High-density DNA functionalization by a combination of Cu-catalyzed and Cu-free click chemistry

Article abstract of DOI:10.1002/chem.201000363

We report the regioselective Cu-free click modification of styrene functionalized DNA with nitrile oxides. A series of modified oligodeoxynucleotides (nine base pairs) was prepared with increasing styrene density. 1,3-Dipolar cycloaddition with nitrile ox

Full text of DOI:10.1002/chem.201000363

FULL PAPER
DOI: 10.1002/chem.201000363
High-Density DNA Functionalization by a Combination of Cu-Catalyzed and
Cu-Free Click Chemistry
Katrin Gutsmiedl, Danila Fazio, and Thomas Carell*[a]
Abstract: We report the regioselective
Cu-free click modification of styrene
functionalized DNA with nitrile oxides.
A series of modified oligodeoxynucleo-
tides (nine base pairs) was prepared
with increasing styrene density. 1,3-Di-
polar cycloaddition with nitrile oxides
allows the high density functionaliza-
tion of the styrene modified DNA di-
rectly on the DNA solid support and in
solution. This click reaction proceeds
smoothly even directly in the DNA
synthesizer and gives exclusively 3,5-
disubstituted isoxazolines. Additionally,
PCR products (300 and 900 base pairs)
were synthesized with a styrene tri-
phosphate and KOD XL polymerase.
The click reaction on the highly modi-
fied PCR fragments allows functionali-
zation of hundreds of styrene units on
these large DNA fragments simulta-
AHCTUNGTRENNnGUN eously. Even sequential Cu-free and
Cu-catalyzed click reaction of PCR
amplicons containing styrene and
alkyne carrying nucleobases was ach-
ieved. This new approach towards
high-density functionalization of DNA
is simple, modular, and efficient.
Keywords: click chemistry · DNA
labeling · nitrile oxides · nucleo-
tides · polymerase chain reaction
Introduction
ly achieved in two different ways. The molecules can be di-
rectly inserted into DNA, for example, as phosphoramidite
building blocks for solid phase oligonucleotide assembly or
as triphosphates via PCR. Alternatively, and particularly im-
portant for sensitive molecules, an anchor molecule can be
inserted into the oligonucleotide during DNA synthesis that
enables the postsynthetic attachment of the desired mole-
cule to the DNA strand.[6] The most prominent method cur-
rently available for the functionalization of DNA is the CuI-
catalyzed alkynes and azides cycloaddition[7] developed by
Meldal[8] and Sharpless.[9] Others and us reported the use of
the CuI-catalyzed click reaction for the high density func-
tionalization of oligonucleotides.[10] The procedure requires
incorporation of alkyne-carrying nucleobase either as phos-
phoramidites or triphosphates into oligonucleotides and the
postsynthetic functionalization of the corresponding alkyne-
DNA with various azides.[11]
The precise functionalization of oligonucleotides is of para-
mount importance in the research areas of DNA based
nano- and biotechnology.[1] In both fields the extraordinary
self-assembly and self-recognition capability of oligonucleo-
tides are either utilized to organize nanomaterials or to bind
to specific medicinally relevant oligonucleotide targets.[2] In
nanotechnological applications, DNA is typically used to as-
semble nanoparticles, inorganic clusters or enzymes in one,
two, and now even three dimensions.[3] DNA in biomolecu-
lar applications is mostly utilized as sequence specific probe
molecule. Functionalized with, for example, fluorescence
donors and acceptors or redox probes, the functionalized
oligonucleotides hybridize specifically with relevant DNA or
RNA molecules to enable their detection and/or isolation
for diagnostic purposes.[4] For all these applications, reporter
and anchor molecules or molecules, which carry a certain
function such as electrical conductivity or magnetism have
to be linked to the DNA or RNA molecule.[5] This is typical-
Although the CuI-catalyzed reaction of alkynes with
azides was found to be high yielding and efficient, the pres-
ence of CuI can cause problems, particularly when the syn-
thesis of DNA–protein conjugates is desired.[12] Up to now
several Cu-free click reactions were developed such as the
reaction of azides with strained cycloalkynes pioneered by
Bertozzi,[13] the reaction of tetrazines with strained al-
kenes,[14] reaction of nitrile oxides with alkynes,[15] reaction
of oxanorbornadienes with azides,[16] and a photoclick reac-
tion discovered by Lin.[17] We recently reported the use of
norbornene-functionalized oligonucleotides, in which the
[a] Dipl.-Chem. K. Gutsmiedl, Dipl.-Chem. D. Fazio, Prof. Dr. T. Carell
Department of Chemistry, Center for Integrated Protein Science
(CiPSM), Butenandtstrasse 5–13, 81377 Munich (Germany)
Fax : (+49)89-2180-77756
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.201000363.
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strained norbornene double bond was found to react quickly
with in situ prepared nitrile oxides.[18] However, this reaction
gives for each norbornene reaction site two regioisomers,
which makes the oligonucleo-
TBDMS protection of 5-iodouridine 3 (yield of 90%) fol-
lowed by Sonogashira coupling of the TBDMS-protected
nucleoside with propargylic alcohol to yield compound 4 in
tide purification complicated.
Here we report that styrene
functionalized oligonucleotides
react fast and smooth with ni-
trile oxides. This reaction fur-
nishes exclusively one regioiso-
mer, the 3,5-disubstituted isoxa-
zolines. A styrene modified tri-
phosphate was found to be
readily accepted by polymerase
so that it can be inserted into
long gene fragments by PCR.
Finally we found that it is possi-
ble to insert alkyne- and sty-
rene-functionalized
building
blocks into PCR amplicons si-
multaneously.[19] The reactions
of azides with the alkyne sites
in DNA (Cu-catalyzed) and of
the nitrile oxides with the sty-
rene units (Cu-free) were found
to be orthogonal so that
alkyne–styrene modified PCR
fragments can be high density
functionalized with two differ-
ent sets of molecules as depict-
ed in Scheme 1.
Scheme 1. Schematic depiction of the postsynthetic functionalization of DNA strands using CuI-catalyzed
alkyne–azide and Cu-free styrene–nitrile oxide click chemistry. (dATP: deoxyadenosine triphosphate; dCTP:
deoxycytidine triphosphate; dGTP: deoxyguanosine triphosphate.).
Results and Discussion
Synthesis of styrene building
blocks: For the postsynthetic
modification of DNA strands
via the nitrile oxide click reac-
tion we chose two different
strategies. For the synthesis of
short
(ODNs) we prepared the sty-
rene phosphoramidite and
oligodeoxynucleotides
1
used it as a building block for
standard DNA solid-phase syn-
thesis (Scheme 2). In order to
generate long gene fragments
with hundreds of styrene units,
we prepared the styrene-modi-
fied deoxyuridine triphosphate
2, which we incorporated using
PCR. In the PCR reaction TTP
is simply replaced by compound
2.
Scheme 2. a) Synthesis of styrene-bearing phosphoramidite 1 for DNA solid-phase synthesis; b) styrene tri-
phosphate 2 and alkyne triphosphate 7 as building blocks for PCR, and c) list of the prepared oligonucleotides
The synthesis of compound 1
(Scheme 2a)
starts
with (ODNs) containing nucleoside 5.[21]
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75% yield.[20] Subsequent Williamson ether synthesis with 4-
vinylbenzylchloride and TBAF deprotection furnished the
styrene-bearing nucleoside 5 in 52% yield over two steps.
For DNA solid-phase synthesis the phosphoramidite build-
ing block 1 was prepared. DMT protection using standard
methods was possible in 85% yield. Final phosphitylation
with 2-cyanoethyl-N,N,N’,N’-tetraisopropylphosphordiami-
dite gave compound 1 in 77% yield.[21] Incorporation of the
styrene-phosphoramidite 1 into ODNs-1–4 was performed
using ultramild DNA solid-phase synthesis (see Sche-
me 2c).[21,22] Excellent oligonucleotide syntheses were ob-
tained, even in the case of ODN-4 where six phosphorami-
dites 1 were coupled in a row. For the PCR-based approach
we converted the styrene nucleoside 5 into the correspond-
ing styrene triphosphate 2 (Scheme 2b).[21]
ciently used as a template. Full elongation by Deep vent
exoÀ was only achieved for T1 and T2.
Encouraged by these results, we started to incorporate
styrene-triphosphate 2 into longer DNA strands. For this we
utilized a real-time (RT) PCR assay with SYBR Green II as
fluorescent DNA intercalator. Cycling times, annealing tem-
peratures, chemical additives, and substrate concentrations
were carefully screened. We first investigated the ability of
the polymerases Pwo, Deep Vent exoÀ and KOD XL to am-
plify a 300 base pair (bp) fragment and a 900 bp fragment of
the Pol h gene. The 300 bp PCR amplicon features 154 thy-
midines, which need to be replaced during the PCR. The
900 bp PCR fragment contains 551 thymidines that need to
be exchanged. We discovered that this is possible with the
KOD XL polymerase, which accepts the triphosphate 2 as a
substrate without problems.[21]
Primer extension studies and RT-PCR investigations: To ex-
plore the ability of triphosphate 2 to function as a substrate
for DNA polymerases, primer extension studies were per-
formed (Figure 1).[21] The experiment, designed by Held
et al.,[23] was carried out with three different templates (T1–
T3) and various polymerases, including the family B poly-
merases Pyrococcus woesi (Pwo) and Thermococcus litoralis
(Deep vent exoÀ) as well as Thermococcus kodakaraensis
(KOD XL). For template T1 only one modified uridine
needs to be incorporated into the primer. Copying of T2
and T3 requires that the polymerase inserts two and eight
modified nucleosides. Elongation of the primer on template
T3 is particularly difficult because it requires reading
through a stretch of three consecutive modified bases. All
primer extensions were performed with a 5’-fluorescein-la-
beled primer P at 728C for 10 min or 1 h (see Figure 1).[21]
The primer extension studies with styrene triphosphate 2
revealed efficient incorporation and full elongation of the
primer on all three templates T1–T3 within 10 min reaction
time with Pwo and KOD XL polymerase. Even T3 was effi-
Figure 2a shows the 300 bp PCR products PCR300·N
(dATP, dCTP, TTP, dGTP) and PCR300·2 (dATP, dCTP,
dGTP, 2). Figure 2c shows the results obtained for the
900 bp fragments PCR900·N and PCR900·2. Both fragments
PCR300·2 and PCR900·2 exhibit a reduced mobility on the
agarose gels. In Figure 2b and d the corresponding fluores-
cence curves of the RT-PCR studies are presented. These
are similar to typical PCR profiles. The fluorescence signals
of the PCR reaction producing PCR300·2 and PCR900·2 in-
crease slower in comparison to the unmodified PCR prod-
ucts. This is likely caused by slower incorporation of the tri-
phosphate 2 compared to the natural triphosphate TTP.
Figure 2. Incorporation of dU*TP 2 into a 300 bp PCR fragment using
the KOD XL polymerase. a) Agarose gel of 300 bp fragments; lane 1:
DNA ladder; lane 2: dATP, dCTP, dGTP, TTP; lane 3: dATP, dCTP,
dGTP, 2; b) corresponding RT-PCR profiles of the 300 bp amplicons with
KOD XL (dashed line: natural triphosphates; solid line: dATP, dCTP,
dGTP, 2); c) agarose gel of 900 bp fragments; lane 1: DNA ladder; lane
2: dATP, dCTP, dGTP, TTP; lane 3: dATP, dCTP, dGTP, 2; d) additional
RT-PCR profiles of the 900 bp PCR fragments (dashed line: natural tri-
phosphates; solid line: dATP, dCTP, dGTP, 2).[21]
Figure 1. Primer extension experiments with dATP, dCTP, dGTP and
dU*TP 2 and the indicated DNA polymerases. Reaction times were
either 10 min or 1 h. T1–T3 are the DNA templates and P is the 5’-fluo-
rescein-labeled primer (Sequence: 5’-Fluo-GGCTTCTTCGAACCGGG-
TAC-3’).[21]
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T. Carell et al.
Cycloaddition of nitrile oxides to short styrene-modified
ODNs: For the cycloaddition reaction of the styrene-modi-
fied DNA strands with the nitrile oxides derived from 8–13
we either mixed N-chlorosuccinimide (NCS) and the aldox-
ime (8, 10, 12 or 13) in the presence of a weak base to pro-
duce the corresponding hydroximoyl chloride in situ (Huis-
genꢁs in situ method)[24] or we directly used the hydroximoyl
chloride (9 or 11) without any base present during the reac-
tion. The concentration of the nitrile oxide was found to be
even under these neutral conditions high enough to allow ef-
ficient reaction (see Scheme 3b).[21] The nitrile oxide solu-
tion was added directly to the solid supported ODNs-1–4
(Scheme 2c) or to an unbuffered solution containing the pu-
rified, deprotected ODNs-1–4.[21]
ing DNA material was in all cases completely transformed.
The corresponding MALDI-TOF spectra are in full agree-
ment with the expected molecular weights for the DNA
products. Significantly, we detected no by-products, which
showed us that the conversion proceeded cleanly in all cases
on all ODNs under the various conditions.
Scheme 3. a) Nitrile oxide precursors 8–13 as well as azide 14, and b) ni-
trile oxide preparation via hydroximoyl chlorides.[21]
Figure 3. Raw MALDI-TOF spectra and HPLC traces of a) the unpuri-
fied click reaction performed with ODN-1 (MW 2868) containing one sty-
rene moiety and nitrile oxide precursor 8 (ODN-1a, MW 2987) on solid
support; b) the raw click product ODN-1c (MW 3134) produced with
solid supported ODN-1 (MW 2868) and dabcyl nitrile oxide precursor 12;
c) the crude click product obtained by click reaction of ODN-2 (MW
3021) comprising of two consecutive styrene moieties and nitrile oxide
precursor 9 (ODN-2a, MW 3259) directly in the DNA synthesizer.[21] (*=
by-products of the DNA synthesis.)
We initially performed the click reaction on solid support-
ed ODN-1 comprising one styrene functionality. This mate-
rial was reacted with the nitrile oxide precursors 9 and 11 in
pure water. Reaction with 12 and 13 required first addition
of NCS to the precursors. This mixture was added to the
supported DNA with some base (NaHCO3). 20-fold excess
of the nitrile oxides was allowed to react with the DNA by
shaking the reaction mixture for 1 h at room temperature.[21]
ODN-2 containing two styrene moieties was reacted with ni-
trile oxide precursor 9 directly in the DNA synthesizer. For
reaction in the synthesizer we simply pumped the nitrile
oxide solution through the DNA cartridge containing solid-
supported ODN-2 and washed after 20 min thoroughly with
acetonitrile.[21] The CPG material was washed afterwards
with DMF as well as with CH2Cl2 and finally dried. The la-
beled DNA was cleaved from the solid support with potassi-
um carbonate (50 mm in MeOH) over 3 h at room tempera-
ture and purified by reversed-phase HPLC.[21]
We also performed the nitrile oxide cycloaddition with
the styrene modified short ODNs in solution after DNA pu-
rification. For these experiments we exclusively used hy-
droximoyl chlorides as precursors for the nitrile oxide solu-
tion without any base and utilized pure water as the solvent
(see Figures S2–S5).
The same clean conversions were also observed for ODN-
4 comprising six consecutive styrene units. Reaction of
ODN-4 with nitrile oxide precursor 9 furnished one clean
reaction product which could undoubtedly be assigned to
the six-fold reacted ODN-4 (see Figure S4).[21]
Figure 3 shows representatively the MALDI-TOF and
HPLC data of the unpurified click product ODN-1a (ODN-
1 + 8, Figure 3a), ODN-1c (ODN-1 + 12, Figure 3b), and
ODN-2a (ODN-2 + 9, Figure 3c). All nitrile oxides depict-
ed in Scheme 3 were found to react quantitatively. The start-
Analysis of click efficiency: Investigation of the click trans-
formation was performed by enzymatic digestions of the var-
ious DNA reaction products.[21] Enzymatic digests were ob-
tained by using a mixture of DNA degrading enzymes (Cro-
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Chem. Eur. J. 2010, 16, 6877 – 6883

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talus adamanteus snake venom phosphodiesterase I and al-
kaline phosphatase; see Supporting Information). The ob-
tained nucleoside mixtures were subsequently analyzed by
HPLC-MS.[21] The results are depicted in Figure 4. Enzymat-
ic digests of ODN-1 (Figure 4a) before the click reaction re-
veals the canonical bases and the styrene nucleoside 5 (re-
tention time=58 min). We found that the digest around the
modified nucleoside is not complete. In most cases we ob-
tained the modified nucleoside, for example 5, and the dinu-
cleotide, for example 5+T. This dinucleotide furnishes the
additional peak with a retention time of 48 min. HPLC-MS
confirmed that this peak is caused by dinucleotide 5+T (see
also Tables S6 and S7). The enzyme mixtures seem to have
problems cleaving the linking phosphodiester bond.[21]
DNA base 15 (retention time=63 min), pyrene-derivative
16 (retention time=67 min), or dabcyl-functionalized nu-
cleoside 17 (retention time=72 min). The HPLC trace at
the diagnostic wavelength of l=310 nm for C5-modified
pyrimidines proved for all digests full conversion as no sty-
rene nucleoside 5 was detected anymore. Additional support
for the completeness of the click transformation was gained
from HPLC chromatograms with a detection wavelength of
l=350 nm for the pyrene system (Figure 4c) and l=475 nm
for the dabcyl-dye (Figure 4d). In both cases only one prod-
uct peak appeared. Most important, independent of the de-
tection wavelength no side reactions of the nitrile oxides
with, for example, the canonical bases were observed.
Further HPLC analysis of the enzymatic digestions of the
reaction products ODN-1a (Figure 4b), ODN-1b (Fig-
ure 4c), and ODN-1c (Figure 4d) showed beside the canoni-
cal DNA bases only the corresponding phenyl-modified
Click modifications of PCR products: We next investigated
the click reaction on styrene modified PCR300·2, obtained
with triphosphate 2. This PCR fragment was reacted with
the nitrile oxide precursor 9. The results of the high density
functionalization experiments were again analyzed by enzy-
matic digestion (different protocol, see Supporting Informa-
tion) after purification of the product DNA strands via
simple ethanol precipitation (see Figure 5).[21]
The HPLC-MS analysis of the digest of the PCR300·2
starting material (Figure 5a) showed the four canonical
DNA bases (T arising from the unmodified primer strands
needed for PCR) and the styrene nucleoside 5 (retention
time=58 min). Analysis of the amplicon PCR300·2a (Fig-
ure 5b) after click reaction with benzonitrile oxide, ethanol
precipitation and enzymatic digestion showed the four natu-
ral DNA bases and the corresponding 5-phenyl-2-isoxazo-
line derivative 15 (retention time=63 min). Surprisingly, no
styrene nucleoside 5 (retention time=58 min) was detected
even though every PCR300·2 molecule contains 154 styrene
modifications. These were obviously all converted into the
corresponding cycloaddition products, which proves the
amazing efficiency of this mild click reaction (see also Fig-
ure S6 and Tables S6–S7).
We next investigated the possibility to combine the nitrile
oxide based modification of styrene containing oligonucleo-
tides with the CuI-catalyzed alkyne–azide click reaction. To
this end we synthesized a 300 bp PCR fragment by using a
mixture of the styrene triphosphate 2 and the alkyne tri-
phosphate 7 (2/7 1:1).[21] In this experiments the 154 thymi-
dines are exchanged by alkyne and styrene carrying thymi-
dine derivatives in a statistical fashion. Figure 5c shows the
enzymatic digestion data of PCR300·2/7. Indeed, this ampli-
con contains next to the canonical bases the styrene building
block 5 (retention time=58 min) and the alkyne nucleoside
6 (retention time=26 min), which proves the successful in-
corporation of both modified triphosphates.[21]
To introduce two different labels we first performed the
Cu-catalyzed click reaction with the sugar azide 14. After
click reaction we purified the sugar-modified PCR product
via ethanol precipitation. We subsequently performed the
nitrile oxide click reaction using the nitrile oxide precursor
9. The final amplicon reaction product was isolated by iso-
propanol precipitation. After intensive washing PCR300·2a/
Figure 4. HPLC analysis of the enzymatic digestions of the nitrile oxide
modified oligonucleotide ODN-1. a) ODN-1 prior to click reaction.
b) ODN-1 reacted with nitrile oxide precursor 8 (ODN-1a) containing
15, c) ODN-1 after click reaction with nitrile oxide precursor 10 (ODN-
1b) containing 16, and d) ODN-1 reacted with nitrile oxide precursor 12
(ODN-1c) containing 17. The additional traces show the HPLC data at
the diagnostic wavelengths l=310, 350, or 475 nm. (*=inosine formed
due to deamination of dA during the assay.)
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T. Carell et al.
potential to change the way of how modified oligonucleo-
tides are prepared today. Further LC-ESI-MS measurements
listed in the Supporting Information support this vision (Fig-
ure S8, Table S8).[21]
Conclusion
We have shown here that the 1,3-dipolar cycloaddition of ni-
trile oxides with styrene modified oligodeoxynucleotides
(9 bp) and long PCR amplicons (300 bp) is an extremely
facile and efficient approach for the high density functionali-
zation of DNA. This mild click reaction on styrene-bearing
DNA gives exclusively the corresponding 3,5-disubstituted
isoxazoline cycloproducts. The reaction is easy to perform
on solid support, directly in the DNA synthesizer or in solu-
tion. The ready availability of all starting materials, short re-
action times and the rather simple procedures involved pro-
vide the DNA material in excellent quantity and quality.
The chemistry is orthogonal to the CuI-catalyzed reaction of
alkyne-modified DNA with azide-carrying molecules allow-
ing the sequential labeling of alkyne- and styrene-modified
PCR amplicons.
Acknowledgements
Figure 5. HPLC data of enzymatic digests of the PCR-product PCR300·2
a) before and b) after the click reaction with 9 to the corresponding prod-
uct PCR300·2a; c) HPLC traces of the enzymatic digestions of the PCR-
fragment PCR300·2/7 c) before and d) after the CuI-catalyzed click-reac-
tion with azide 14 and the Cu-free click reaction with benzonitrile oxide
precursor 9 to the corresponding product PCR300·2a/7e, and e) depiction
of the detected nucleosides. The nucleobase T comes from the unmodi-
fied primers that were used for the PCR experiment.
We thank the Excellence Clusters CiPSM and Nano-Initiative Munich
(NIM) as well as SFB 749 for generous financial support. Further sup-
port from the Fonds of the German Chemical Industry is gratefully ac-
knowledged.
[1] a) N. C. Seeman, Nature 2003, 421, 427–431; b) N. C. Seeman, Mol.
Biotechnol. 2007, 37, 246–257; c) U. Feldkamp, C. M. Niemeyer,
Angew. Chem. 2006, 118, 1888–1910; Angew. Chem. Int. Ed. 2006,
45, 1856–1876; d) M. Endo, H. Sugiyama, ChemBioChem 2009, 10,
2420–2443.
7e was subjected to analysis by enzymatic digest. Figure 5d
shows the results obtained for the double labeled
PCR300·2a/7e fragment: Next to the canonical DNA bases
only the double peaks of the sugar click product 18 (reten-
tion times=27.1 and 27.7 min) and of the 5-phenyl-2-isoxa-
zoline nucleobase 15 (retention time=63 min) were detect-
ed. The clean reaction is nicely proven at the detection
wavelength of l=310 nm, which shows only three main
products. The sugar nucleoside gives two signals because of
rapid anomerization of the sugar moiety. The obtained
result is in two respects remarkable. First, because the two
click reactions are truly orthogonal. The azide reaction pro-
ceeds smoothly at the alkynes, leaving the styrene units
intact. More important is, however, the observation that no
unreacted alkyne or styrene building block was detected,
showing that both reactions are extremely high yielding
even on complex oligonucleotide substrates. One should
keep in mind that a 300 bp DNA fragment has a molecular
weight of approximately 90 kDa, which is in the range of a
normal size protein. Chemistry that enables the quantitative
conversion of 154 reaction sites on such complex chemical
entities, which are surely folded in aqueous solution, has the
[2] D. A. Rusling, G. Peng, N. Srinivasan, K. R. Fox, T. Brown, Nucleic
Acids Res. 2008, 37, 1288–1296.
[3] a) M. K. Beissenhirtz, I. Willner, Org. Biomol. Chem. 2006, 4, 3392–
3401; b) I. Willner, B. Willner, E. Katz, Bioelectrochemistry 2007, 70,
2–11; c) F. A. Aldaye, A. L. Palmer, H. F. Sleiman, Science 2008,
321, 1795–1799; d) A. Heckel, M. Famulok, Biochimie 2008, 90,
1096–1107; e) C. M. Niemeyer, Angew. Chem. 2010, 122, 1220–
1238; Angew. Chem. Int. Ed. 2010, 49, 1200–1216; f) L. Xu, D. Liu,
Chem. Soc. Rev. 2010, 39, 150–155; g) L. Rçglin, O. Seitz, Org.
Biomol. Chem. 2008, 6, 3881–3887; h) J. S. Hartig, S. H. Najafi-
Shoushtari, I. Grune, A. Yan, A. D. Ellington, M. Famulok, Nat.
Biotechnol. 2002, 20, 717–722; i) M. Fischler, A. Sologubenko, J.
Mayer, G. Clever, G. Burley, J. Gierlich, T. Carell, U. Simon, Chem.
Commun. 2008, 169–171; j) J. Elbaz, R. Tel-Vered, R. Freeman,
H. B. Yildiz, I. Willner, Angew. Chem. 2009, 121, 139–143; Angew.
Chem. Int. Ed. 2009, 48, 133–137; k) C. T. Wirges, J. Timper, M.
Fischler, A. S. Sologubenko, J. Mayer, U. Simon, T. Carell, Angew.
Chem. 2009, 121, 225–229; Angew. Chem. Int. Ed. 2009, 48, 219–
223.
[4] a) G. Mayer, D. Ackermann, N. Kuhn, M. Famulok, Angew. Chem.
2008, 120, 985–987; Angew. Chem. Int. Ed. 2008, 47, 971–973; b) K.
Borjesson, S. Preus, A. H. El-Sagheer, T. Brown, B. Albinsson, L. M.
Wilhelmsson, J. Am. Chem. Soc. 2009, 131, 4288–4293; c) V. Rain-
ˇ
dlovꢂ, R. Pohl, M. Sanda, M. Hocek, Angew. Chem. 2010, 122,
1082–1084; Angew. Chem. Int. Ed. 2010, 48, 1064–1066; d) H.
6882
www.chemeurj.org
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Cahovꢂ, L. Havran, P. Brazdilova, H. Pivonkova, R. Pohl, M. Fojta,
ence 2008, 320, 664–667; d) E. M. Sletten, C. R. Bertozzi, Angew.
Chem. 2009, 121, 7108–7133; Angew. Chem. Int. Ed. 2009, 48, 6974–
6998; e) P. V. Chang, J. A. Prescher, E. M. Sletten, J. M. Baskin, I. A.
Miller, N. J. Agard, A. Lo, C. R. Bertozzi, Proc. Natl. Acad. Sci.
USA 2010, 107, 1821–1826; f) A. A. Poloukhtine, N. E. Mbua, M. A.
Wolfert, G. J. Boons, V. V. Popik, J. Am. Chem. Soc. 2009, 131,
15769–15776; g) X. Ning, J. Guo, M. A. Wolfert, G. J. Boons,
Angew. Chem. 2008, 120, 2285–2287; Angew. Chem. Int. Ed. 2008,
47, 2253–2255; h) M. F. Debets, S. S. van Berkel, S. Schoffelen, F. P.
Rutjes, J. C. van Hest, F. L. van Delft, Chem. Commun. 2010, 97–99.
[14] a) M. L. Blackman, M. Royzen, J. M. Fox, J. Am. Chem. Soc. 2008,
130, 13518–13519; b) N. K. Devaraj, R. Weissleder, S. A. Hilder-
brand, Bioconjugate Chem. 2008, 19, 2297–2299; c) N. K. Devaraj,
R. Upadhyay, J. B. Haun, S. A. Hilderbrand, R. Weissleder, Angew.
Chem. 2009, 121, 7147–7150; Angew. Chem. Int. Ed. 2009, 48, 7013–
7016.
[15] a) I. Singh, J. S. Vyle, F. Heaney, Chem. Commun. 2009, 3276–3278;
b) V. Algay, I. Singh, F. Heaney, Org. Biomol. Chem. 2010, 8, 391–
397; c) I. Singh, F. Heaney, Org. Biomol. Chem. 2010, 8, 451–456.
[16] a) S. S. van Berkel, A. T. Dirks, M. F. Debets, F. L. van Delft, J. J.
Cornelissen, R. J. M. Nolte, F. P. Rutjes, ChemBioChem 2007, 8,
1504–1508; b) S. S. van Berkel, A. T. Dirks, S. A. Meeuwissen, D. L.
Pingen, O. C. Boerman, P. Laverman, F. L. van Delft, J. J. Cornelis-
sen, F. P. Rutjes, ChemBioChem 2008, 9, 1805–1815.
M. Hocek, Angew. Chem. 2008, 120, 2089–2092; Angew. Chem. Int.
Ed. 2008, 47, 2059–2062; e) M. Vrabel, P. Horakova, H. Pivonkova,
L. Kalachova, H. Cernocka, H. Cahova, R. Pohl, P. Sebest, L.
Havran, M. Hocek, M. Fojta, Chem. Eur. J. 2009, 15, 1144–1154.
[5] a) G. H. Clever, C. Kaul, T. Carell, Angew. Chem. 2007, 119, 6340–
6350; Angew. Chem. Int. Ed. 2007, 46, 6226–6236; b) J. Mꢃller, Eur.
J. Inorg. Chem. 2008, 3749–3763; c) K. Tanaka, G. H. Clever, Y.
Takezawa, Y. Yamada, C. Kaul, M. Shionoya, T. Carell, Nat. Nano-
technol. 2006, 1, 190–194; d) H. Yang, C. K. McLaughlin, F. A.
Aldaye, G. D. Hamblin, A. Z. Rys, I. Rouiller, H. F. Sleiman, Nat.
Chem. Biol. 2009, 5, 390–396; e) S. Johannsen, N. Megger, D.
Bçhme, R. K. Sigel, J. Mꢃller, Nat. Chem. 2010, 2, 229–234.
[6] S. H. Weisbrod, A. Marx, Chem. Commun. 2008, 5675–5685.
[7] a) R. Huisgen, Angew. Chem. 1963, 75, 604–637; Angew. Chem. Int.
Ed. Engl. 1963, 2, 565–632; b) R. Huisgen, G. Szeimies, L. Mçbius,
Chem. Ber. 1967, 100, 2494–2507; c) V. Hong, S. I. Presolski, C. Ma,
M. G. Finn, Angew. Chem. 2009, 121, 10063–10067; Angew. Chem.
Int. Ed. 2009, 48, 9879–9883.
[8] C. W. Tornoe, C. Christensen, M. Meldal, J. Org. Chem. 2002, 67,
3057–3064.
[9] V. V. Rostovtsev, L. G. Green, V. V. Fokin, K. B. Sharpless, Angew.
Chem. 2002, 114, 2708–2711; Angew. Chem. Int. Ed. 2002, 41, 2596–
2599.
[10] P. M. E. Gramlich, C. T. Wirges, A. Manetto, T. Carell, Angew.
Chem. 2008, 120, 8478–8487; Angew. Chem. Int. Ed. 2008, 47, 8350–
8358.
[17] a) W. Song, Y. Wang, J. Qu, M. M. Madden, Q. Lin, Angew. Chem.
2008, 120, 2874–2877; Angew. Chem. Int. Ed. 2008, 47, 2832–2835;
b) W. Song, Y. Wang, J. Qu, Q. Lin, J. Am. Chem. Soc. 2008, 130,
9654–9655; c) Y. Wang, W. Song, W. J. Hu, Q. Lin, Angew. Chem.
2009, 121, 5434–5437; Angew. Chem. Int. Ed. 2009, 48, 5330–5333.
[18] K. Gutsmiedl, C. T. Wirges, V. Ehmke, T. Carell, Org. Lett. 2009, 11,
2405–2408.
[19] a) M. D. Best, Biochemistry 2009, 48, 6571–6584; b) P. Kele, G.
Mezo, D. Achatz, O. S. Wolfbeis, Angew. Chem. 2009, 121, 350–353;
Angew. Chem. Int. Ed. 2009, 48, 344–347; c) P. Kele, X. Li, M. Link,
K. Nagy, A. Herner, K. Lorincz, S. Beni, O. S. Wolfbeis, Org.
Biomol. Chem. 2009, 7, 3486–3490.
[20] M. Kuwahara, J. Nagashima, M. Hasegawa, T. Tamura, R. Kitagata,
K. Hanawa, S. Hososhima, T. Kasamatsu, H. Ozaki, H. Sawai, Nu-
cleic Acids Res. 2006, 34, 5383–5394.
[21] See the Supporting Information.
[22] B. Horten, Nature 1998, 396, 391–392.
[11] a) G. A. Burley, J. Gierlich, M. R. Mofid, H. Nir, S. Tal, Y. Eichen,
T. Carell, J. Am. Chem. Soc. 2006, 128, 1398–1399; b) J. Gierlich,
G. A. Burley, P. M. Gramlich, D. M. Hammond, T. Carell, Org. Lett.
2006, 8, 3639–3642; c) J. Gierlich, K. Gutsmiedl, P. M. Gramlich, A.
Schmidt, G. A. Burley, T. Carell, Chem. Eur. J. 2007, 13, 9486–9494;
d) P. M. E. Gramlich, S. Warncke, J. Gierlich, T. Carell, Angew.
Chem. 2008, 120, 3491–3493; Angew. Chem. Int. Ed. 2008, 47, 3442–
3444; e) F. Seela, V. R. Sirivolu, Org. Biomol. Chem. 2008, 6, 1674–
1687; f) V. R. Sirivolu, P. Chittepu, F. Seela, ChemBioChem 2008, 9,
2305–2316; g) F. Seela, H. Xiong, P. Leonard, S. Budow, Org.
Biomol. Chem. 2009, 7, 1374–1387.
[12] a) S. Thyagarajan, N. N. Murthy, A. A. Narducci Sarjeant, K. D.
Karlin, S. E. Rokita, J. Am. Chem. Soc. 2006, 128, 7003–7008; b) F.
Wolbers, P. ter Braak, S. Le Gac, R. Luttge, H. Andersson, I.
Vermes, A. van den Berg, Electrophoresis 2006, 27, 5073–5080.
[13] a) N. J. Agard, J. A. Prescher, C. R. Bertozzi, J. Am. Chem. Soc.
2004, 126, 15046–15047; b) J. M. Baskin, J. A. Prescher, S. T. Laugh-
lin, N. J. Agard, P. V. Chang, I. A. Miller, A. Lo, J. A. Codelli, C. R.
Bertozzi, Proc. Natl. Acad. Sci. USA 2007, 104, 16793–16797;
c) S. T. Laughlin, J. M. Baskin, S. L. Amacher, C. R. Bertozzi, Sci-
[23] H. A. Held, S. A. Benner, Nucleic Acids Res. 2002, 30, 3857–3869.
[24] W. Fliege, R. Huisgen, Liebigs Ann. Chem. 1973, 2038–2047.
Received: February 10, 2010
Published online: May 10, 2010
Chem. Eur. J. 2010, 16, 6877 – 6883
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