG is predominantly in the demethylated
state during cellular reprogrammng. The fluorous building
blocks 8a−c were thus incorporated into the CpG region of
this sequence in Oct4 promoter according to the standard
protocol. Briefly, all oligonucleotides are synthesized by
standard phosphoramidite techniques and deprotected at 52
°C in saturated ammonia hydroxide for 5 h. The resulting
mixtures are then purified by HPLC to provide oligonucleo-
tides 9a−c (Figure 2A). As a control, 5-methylcytosine was also
incorporated into this sequence to generate oligonucleotide 9d.
The purity and identity of the oligonucleotide were determined
by reverse-phase HPLC and matrix-assisted laser desorption/
ionization time-of-flight (MALDI-TOF) mass spectrometry,
respectively. The calculated and measured molecular weights of
9a−d are listed in Figure 2A. One representative HPLC
chromatogram of 9c is shown in Figure 2B with the MS
spectrum of 9c in the inset. These results suggest that the
desired oligonucleotides were obtained with high purity
(>95%).
The fluorinated oligonucleotides 9a−c were then annealed
with the antisense pair 5′-CTACCGGATTGC to generate
double-stranded DNAs 10a−c. Similarly, the nonfluorinated
oligonucleotide 9d was also annealed to form double-stranded
10d as a control. Next, the melting temperatures (Tm) of 10a−
d were measured (Figure 2A) to investigate the effect of
fluorous tags. The Tm values decrease as the size of the fluorous
tag increases, suggesting that the fluorous tags interfere with
base pairing. Nonetheless, the Tm values of 10a−d were all in
the range of 53−58 °C, indicating that stable double helix
structures were formed for all double-stranded DNAs.
One key feature of using fluorous 5mC derivatives is to
enrich the fluorous-tagged products. To test the separation
efficiency of the fluorous 5mC derivatives, 9a was mixed with
9d and the resulting mixture was loaded onto a column with
fluorous silica. Elution with the column with 10% CH3CN in
0.1 M triethylammonium acetate (TEAA) buffer yielded the
first fraction. The second fraction was obtained by eluting the
column with 100% CH3CN. Both fractions were then analyzed
by HPLC (Figure 3) to assess the separation efficiency. The
same sets of experiments were carried out for 9b and 9c. As
expected, only the nonfluorous 9d was eluted when 10%
CH3CN in 0.1 M TEAA buffer was applied. In contrast, elution
with 100% CH3CN provides 9a−c. While the separation of 9a
(Rf = C2F5) from 9d requires careful elution, 9b (Rf = C6F13)
and 9c (Rf = C8F17) can be easily separated from the
nonfluorous 9d.
In summary, we have developed fluorous 5-methylcytosine
derivatives and successfully incorporated them into one CpG
region that exists in the Oct4 gene promoter. The modified
oligonucleotides are effectively separated from the nonfluorous
oligonucleotide with 5-methylcytosine incorporated. We
acknowledge that these fluorous oligonucleotides have not
been tested in DNA demethylation reactions in cells, which
requires extensive studies on suitable oligonucleotide sequences
and cellular systems. Nonetheless, the fact that fluorous 5mC
derivatives can be incorporated into oligonucleotides and
separated from nonfluorous species has laid the foundation to
investigate the cellular mechanism of DNA demethylation
through enriching and analyzing the reaction products.
EXPERIMENTAL SECTION
■
All of the solvents were purchased from suppliers as anhydrous grade.
1H and 13C NMR spectra were recorded at room temperature in
CDCl3 (containing 1% TMS) solutions on a 300 or 400 MHz
spectrometer. HPLC analyses were performed on reverse-phase
columns. Low- and high-resolution mass spectra were obtained from
a Qh-FTICR mass spectrometer.
Synthesis of Compound 5. Imidazole (1.59 g, 23.32 mmol) and
DMAP (64.8 mg, 0.53 mmol) were added to a solution of 5-iodo-2′-
deoxycytidine (1.87 g, 5.30 mmol) in dry DMF (25 mL) at room
temperature. Subsequently, triisopropylsilyl trifluoromethanesulfonate
(3.57 g, 11.66 mmol) was added dropwise. After stirring for 16 h at
room temperature, saturated NH4Cl solution was added. The mixture
was extracted with chloroform three times. The combined organic
layers were dried over sodium sulfate and concentrated under vacuum
to give a crude product of 5-iodo-3′,5′-O-bis(triisopropylsilyl)-2′-
deoxycytidine, which was resuspended in dry pyridine (25 mL).
DMAP (64.80 mg, 0.53 mmol) and benzoic anhydride (2.40 g, 10.60
mmol) were then added. After stirring at 35 °C for 5 h, the pyridine
was removed under reduced pressure. The residue was purified
through flash chromatography (hexanes/EtOAc = 9:1) to yield
1
compound 5 as light yellow solid (3.26 g, 80%): H NMR (CDCl3,
300 MHz) δ 1.01−1.31 (m, 42H), 2.00−2.09 (m, 1H), 2.45 (dd, J =
12.0, 5.1 Hz, 1H), 3.89 (dd, J = 10.8, 2.7 Hz, 1H), 3.99 (dd, J = 10.8,
2.7 Hz, 1H), 4.09 (d, J = 1.5 Hz, 1H), 4.62 (d, J = 5.4 Hz, 1H), 6.30
(dd, J = 9.0, 5.1 Hz, 1H), 7.42−7.47 (m, 2H), 7.51−7.56 (m, 1H),
8.19 (s, 1H), 8.38 (d, J = 6.9 Hz, 1H); 13C NMR (CDCl3, 75 MHz) δ
11.9, 12.0, 17.92, 17.94, 18.1, 42.6, 63.5, 69.4, 73.0, 86.3, 89.1, 128.1,
130.11, 132.6, 136.6, 145.2, 147.2, 156.7, 179.6; MS (ESI) m/z 770.3
[M + H]+.
Compound 7b. Trifluoromethyl benzene (4.3 μL, 35 μM) was
added to a solution of 5 (54 mg, 70 μmol) in anhydrous THF (0.3
mL). The reaction solution was then heated to 50 °C. PdCl2(dppf)-
(CH2Cl2) (5.7 mg, 7 μmol) was added at the same temperature. After
stirring for 5 min, 3,3,4,4,5,5,6,6,7,7,8,8,8-tridecafluorooctyl zinc iodide
(0.18 mL, 0.6 M) was added dropwise, followed by the addition of CuI
(13.3 mg, 70 μmol). The stirring was continued at the same
temperature for 3 h. Another portion of the zinc reagent (0.18 mL, 0.6
M) was then added, and the reaction mixture was heated to 60 °C.
After 14 h, the reaction mixture was cooled to room temperature and
added into water, extracted with EtOAc three times. The combined
organic fractions were dried over sodium sulfate. Evaporation of the
solvent and flash chromatography (hexanes/EtOAc = 95:5) gave the
1
compound 7b as colorless oil (28 mg, 41%): H NMR (CDCl3, 300
MHz) δ 1.01−1.23 (m, 42H), 1.99−2.08 (m, 1H), 2.41 (dd, J = 12.9
Hz, 5.1 Hz, 1H), 2.44−2.56 (m, 2H), 2.82 (t, J = 8.1 Hz, 2H), 3.91
(dd, J = 11.1, 2.7 Hz, 1H), 3.41 (dd, J = 11.4, 2.1 Hz, 1H), 4.08 (d, J =
2.1 Hz, 1H), 4.63 (d, J = 5.4 Hz, 1H), 6.38 (dd, J = 8.7, 5.4 Hz, 1H),
7.41−7.46 (m, 2H), 7.51−7.56 (m, 1H), 7.79 (s, 1H), 8.25−8.29 (m,
2H); 13C NMR (CDCl3, 75 MHz) δ 11.8, 12.0, 17.9, 20.2, 30.2 (t, J =
21.6 Hz), 42.3, 63.6, 72.9, 85.6, 88.8, 112.2, 128.2, 129.8, 132.6, 136.9,
138.0, 147.6, 158.8, 179.6; 19F NMR (CDCl3, 282 MHz) δ −81.4 (t, J
= 10.2 Hz), −115.6 (m), −122.2, −123.3, −124.1, −126.5; MS (ESI)
m/z 990.3 [M + H]+; HRMS (ESI) calcd for C42H61F13N3O5Si2 [M +
H]+ 990.3942, found 990.3938.
Compound 7a. In a similar manner to 7b, 7a (72 mg, 30%) was
synthesized from 6a (1.5 mL, 0.9 mmol) and 5 (233 mg, 0.3 mmol) as
1
colorless oil: H NMR (CDCl3, 300 MHz) δ 0.88−1.23 (m, 42H),
1.98−2.07 (m, 1H), 2.38−2.52 (m, 3H), 2.80 (t, J = 8.1 Hz, 2H), 3.90
(dd, J = 11.4, 2.4 Hz, 1H), 3.99 (dd, J = 11.4, 2.4 Hz, 1H), 4.08 (d, J =
1.8 Hz, 1H), 4.62 (d, J = 5.4 Hz, 1H), 6.38 (dd, J = 8.7, 5.4 Hz, 1H),
7.42−7.47 (m, 2H), 7.51−7.56 (m, 1H), 7.78 (s, 1H), 8.25−8.29 (m,
2H); 13C NMR (CDCl3, 75 MHz) δ 11.8, 12.0, 17.91, 17.93, 20.3, 30.0
(t, J = 21.4 Hz), 42.3, 63.6, 72.9, 85.6, 88.8, 112.3, 128.2, 129.8, 132.6,
136.9, 137.9, 147.6, 158.8, 179.5; 19F NMR (CDCl3, 282 MHz) δ
−85.9, −119.1 (t, J = 17.8 Hz); MS (ESI) m/z 790.3 [M + H]+;
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Note
HRMS (ESI) calcd for C38H60CsF5N3O5Si2 [M + Cs]+ 922.305, found
922.303.
Compound 7c. In a similar manner to 7b, 7c (116 mg, 35%) was
135.05, 135.06, 135.12, 136.9, 138.2, 138.3, 144.0, 147.6, 158.8, 179.5;
19F NMR (CDCl3, 282 MHz) δ −86.0 (d, J = 5.4 Hz), −119.0 (m);
31P NMR (CDCl3, 121 MHz) δ 149.6, 150.0; MS (ESI) m/z 980.3 [M
+ H]+, HRMS (ESI) calcd for C50H55CsF5N5O8P [M + Cs]+
1112.276, found 1112.289.
synthesized from 6c (1.5 mL, 0.9 mmol) and 5 (233, 0.3 mmol) as
1
white solid: H NMR (CDCl3, 300 MHz) δ 1.00−1.23 (m, 42H),
1.99−2.08 (m, 1H), 2.42 (dd, J = 12.6, 5.1 Hz, 1H), 2.44−2.56 (m,
2H), 2.82 (t, J = 8.1 Hz, 2H), 3.90 (dd, J = 11.4, 2.4 Hz, 1H), 4.00 (dd,
J = 11.4, 2.4 Hz, 1H), 4.08 (d, J = 1.8 Hz, 1H), 4.63 (d, J = 5.1 Hz,
1H), 6.38 (dd, J = 8.7, 5.4 Hz, 1H), 7.41−7.46 (m, 2H), 7.51−7.56
(m, 1H), 7.79 (s, 1H), 8.25−8.28 (m, 2H); 13C NMR (CDCl3, 75
MHz) δ 11.8, 12.0, 17.89, 17.92, 17.94, 20.2, 30.2 (t, J = 21.7 Hz),
42.3, 63.6, 72.9, 85.6, 88.8, 112.3, 128.2, 129.8, 132.6, 136.9, 138.0,
147.6, 158.8, 179.6; 19F NMR (CDCl3, 282 MHz) δ −81.3 (t, J = 10.5
Hz), −115.3, −121.9, −122.1, −122.9, −123.9, −126.3; MS (ESI) m/z
1090.3 [M + H]+; HRMS (ESI) calcd for C44H60CsF17N3O5Si2 [M +
Cs]+ 1222.285, found 1222.284.
Compound 8c. In a similar manner to 8b, 8c (62 mg, 52%) was
synthesized from 7c (103 mg, 94 μmol) as a light yellow foam that
1
existed as a 2:3 mixture of two stereoisomers: H NMR (CDCl3, 300
MHz) δ 1.06−1.09 (2H), 1.14−1.25 (m, 10H), 1.84−2.17 (m, 2H),
2.37−2.51 (m, 4H), 2.57−2.67 (m, 2H), 3.34−3.40 (m, 1H), 3.53−
3.72 (m, 4H), 3.76−3.78 (m, 6H), 3.83−3.87 (m, 1H), 4.20−4.26 (m,
1H), 4.70−4.75 (m, 1H), 6.42 (dd, J =13.2, 6.3 Hz, 1H), 6.82−6.86
(m, 4H), 7.24−7.31 (m, 7H), 7.35−7.44 (m, 4H), 7.49−7.54 (m, 1H),
7.91−7.95 (m, 1H), 8.23 (d, J = 7.8 Hz, 1H); 13C NMR (CDCl3, 75
MHz) δ 19.4, 20.1, 20.2, 20.3, 20.4, 24.39, 24.49, 24.53, 24.61, 29.8 (t,
J = 21.6 Hz), 40.5, 43.1, 43.2, 43.3, 43.4, 55.1, 58.0, 58.1, 58.2, 58.3,
62.9, 63.0, 73.4, 73.6, 73.7, 74.0, 85.5, 85.6, 85.8, 86.1, 86.9, 112.5,
112.6, 113.2, 117.3, 117.5, 127.3, 128.0, 128.1, 128.2, 129.8, 130.0,
130.1, 132.6, 135.0, 135.1, 136.8, 138.3, 138.4, 144.0, 147.62, 147.65,
158.8, 179.6; 19F NMR (CDCl3, 282 MHz) δ −81.3, −115.2, −121.9,
−122.2, −122.9, −123.9, −126.3; 31P NMR (CDCl3, 121 MHz) δ
149.6, 150.0; MS (ESI) m/z 1280.3 [M + H]+; HRMS (ESI) calcd for
C56H55CsF17N5O8P [M + Cs]+ 1412.257, found 1412.272.
Synthesis of Oligonucleotides. The oligonucleotides 9a−d
were synthesized with a DNA synthesizer and cleaved from the
controlled pore glass (CPG) support with concentrated ammonium
hydroxide (3 mL, 2 h). The deprotection was achieved by heating the
resultant ammonium hydroxide solution at 52 °C for 5 h. Purification
of the oligonucleotides was carried out by reverse-phase HPLC using a
C18 column (21.2 mm × 150 mm). The molecular weights of the
oligonucleotides were determined by MALDI-TOF mass spectrosco-
py.
Synthesis of 8b. TBAF (0.09 mL, 1.0 M in THF) was added to a
solution of 7b (40 mg, 40 μmol) in fresh distilled THF (1 mL). After
stirring at room temperature for 30 min, the solvent was removed
under reduced pressure. Flash chromatography (CH2Cl2/MeOH =
1
95:5) gave the deprotected product as white solid (20 mg, 75%): H
NMR (CD3OD, 300 MHz) δ 2.04−2.37 (m, 2H), 2.58−2.73 (m, 2H),
2.86−2.91 (m, 2H), 3.77 (dd, J = 12.3 Hz, J = 3.3 Hz, 1H), 3.87 (dd, J
= 12.3 Hz, J = 2.7 Hz, 1H), 3.95−3.99 (m, 1H), 4.41−4.45 (m, 1H),
6.29 (t, J = 6.3 Hz, 1H), 7.44 (t, J = 7.5 Hz, 2H), 7.56 (t, J = 7.5 Hz,
1H), 8.22−8.27 (m, 3H).
The deprotected product (42 mg, 63 μmol) was dried by first
coevaporation twice with dry pyridine (0.5 mL) and then under
vacuum for 3 h before redissolved in pyridine (0.5 mL) upon heating.
DMAP (1.9 mg, 16 μmol) and DMTrCl (43 mg, 126 μmol) were
added to the resulting solution at 0 °C. The reaction mixture was
warmed to room temperature and stirred for 16 h. The solvent was
then removed under reduced pressure, and the residue was purified by
flash chromatography (CH2Cl2/MeOH = 95:5) to yield DMTr-on
product as a white foam (38 mg, 62%). To a solution of the DMTr-on
compound (38 mg, 39 μmol) in dry CH2Cl2 (0.5 mL) was added
triethylamine (16 μL, 117 μmol) at room temperature. After stirring
for 15 min, cyanoethyl(diisopropylamino)phosphino chloride (17 μL,
78 μmol) was added dropwise. The stirring was continued for another
2 h, and the solvent was then removed under reduced pressure. Flash
chromatography (CH2Cl2/acetone = 95:5) gave the compound 8b as a
white foam (27 mg, 59%). As judged by 31P NMR, 8b was presented
as a 3:5 mixture of two stereoisomers: 1H NMR (CDCl3, 300 MHz) δ
1.06−1.08 (4H), 1.16−1.19 (m, 8H), 1.95−2.17 (m, 2H), 2.36−2.51
(m, 4H), 2.57−2.70 (m, 2H), 3.33−3.40 (m, 1H), 3.51−3.72 (m, 4H),
3.76−3.78 (m, 6H), 3.81−3.89 (m, 1H), 4.19−4.25 (m, 1H), 4.69−
4.74 (m, 1H), 6.38−6.45 (m, 1H), 6.82−6.86 (m, 4H), 7.23−7.33 (m,
7H), 7.39−7.44 (m, 4H), 7.50−7.55 (m, 1H), 7.89−7.94 (1H), 8.23
(d, J = 6.9 Hz, 1H); 13C NMR (CDCl3, 75 MHz) δ 19.4, 20.37, 20.47,
24.4, 24.5, 24.6, 29.8 (t, J = 25.2 Hz), 40.5, 43.1, 43.3, 43.4, 55.1, 58.1,
58.4, 62.9, 63.1, 73.8, 74.0, 85.5, 85.6, 85.7, 85.8, 86.9, 112.5, 112.6,
113.2, 117.3, 117.5, 127.2, 128.0, 128.1, 128.2, 129.8, 130.1, 132.6,
134.8, 135.06, 135.12, 136.9, 138.4, 144.0, 147.7, 158.8, 179.6; 19F
NMR (CDCl3, 282 MHz) δ −81.3, −115.2, −122.1, −123.1, −123.9,
−126.3; 31P NMR (CDCl3, 121 MHz) δ 149.6, 150.1; MS (ESI) m/z
1180.3 [M + H]+, HRMS (ESI) calcd for C54H55CsF13N5O8P [M +
Cs]+ 1312.263, found 1312.261.
ASSOCIATED CONTENT
* Supporting Information
■
S
The experimental protocols and NMR spectra of key
compounds. This material is available free of charge via the
Internet at http://pubs.acs.org.
AUTHOR INFORMATION
Corresponding Author
*E-mail: qszhang@unc.edu.
■
ACKNOWLEDGMENTS
We thank Drs. Rudy Juliano and Osamu Nakagawa (University
of North Carolina) for the help with oligonucleotide synthesis.
■
REFERENCES
■
(1) (a) Lande-Diner, L.; Zhang, J.; Ben-Porath, I.; Amariglio, N.;
Keshet, I.; Hecht, M.; Azuara, V.; Fisher, A. G.; Rechavi, G.; Cedar, H.
J. Biol. Chem. 2007, 282, 12194−12200. (b) Shames, D. S.; Minna, J.
D.; Gazdar, A. F. Curr. Mol. Med. 2007, 7, 85−102. (c) Sontag, L. B.;
Lorincz, M. C.; Georg Luebeck, E. J. Theor. Biol. 2006, 242, 890−899.
(2) Altun, G.; Loring, J. F.; Laurent, L. C. J. Cell. Biochem. 2010, 109,
1−6.
Compound 8a. In a similar manner to 8b, 8a (29 mg, 41%) was
(3) (a) Gehring, M.; Reik, W.; Henikoff, S. Trends Genet. 2009, 25,
82−90. (b) Ma, D. K.; Guo, J. U.; Ming, G. L.; Song, H. Cell Cycle
2009, 8, 1526−1531. (c) Zhu, J. K. Annu. Rev. Genet. 2009, 43, 143−
166.
synthesized from 7a (58 mg, 73 μmol) as colorless oil that existed as a
1
4:5 mixture of two stereoisomers: H NMR (CDCl3, 300 MHz) δ
1.06−1.08 (3H), 1.16−1.19 (m, 9H), 1.93−2.17 (m, 2H), 2.35−2.49
(m, 4H), 2.61−2.66 (m, 2H), 3.33−3.40 (m, 1H), 3.51−3.66 (m, 4H),
3.79 (m, 6H), 3.81−3.85 (m, 1H), 4.19−4.26 (m, 1H), 4.68−4.74 (m,
1H), 6.41 (dd, J =13.8, 6.0 Hz, 1H), 6.82−6.86 (m, 4H), 7.25−7.34
(m, 7H), 7.39−7.45 (m, 4H), 7.49−7.55 (m, 1H), 7.89−7.93 (1H),
8.23 (d, J = 7.8 Hz, 1H); 13C NMR (CDCl3, 75 MHz) δ 19.5, 20.1,
20.2, 20.3, 20.4, 24.38, 24.48, 24.54, 24.6, 29.5 (t, J = 21.2 Hz), 40.4,
40.5, 43.10, 43.18, 43.27, 43.34, 55.1, 58.0, 58.1, 58.2, 58.3, 62.9, 63.0,
73.4, 73.7, 74.0, 85.5, 85.6, 85.7, 85.8, 86.06, 86.11, 86.9, 112.5, 112.6,
113.2, 117.3, 117.5, 127.2, 128.0, 128.1, 129.8, 130.0, 130.1, 132.6,
(4) (a) Guo, J. U.; Su, Y.; Zhong, C.; Ming, G. L.; Song, H. Cell 2011,
145, 423−434. (b) He, Y. F.; Li, B. Z.; Li, Z.; Liu, P.; Wang, Y.; Tang,
Q.; Ding, J.; Jia, Y.; Chen, Z.; Li, L.; Sun, Y.; Li, X.; Dai, Q.; Song, C.
X.; Zhang, K.; He, C.; Xu, G. L. Science 2011, 333, 1303−1307. (c) Ito,
S.; Shen, L.; Dai, Q.; Wu, S. C.; Collins, L. B.; Swenberg, J. A.; He, C.;
Zhang, Y. Science 2011, 333, 1300−1303.
(5) (a) Cornils, B. Angew. Chem., Int. Ed. Engl. 1997, 36, 2057−2059.
(b) Studer, A.; Hadida, S.; Ferritto, R.; Kim, S. Y.; Jeger, P.; Wipf, P.;
10267
dx.doi.org/10.1021/jo2015399 | J. Org. Chem. 2011, 76, 10263−10268

The Journal of Organic Chemistry
Note
Curran, D. P. Science 1997, 275, 823−826. (c) Neil, E.; Marsh, G.
Chem. Biol. 2000, 7, R153−R157.
(6) Horvath, I. T.; Rabai, J. Science 1994, 266, 72−75.
(7) (a) Bergbreiter, D. E. Chemtracts: Inorg. Chem. 1995, 7, 14−18.
(b) DiMagno, S. G.; Dussault, P. H.; Schultz, J. A. J. Am. Chem. Soc.
1996, 118, 5312−5313. (c) Pozzi, G.; Banfi, S.; Manfredi, A.;
Montanari, F.; Quici, S. Tetrahedron 1996, 52, 11879−11888.
(8) Curran, D. P.; Hadida, S.; He, M. J. Org. Chem. 1997, 62, 6714−
6715.
(9) (a) Curran, D. P. Synlett 2001, 1488−1496. (b) Zhang, Q.; Luo,
Z.; Curran, D. P. J. Org. Chem. 2000, 65, 8866−8873. (c) Matsugi, M.;
Curran, D. P. J. Org. Chem. 2005, 70, 1636−1642.
(10) (a) Curran, D. P.; Luo, Z. J. Am. Chem. Soc. 1999, 121, 9069−
9072. (b) Dandapani, S.; Curran, D. P. Tetrahedron 2002, 58, 3855−
3864. (c) Luo, Z.; Williams, J.; Read, R. W.; Curran, D. P. J. Org. Chem.
2001, 66, 4261−4266.
(11) (a) Luo, Z.; Zhang, Q.; Oderaotoshi, Y.; Curran, D. P. Science
2001, 291, 1766−1769. (b) Zhang, W.; Luo, Z.; Chen, C. H.; Curran,
D. P. J. Am. Chem. Soc. 2002, 124, 10443−10450. (c) Zhang, Q.; Lu,
H.; Richard, C.; Curran, D. P. J. Am. Chem. Soc. 2004, 126, 36−37.
(12) Brittain, S. M.; Ficarro, S. B.; Brock, A.; Peters, E. C. Nat.
Biotechnol. 2005, 23, 463−468.
(13) Pearson, W. H.; Berry, D. A.; Stoy, P.; Jung, K. Y.; Sercel, A. D.
J. Org. Chem. 2005, 70, 7114−7122.
(14) El Safadi, Y.; Paillart, J. C.; Laumond, G.; Aubertin, A. M.;
Burger, A.; Marquet, R.; Vivet-Boudou, V. J. Med. Chem. 2010, 53,
1534−1545.
(15) Negishi, E.-i.; King, A. O.; Okukado, N. J. Org. Chem. 1977, 42,
1821−1823.
(16) Kwiatkowski, M.; Samiotaki, M.; Lamminmaki, U.; Mukkala, V.
M.; Landegren, U. Nucleic Acids Res. 1994, 22, 2604−2611.
(17) (a) Seela, F.; Ramzaeva, N.; Leonard, P.; Chen, Y.; Debelak, H.;
Feiling, E.; Kroschel, R.; Zulauf, M.; Wenzel, T.; Frohlich, T.;
Kostrzewa, M. Nucleosides, Nucleotides Nucleic Acids 2001, 20, 1421−
1424. (b) Pike, A. R.; Ryder, L. C.; Horrocks, B. R.; Clegg, W.;
Connolly, B. A.; Houlton, A. Chemistry 2004, 11, 344−353. (c) Seela,
F.; Feiling, E.; Gross, J.; Hillenkamp, F.; Ramzaeva, N.; Rosemeyer, H.;
Zulauf, M. J. Biotechnol. 2001, 86, 269−279.
(18) Zaehres, H.; Lensch, M. W.; Daheron, L.; Stewart, S. A.;
Itskovitz-Eldor, J.; Daley, G. Q. Stem Cells 2005, 23, 299−305.
10268
dx.doi.org/10.1021/jo2015399 | J. Org. Chem. 2011, 76, 10263−10268