Total synthesis of the hypermodified RNA bases wybutosine and hydroxywybutosine and their quantification together with other modified RNA bases in plant materials

Article abstract of DOI:10.1002/chem.201204209

We report an efficient synthesis of the hypermodified natural tRNA modifications wybutosine (yW) and hydroxywybutosine (OHyW). We also describe the preparation of isotopically labeled analogues for precise quantification of yW and OHyW in different tissue

Full text of DOI:10.1002/chem.201204209

DOI: 10.1002/chem.201204209
Total Synthesis of the Hypermodified RNA Bases Wybutosine and
Hydroxywybutosine and Their Quantification Together with Other Modified
RNA Bases in Plant Materials
Antje Hienzsch, Christian Deiml, Veronika Reiter, and Thomas Carell*[a]
Abstract: We report an efficient syn-
thesis of the hypermodified natural
tRNA modifications wybutosine (yW)
and hydroxywybutosine (OHyW). We
also describe the preparation of isotop-
ically labeled analogues for precise
quantification of yW and OHyW in dif-
ferent tissues including plant materials.
The synthesis involved the formation
of the unusual tricyclic ring structure of
the bases by using a catalytic, intramo-
lecular hydroamination reaction. The
basis for the synthesis is also a stereo-
selective coupling reaction that allows
the introduction of the fully substituted
side chains to the tricyclic core struc-
ture. The isotopologues of yW and
OHyW, together with other isotopically
labeled tRNA modifications, were ulti-
mately used in LC-MS quantification
experiments to investigate the role of
the modified bases in the translational
process. Quantification was performed
in the plant species Arabidopsis thali-
ana.
Keywords: isotopic labeling · liquid
chromatography · natural products ·
nucleosides · RNA
Introduction
OHyW 2 is present in various eukaryotes, yW 1 has so far
only been isolated from yeasts. Research in recent years has
helped to gain initial insight into the role of yW and OHyW
during translation. yW contributes to the stabilization of the
tRNA by means of its large and hydrophilic side chain,
which reduces the conformational flexibility of the antico-
don loop.[2] Additionally, the hydrophobic tricyclic core
structure enhances the codon–anticodon interaction due to
increased base stacking.[3] When the yW base is excised
from the tRNAPhe, the conformation of the anticodon is
strongly altered.[4] This structural change leads to a destabili-
zation of the codon–anticodon interaction at the ribosomal
A and P site.[3,5] Although the stabilizing effect of the yW
modification has been clearly demonstrated, no significant
influence on the aminoacylation process and hence the pro-
tein synthesis has been observed so far.[6] Recently it was
shown that the absence of yW promotes À1 frameshift
events.[7] For the tRNAPhe, it was clearly demonstrated that
the frameshift rates are increased in vivo[7,8] and in vitro[9]
when the yW base is replaced by m1G. A major question as-
sociated with yW and OHyW is their distribution in the bio-
sphere, and particularly in plants in which they were recent-
ly detected. To investigate this question, we synthesized yW
and OHyW also in an isotope-labeled form to enable a pre-
cise quantification of the compounds.
The hypermodified nucleosides wybutosine (yW, 1) and hy-
droxywybutosine (OHyW, 2; Figure 1) belong to the large
family of modified transfer RNA (tRNA) nucleosides.[1]
They occur exclusively in the anticodon stem loop of eucary-
otic phenylalanine tRNA (tRNAPhe) at position 37, at the 3’-
position adjacent to the Phe anticodon (34GAA36). Whereas
Figure 1. Depiction of the hypermodified tRNA nucleosides wybutosine
1 (yW) and hydroxywybutosine 2 (OHyW), as well as their isotopically
labeled analogues 3 and 4.
Herein we report that a novel copper(I)-mediated cross-
coupling reaction between the tricyclic core structure and
the fully functionalized side chains allows the fast and effi-
cient synthesis of the natural wybutosine RNA bases. The
chemistry also enabled the total synthesis of hydroxywybu-
tosine in addition to stable isotope-labeled compounds. By
using the synthetic material, we ultimately quantified the
levels of these modified bases in several tissues.
[a] Dr. A. Hienzsch, C. Deiml, V. Reiter, Prof. Dr. T. Carell
Center for Integrated Protein Science (CIPS) at the
Department of Chemistry
Ludwig-Maximilians-Universitꢀt Mꢁnchen
Butenandtstrasse 5–13, 81377 Mꢁnchen (Germany)
Fax : (+49)2180-77756
E-mail: thomas.carell@cup.uni-muenchen.de
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/chem.201204209.
4244
ꢂ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Eur. J. 2013, 19, 4244 – 4248

FULL PAPER
Results and Discussion
The synthesis starts with acetyl-protected guanosine 5. We
introduced a propargyl group at the 1-position for a later
catalytic hydroamination reaction to give derivative
6
(Scheme 1). To achieve the cycloreaction, we tested various
Scheme 2. a) HBr in AcOH, 1008C, pressure, 5h, 96%; b) AcCl, MeOH,
RT, 24 h, c) dimethyl dicarbonate, NaHCO3, dioxane/H2O, 16 h, 08C;
À
d) NaI, acetone, reflux, 2 h, 83% over 3 steps; e) PhSe SePh, NaBH4,
MeOH, RT, 20 min, 96%; f) 30% H2O2, THF, reflux, 1 h, 85%; g) meta-
chloroperbenzoic acid (mCPBA), CH2Cl2, (1:3), 78%; h) MgI2, toluene,
À608C, 88%; i) trimethylsilyl chloride (TMSCl), pyridine, 408C, 90%.
nated precursor 8. Finally, simple deprotection of 12 with
NaOMe gave pure wybutosine 1 (76%).[12]
We next extended the study to the synthesis of OHyW 2.
For its synthesis we first needed the OH-containing side
chain. Thus, bromide 13 was converted to an intermediate
phenylselenide,[13] which after treatment with H2O2 gave the
vinylglycine derivative 14 (Scheme 2).[15] Epoxidation with
mCPBA delivered the corresponding epoxide diastereomers
(syn/anti 3:1), which were separated by flash chromatogra-
phy. Mild opening of the epoxide ring 15 with MgI2 at low
temperatures gave the free alcohol.[16] This compound was
subsequently converted to the trimethylsilylether 10, which
was used for the coupling reaction. The copper-mediated
cross-coupling proceeded smoothly under the conditions de-
scribed above. The protected hydroxywybutosine derivative
16 was isolated in 63% yield (Scheme 3). Final deprotection
Scheme 1. a) NaH, DMF, propargyl bromide, 88%; b) Zn, 1,2-dibromo-
ethane, CH3CN, 50%, 45% recovery of 6; c) CH2I2, Zn(Et)2, 1,2-dime-
thoxyethane (DME), Et2O, toluene, CH2Cl2, 75%; d) I2, CF3CO2Ag,
CH2Cl2, 93%.
Lewis acids. The best results were finally obtained by treat-
ing propargyl guanosine 6 with in situ prepared ZnBr2 in
acetonitrile heated to reflux, which led to the tricyclic struc-
ture in about 50% yield. Although the yield is moderate, we
were able to recover 45% of the starting material, so that
the yield on the basis of recovered starting material is 91%.
Subsequently, methylation with Zn(Et)2 and iodomethane[10]
gave the strongly fluorescent wyosine derivative 7,[11] fol-
lowed by the introduction of iodine to generate key inter-
mediate 8.[12]
We next prepared the side chains 9 and 10 as the required
second building blocks for the intended cross-coupling reac-
tion. As starting material for those precursors, (S)-(À)-a-
amino-g-butyrolactone hydrobromide (11) was chosen.
Compound 11 was first transformed to the corresponding
(S)-2-carboxymethylamino-4-iodobutyric acid methyl ester
(9) by using an established four-step procedure with an over-
all yield of 83% (Scheme 2).[13]
We next investigated the cross-coupling reaction between
8 and 9. The best results were achieved using a copper(I)-
mediated coupling approach. For the reaction we first per-
formed a smooth magnesium–iodine exchange of 8 that pro-
ceeded well within 30 min at À788C. The addition of
CuCN·2LiCl, followed by 9 and reaction at À258C for 16 h,
gave the product 12 in fair yields (50%).[14] The product was
obtained together with reduced wyosine 7, which was sepa-
rated from the product by column chromatography. The ap-
pearance of 7 as the main side product shows that the yield
of the coupling reaction is limited by reduction of the iodi-
Scheme 3. a) sec-BuMgCl, À788C, 30 min; b) CuCN·2LiCl, À608C,
15 min; c) 9 or 10 (0.5 equiv), À60 to À258C, 16 h; R = H, 50%, R =
OH, 63%; d) NaOMe, MeOH, NaH2PO4, H2O, 0 8C, 75–76%.
Chem. Eur. J. 2013, 19, 4244 – 4248
ꢂ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemeurj.org
4245

T. Carell et al.
from cell culture or from specific tissues. Those samples
were subjected to LC-MS analysis, and the ratio of the mass
signals from the natural nucleoside and its isotopologue was
determined. On the basis of calibration curves measured
with the isotopologues, the precise amount of a certain mod-
ification in the original sample was determined, even if the
modified bases were present only in minimal amounts.[18]
The unlabeled compounds were used to verify the structural
assignment.
Together with the quantification of yW and OHyW, 16
further modified RNA bases were quantified analogously in
Arabidopsis thaliana. The compounds and a list of the data
are reported in Table 1. We also compared RNA isolated
from a plant cell culture of A. thaliana with RNA material
isolated from whole plants grown on synthetic media for
15 days. The intention was to study how much cell-culture
conditions influence the diversity of the translational appa-
ratus. The investigated modifications were located at various
positions in the tRNA. Some were found in the anticodon
loop, others in scaffold regions of tRNA. To give a compre-
hensive picture, the data comprise the ubiquitous modifica-
Scheme 4. a) NaH, DMF, 4 ꢃ mol. sieves, [D5]bromoacetone, 6 h, 67%,
RT; b) CH2I2, ZnEt2, DME, Et2O, CH2Cl2, 108C, 5 min, 75%; c) I2,
CF3CO2Ag, CH2Cl2, RT, 10 min, 91%; d) iPrMgCl/sec-BuMgCl, CuCN·2
LiCl, 9 or 10, THF, À78 to À258C, 50 or 63%; e) NaOMe, MeOH,
NaH2PO4, H2O, 0 8C, 72–76%.
Table 1. Quantitative data for the investigated tRNA modifications in A. thaliana grown in cell culture and as
A. thaliana plants. All tRNA nucleoside values are given per 1000 tRNA molecules [%]. These data reveal a
higher modification level of cells grown in cell culture than in intact plants. The upper table shows modifica-
tions found in the scaffold region or in the anticodon of tRNAs and the lower table includes the modifications
found in position 37, at the 3’-position adjacent to the anticodon of the tRNAs.
with NaOMe completed the
synthesis of the desired OHyW
2 (75%).
To detect and quantify both
natural products in biological
tRNA samples, an isotope label
needs to be introduced into the
synthetic product (Scheme 4).
We considered the “third ring”
to be a suitable and economic
site to introduce deuterium
labels. Due to availability of la-
Modification
Gr
m2G
ac4C
Am
m7G
m2 G
Cm
m5C
Gm+m1G
2
A. th. cell culture
A. th. plants
15.7
10.8
430.5
316.0
99.6
78.3
34.7
29.2
388.9
368.0
601.6
607.2
257.7
180.5
1227.6
984.1
855.7
762.6
Modification
OHyW
yW
m6A
m6t6A
io6A
t6A
m2A
ms2io6A
8.7
38.7
i6A
A. th. cell culture
A. th. plants
2.6
1.1
26.0
17.1
20.6
14.5
14.6
12.9
53.8
59.6
112.9
150.5
41.7
60.6
5.4
27.6
beled acetone, we decided to introduce the third ring with
[D5]bromoacetone.[17] Consequently, bromine was treated
with [D6]acetone in the presence of [D1]acetic acid to yield,
after careful distillation, pure [D5]bromoacetone. Reaction
tions m5C, m7G, m1G, m2 G, and m2G, as well as the rarely
2
present modified nucleosides i6A and OHyW, which could
indeed be detected in small amounts.
In 2010, the quantification of A. thaliana tRNA modifica-
tions on the basis of UV and high-performance liquid chro-
matograms was reported; in this study, none of the wybuto-
sine derivatives or of other minor modifications were detect-
ed.[19] In contrast, by using our isotope method and synthetic
material, both yW and OHyW were clearly detected togeth-
er with the other rare A. thaliana modifications i6A, io6A,
and Gr. In addition, our analysis shows that no queuosine
derivatives were present, even though their existence in
plant RNA was predicted. Just recently they were shown to
occur in maize and pea plants.[20] The detection of yW and
HOyW in plants shows that the bases are present side-by-
side and more widely distributed than previously thought.
The quantification data are presented in Table 1. The
values represent the measured number of each modification
per 1000 tRNA molecules [%]. Instead of giving the exact
concentration, this representation allows one to quickly rec-
ognize to what extent the analyzed tRNA set is modified.
of
[D5]bromoacetone
with
5
gave
[D3]-
and
[D4]desmethylwyosine 17 in a ratio of 4:1.
We explain the unexpected product mixture with a keto–
enol tautomerization of the bromoacetone under the reac-
tion conditions. In this way, the deuterium atoms at the
methylene position of bromoacetone are exchanged with hy-
drogen atoms, which are likely donated by the guanosine
compound 5. Subsequently the desmethylwyosine 17 was
treated in analogy to the natural modification to 19, to give
after copper-mediated coupling with 9 and 10, followed by
deprotection, the desired [D3]-labeled wybutosine 3 and hy-
droxywybutosine 4, respectively.
The prepared isotope- and unlabeled wybutosines (1 and
3) and hydroxywybutosines (2 and 4) were next used for iso-
tope dilution-based LC-MS quantification. Briefly, we added
defined amounts of the isotopically labeled nucleosides to
enzymatically hydrolyzed total tRNA samples, extracted
4246
www.chemeurj.org
ꢂ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Eur. J. 2013, 19, 4244 – 4248

Wybutosine and Hydroxywybutosine
FULL PAPER
From the data, furthermore, it is evident that the amount
of modifications, measured in RNA isolated from cell cul-
ture and whole plants, are quite different. Twelve of 18 ob-
served modifications are more abundant in cells kept in cul-
adenine scaffold that bears an isoprenoid side chain at the
N6 nitrogen. Additionally, they are modified with different
C5 and C6 carbohydrates at various positions on the purine
scaffold or they carry a methylthio group at the 2-position
of the purine rings.[26] Therefore their structure is essentially
the same as that of the tRNA modifications i6A, io6A, and
ms2io6A. Due to the common appearance of the cytokinin
nucleobases and their corresponding nucleosides to the
tRNA nucleosides, it is suggested that both substrate classes
share the same biosynthetic enzymes, which are indeed
known to have a broad substrate specificity.[27] As cytokinins
are very important signal transducers during development
and differentiation of plants, a higher level of the corre-
sponding biosynthetic enzymes in developing plants could
lead to an increased level of i6A modifications in their
tRNAs.
ture than in plants. One modification, m2 G, is nearly un-
2
changed, and five modifications are more abundantly pres-
ent in plants. These five modifications can all be found in or
at the 3’-position adjacent to the anticodon and are there-
fore directly involved in the recognition of the anticodon
loop by the ribosome. It is also interesting that three of the
five modifications belong to the family of the i6A modifica-
tions; the others are m2A and t6A.
Our results for methyl RNA modifications are in good
agreement with previously published observations for tobac-
co plants, in which the methylation patterns were compared
between intact tobacco plants and the corresponding tobac-
co cell-culture samples. In this study, the authors also detect-
ed more methylated RNA nucleosides (m5C, m1G, m2G,
m2 G, m7G, m5U) in the cell-culture material than in whole
Conclusion
2
tobacco plants.[21] This difference was attributed to the un-
equal differentiation status of the plant cells in culture and
of the cells isolated from real plants. Since plant cells in cul-
ture experience the same environmental conditions and
have to be ready for development into different plant
organs, they all seem to express a similar set of proteins,
which gives a more uniform mRNA and tRNA composition
and hence also of the present modifications. In contrast,
cells in intact plants are already differentiated and therefore
their sets of proteins, mRNAs and tRNAs, are modified ac-
cording to their specific needs. This close connection of
tRNA composition and differentiation status has already
been shown for wheat and mimosa tissues.[22] Another factor
is the general difference in the culture conditions. Since cells
in cell culture can uptake nutrients from the culture
medium, they are, in contrast to real plants, not forced to
perform photosynthesis to grow.[23] Photosynthesis takes
place in chloroplasts, and these organelles have their own
DNA and ribosomes, and they produce their own set of
chloroplast tRNAs. These chloroplast tRNAs are known to
have fewer modification sites[24] and display an overall re-
duced modification content.[25] Therefore, they more resem-
ble prokaryotic tRNAs[24] and their reduced amount in cell-
culture total tRNA sets might dilute the number of modifi-
cations. The higher amount of m2A in plants can also be ex-
plained by this model. The modification m2A is prevalent in
procaryotic organisms but was also found in plants.[19] Since
photosynthesis in real plants generates higher levels of
chloroplasts and chloroplast tRNA is basically a prokaryotic
RNA, it is the higher tRNA level from the chloroplasts that
makes this modification more abundant in tRNAs isolated
from real plants.
In summary, we have developed a short and convenient syn-
thesis of the hypermodified nucleosides wybutosine and hy-
droxywybutosine. The procedures allowed us to synthesize
these molecules as isotope-labeled analogues as well, which
were needed for the quantification of tRNA modifications
of the plant species A. thaliana.
These data reveal different modification levels in cell-cul-
ture samples as opposed to whole plants. Our data show
that yW and OHyW are clearly detectable in tRNA from
plants, which indicates that they are more broadly present in
tRNAs than thought so far. The data show in addition a
larger modification content in plants than in plant cell cul-
tures. In agreement with a recent study, we believe that cul-
ture conditions, differentiation status, and the requirement
for specific needs within tissues of higher organisms have a
strong impact on the formation and distribution of tRNA
modifications within a cell.[18] In this study, the protein pro-
duction level and hence the translational efficiency was cor-
related to the tRNA modification level within different tis-
sues of mouse and pig. This provides support for the as-
sumption that the tRNA modification level adapts to envi-
ronmental conditions and to the metabolomic requirement
of the cells.
Experimental Section
(aS)-a-[(Methoxycarbonyl)amino]-4,6-dimethyl-9-oxo-3-(2’,3’,5’-tri-O-
acetyl-b-d-ribofuranosyl)-4,9-dihydro-3H-imidazoACTHNUGTRNEUNG[1,2-a]purine-7-butano-
ic acid methyl ester (12): Compound 8 (509 mg, 0.867 mmol) was put into
a dry and argon-flushed 10 mL Schlenk flask, dissolved in anhydrous
THF (2.5 mL), and cooled to À788C. Freshly titrated iPrMgCl (477 mL,
0.954 mmol, 2m in THF, 1.1 equiv) was added dropwise, and the reaction
mixture was stirred at À788C for 30 min. A solution of CuCN·2LiCl
(434 mL, 0.434 mmol, 1m in THF, 0.5 equiv) was added dropwise and the
solution was allowed to warm to À608C within 15 min, at which point 9
(131 mg, 0.433 mmol, 0.5 equiv), dissolved in anhydrous THF (1 mL),
was slowly added. The reaction mixture was slowly warmed to À258C
and stirred at this temperature for 16 h. The reaction was allowed to
The higher level of modifications of the i6A family in
whole plant samples can be attributed to a class of cytoki-
nins in plants that act as phytohormones. These cytokinins
are involved in growth and differentiation and therefore in-
fluence various processes like cell division, photosynthesis,
aging, and metabolism. One class of such cytokinins have an
Chem. Eur. J. 2013, 19, 4244 – 4248
ꢂ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
www.chemeurj.org
4247

T. Carell et al.
warm to room temperature and quenched by the addition of an aqueous
saturated NH4Cl/NH4OH (2 mL, 9:1) solution followed by CH2Cl2
(15 mL). The solution was vigorously stirred until the aqueous layer
became dark blue. The layers were separated, and the aqueous layer was
extracted twice with CH2Cl2. The combined organic layers were washed
with brine, dried over MgSO4, and concentrated. The crude product was
purified by flash column chromatography on silica gel (CH2Cl2/MeOH
500:10) to afford the pure product 12 (136 mg, 0.214 mmol, 50%) as a
colorless foam. Side product 7 was isolated as well as a white foam
(260 mg, 0.564 mmol). M.p. 968C; 1H NMR (600 MHz, CDCl3, 258C,
TMS): d=2.00–2.07 (m, 1H; CbH), 2.11–2.18 (m, 1H; CbH), 2.11 (s, 3H;
COCH3), 2.13 (s, 3H; C6CH3), 2.17 (s, 3H; COCH3), 2.22 (s, 3H;
COCH3), 2.95–3.00 (m, 1H; CgH), 3.27–3.32 (m, 1H; CgH), 3.69 (s, 3H;
OCH3), 3.70 (s, 3H; OCH3), 4.13 (s, 3H; N4CH3), 4.29–4.34 (m, 3H; CaH
and C5’H), 4.49 (dd, 3J=2.9, 3.2 Hz, 1H; C4’H), 5.48 (dd, 3J=3.6, 5.4 Hz,
1H; C3’H), 5.83 (dd, 3J=5.6, 5.8 Hz, 2H; C2’H, NH), 6.19 (d, 3J=6.0 Hz,
1H; C1’H), 7.73 ppm (s, 1H; C2H); 13C NMR (150 MHz, CDCl3, 258C,
TMS): d=12.4 (C6CH3), 20.3 (COCH3), 20.5 (COCH3), 20.7 (COCH3),
20.9 (Cg), 32.8 (Cb), 33.7 (N4CH3), 52.2 (OCH3), 52.3 (OCH3), 53.6 (Ca),
62.7 (C5’), 70.6 (C3’), 72.6 (C2’), 80.9 (C4’), 85.8 (C1’), 117.3 (CC9), 121.8
(C7), 133.4 (C6), 134.5 (C2), 139.4 (N3CN4), 142.5 (N4CN5), 154.2 (C9),
157.0 (COO), 169.1 (CAcOO), 169.5 (CAcOO), 170.1 (CAcOO), 172.9 ppm
(COO); IR (ATR, neat): n˜max =3602 (s), 3363 (s), 3132 (s), 2954 (s), 2848
(s), 2360 (s), 1745 (l), 1698 (l), 1587 (m), 1568 (m), 1532 (m), 1447 (s),
1366 (m), 1217 (l), 1096 (m), 1051 (m), 903 (s), 821 (s), 784 (s), 764 (s),
705 (s), 627 cmÀ1 (s); HRMS (ESI): m/z: calcd for C27H35N6O12 [M+H+]+:
635.2313; found: 635.2322.
Sci. USA 1994, 91, 2125–2129; b) D. R. Kearns, K. L. Wong, Y. P.
Wong, Proc. Natl. Acad. Sci. USA 1973, 70, 3843–3846.
[5] V. Katunin, N. Soboleva, V. Mahkno, E. Sedelnikova, S. Zhenodaro-
va, S. Kirillov, Biochimie 1994, 76, 51–57.
[6] a) R. Thiebe, H. G. Zachau, Eur. J. Biochem. 1968, 5, 546–555;
b) G. Krauss, F. Peters, G. Maass, Nucl. Acids Res. 1976, 3, 631–640;
c) W. Wintermeyer, H. G. Zachau, Eur. J. Biochem. 1979, 98, 465–
475; d) D. W. Smith, A. L. McNamara, J. F. Mushinski, D. L. Hat-
field, J. Biol. Chem. 1985, 260, 147–151.
[7] W. F. Waas, Z. Druzina, M. Hanan, P. Schimmel, J. Biol. Chem.
2007, 282, 26026–26034.
[8] B. A. Carlson, B. J. Lee, D. L. Hatfield, Biochem. Biophys. Res.
Commun. 2008, 375, 86–90.
[9] B. A. Carlson, J. F. Mushinski, D. W. Henderson, S. Y. Kwon, P. F.
Crain, B. J. Lee, D. L. Hatfield, Virology 2001, 279, 130–135.
[10] H. Bazin, X. X. Zhou, C. Glemarec, J. Chattopadhyaya, Tetrahedron
Lett. 1987, 28, 3275–3278.
[11] S.-i. Nakatsuka, T. Ohgi, T. Goto, Tetrahedron Lett. 1978, 19, 2579–
2582.
[12] T. Itaya, M. Morisue, M. Shimomichi, M. Ozasa, S. Shimizu, S. Na-
kagawa, J. Chem. Soc. Perkin Trans. 1 1994, 2759–2765.
[13] D. M. Bartley, J. K. Coward, J. Org. Chem. 2005, 70, 6757–6774.
[14] W. Dohle, D. M. Lindsay, P. Knochel, Org. Lett. 2001, 3, 2871–2873.
[15] Y. Feng, J. K. Coward, J. Med. Chem. 2005, 48, 655.
[16] C. Bonini, G. Righi, Tetrahedron 1992, 48, 1531–1538.
[17] S. Porcher, Helv. Chim. Acta 2008, 91, 1219–1235.
[18] C. Brandmayr, M. Wagner, T. Brꢁckl, D. Globisch, D. Pearson,
A. C. Kneuttinger, V. Reiter, A. Hienzsch, S. Koch, I. Thoma, P.
Thumbs, S. Michalakis, M. Mꢁller, M. Biel, T. Carell, Angew. Chem.
Int. Ed. 2012, 51, 11162–11165.
[19] P. Chen, G. Jager, B. Zheng, BMC Plant Biol. 2010, 10, 201.
[20] L. Y. Atanasova, Compt. Rend. Acad. bulg. Sci. 2011, 64, 67–74.
[21] L. Jones, T. Scott, Plant Physiol. 1981, 67, 535–538.
[22] a) B. S. Vold, P. S. Sypherd, Proc. Natl. Acad. Sci. USA 1968, 59,
453–458; b) G. N. Brown, Plant Physiol. 1969, 44, 272–276.
[23] K. E. Koch, Annu. Rev. Plant Physiol. Plant Mol. Biol. 1996, 47,
509–540.
Acknowledgements
We thank SFB 749 and the DFG Normalverfahren CA275/8-4 for finan-
cial support.
[1] T. Carell, C. Brandmayr, A. Hienzsch, M. Mꢁller, D. Pearson, V.
Reiter, I. Thoma, P. Thumbs, M. Wagner, Angew. Chem. Int. Ed.
2012, 51, 7110–7131.
[2] a) F. H. C. Crick, J. Mol. Biol. 1966, 19, 548–555; b) A. Maelicke,
F. V. D. Haar, F. Cramer, Biopolymers 1973, 12, 27–43; c) L. S. Kan,
P. O. P. Tsꢄo, F. Von der Haar, M. Sprinzl, F. Cramer, Biochemistry
1975, 14, 3278–3291.
[3] A. L. Konevega, N. G. Soboleva, V. I. Makhno, Y. P. Semenikov, W.
Wintermeyer, M. V. Rodnina, V. I. Katunin, RNA 2004, 10, 90–101.
[4] a) V. Dao, R. Guenther, A. Malkiewicz, B. Nawrot, E. Sochacka, A.
Kraszewski, J. Jankowska, K. Everett, P. F. Agris, Proc. Natl. Acad.
[24] L. Marechal-Drouard, J. H. Weil, A. Dietrich, Annu. Rev. Plant
Physiol. Plant Mol. Biol. 1993, 44, 13–32.
[25] G. R. Bjçrk, J. Kohli in Chromatography and Modification of Nu-
cleosides: Biological roles and function of modification (Eds.: C. W.
Gehrke, K. C. T. Kuo), Elsevier, Amsterdam, 1990, pp. 813–867.
[26] H. Sakakibara, Annu. Rev. Plant Biol. 2006, 57, 431–449.
[27] a) M. Allen, W. Qin, F. Moreau, B. Moffatt, Physiol. Plant. 2002,
115, 56–68; b) K. M. Schnorr, C. Gaillard, E. Biget, P. Nygaard, M.
Laloue, Plant J. 1996, 9, 891–898.
Received: November 26, 2012
Published online: February 18, 2013
4248
www.chemeurj.org
ꢂ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Chem. Eur. J. 2013, 19, 4244 – 4248