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Lipid-Anchored Oligonucleotides for Stable Double-Helix Formation in Distinct Membrane Domains.

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Communications
DOI: 10.1002/anie.200600822
Nanobiotechnology
Lipid-Anchored Oligonucleotides for Stable DoubleHelix Formation in Distinct Membrane Domains**
Anke Kurz, Andreas Bunge, Anne-Katrin Windeck, Maximilian Rost,
Wolfgang Flasche, Anna Arbuzova, Denise Strohbach, Sabine M!ller,
J!rgen Liebscher,* Daniel Huster,* and Andreas Herrmann*
Angewandte
Chemie
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2006 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
Angew. Chem. Int. Ed. 2006, 45, 4440 –4444
Angewandte
Chemie
L
ipid-anchored RNA and DNA oligonucleotides have
emerged as attractive chimeric molecules for various
applications in nanobiotechnology, in cell biology, and for
the development of therapeutic strategies in medicine.[1, 2] For
example, lipophilic oligonucleotides have been synthesized
to improve cellular uptake of antisense single-stranded DNA
(ssDNA) and interference RNA.[1, 3] Recently, efficient
silencing of an endogenous apolipoprotein B in mice has
been reported upon injection of short interfering RNA
molecules (siRNA) coupled to a cholesterol anchor. [4]
Depending on the application, lipid-anchored oligonucleotides should meet various requirements, such as easy and
flexible synthesis, efficient and stable membrane incorporation, and selective binding of complementary DNA strands.
For various purposes a distinct lateral organization of
oligonucleotides in membranes is desirable. By selecting
appropriate lipid anchors, the enrichment of lipophilic
oligonucleotides in specific lipid domains can be achieved[5]
which allows distinct functional compartments to be created
on a membrane surface. Herein we describe the synthesis and
application of a lipidated ssDNA molecule that stably inserts
into lipid membranes, displays the oligonucleotide on the
vesicular surface, binds complementary DNA strands by
formation of Watson–Crick base pairs, and preferentially
sequesters into liquid-disordered membrane domains.
The oligonucleotide LT23 mer is a 23 mer consisting of
21 thymidine units and two lipophilic nucleotides L in positions 1 and 8 with a-tocopherol units as lipophilic anchors
(Scheme 1). LT23 mer was synthesized on a DNA synthesizer
applying phosphoramidite methodology (Supporting Infor[*] Dr. A. Kurz,[+] Dr. A.-K. Windeck, M. Rost, Dr. A. Arbuzova,
Prof. Dr. A. Herrmann
Institute of Biology/Biophysics
Humboldt University of Berlin
Invalidenstrasse 42, 10115 Berlin (Germany)
Fax: (+ 49) 30-2093-8585
E-mail: andreas.herrmann@rz.hu-berlin.de
Dipl.-Phys. A. Bunge,[+] Priv.-Doz. Dr. D. Huster
Junior Research Group “Structural Biology of Membrane Proteins”
Institute of Biotechnology
Martin-Luther University Halle-Wittenberg
Kurt-Mothes-Strasse 3, 06120 Halle (Germany)
Fax: (+ 49) 345-55-27013
E-mail: daniel.huster@biochemtech.uni-halle.de
Dr. W. Flasche, Prof. Dr. J. Liebscher
Institute of Chemistry
Humboldt University Berlin
Brook-Taylor-Strasse 2, 12489 Berlin (Germany)
Fax: (+ 49) 30-2093-7552
E-mail: liebscher@chemie.hu-berlin.de
Dipl.-Chem. D. Strohbach, Prof. Dr. S. MJller
Institute of Biochemistry
Ernst-Moritz-Arndt University Greifswald
Soldmannstrasse 16, 17487 Greifswald (Germany)
[+] These authors contributed equally.
[**] This study was supported by the BMBF grant No. 0312018. We
thank Mrs Charlott Peters for preparative support.
Supporting information for this article is available on the WWW
under http://www.angewandte.org or from the author.
Angew. Chem. Int. Ed. 2006, 45, 4440 –4444
Scheme 1. Structure of the lipophilic nucleoside L building block used
as a monomer in the synthesis of lipid modified oligonucleotides. The
sequence of the LT23 mer is given below
mation). The lipophilic tocopherylpropinylcytidine L was
obtained by Sonogashira coupling of 5-iodocytidine with
racemic O-propargyltocopherol using a recently published
procedure[6] and transformed into the 5’-DMTr-protected 3’diisopropylaminocyanethylphosphor amidite by conventional
methods. This method provides a high flexibility because
lipophilic nucleotides can be introduced in any position of an
oligonucleotide.
Fluorescence microscopy was used to visualize the
membrane affinity of LT23 mer and the ability to bind a
complementary A20 mer DNA strand. Giant unilamellar
vesicles (GUVs) were prepared containing 1 mol %
LT23 mer and the fluorescent phospholipid analogue NNBD-PE (0.5 mol %; see Experimental Section). To demonstrate binding of the complementary DNA strand, an
adenosine 20 mer tagged either on its 3’ (3’Rh-A20 mer) or
on its 5’ terminus (5’Rh-A20 mer) with the fluorophore
rhodamine (Rh) was added. The binding of these complementary strands to the LT23 mer of the GUV membranes
could be identified by the Rh fluorescence (Figure 1, only
Figure 1. Binding of a Rh-A20 mer to N-NBD-PE-labeled (0.5 mol %)
POPC–GUVs containing LT23 mer (about 1 mol %) at room temperature. Left: differential interference contrast (DIC) image. Middle:
Fluorescence of N-NBD-PE and right: fluorescence of 5’Rh-A20 mer.
shown for 5’Rh-A20 mer). Fluorescence was homogeneous
over the whole membrane indicating that LT23 mer is
homogeneously distributed within the membrane. The
GUVs could not be labeled by Rh-tagged T20 mer. In the
absence of LT23 mer no labeling of GUVs with fluorescent
A20 mers was observed (not shown).
To examine further binding of complementary adenine
oligonucleotides to membrane-bound LT23 mer we employed
a fluorescence resonance energy transfer (FRET) assay.
2006 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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Large unilamellar vesicles (LUVs) containing either POPC
and LT23 mer or only POPC (control) were prepared in the
presence of N-NBD-PE (0.5 mol %). The fluorescent NBD
moiety, which serves as a donor for FRET, is covalently
attached to the lipid head group. LUVs were incubated with
the 3’Rh-A20 mer with Rh being the fluorescence acceptor.
We measured a strong FRET when LUVs containing
LT23 mer were incubated with 3’Rh-A20 mer (Figure 2 a).
FRET was not observed for control LUVs (Figure 2 b).
Figure 2. Fluorescence of N-NBD-PE-labeled POPC–LUVs a) with or
b) without LT23 mer, were incubated in the absence (c) or presence (a) of 3’Rh-A20 mer at room temperature. A strong FRET was
observed when 3’Rh-A20 mers were added to LT23 mer containing
LUVs (LT23 mer:3’Rh-A20 mer = 1:1). Vertical dotted lines are fluorescence maxima of NBD and rhodamine, respectively. c) Kinetics of 3’RhA20 mer (500 nm) binding to LT23 mer-containing LUVs measured
by the decrease of NBD fluorescence owing to FRET (excitation
460 nm; emission 532 nm). At t = 280 s an unlabelled T20 mer (5 mm)
was added. For three selected time points fluorescence spectra are
shown indicating the absence, increase, and decrease of FRET,
respectively.
Binding of 3’Rh-A20 mer to membrane-bound LT23 mer
was rapid, with a half time of about 20 s (Figure 2 c), and
reversible. When (unlabelled) T20 mers were added in 10-fold
excess with respect to LT23 mer, a decrease of FRET was
observed owing to the competition of the T20 mer for 3’RhA20 mer (Figure 2 c).
The structure of the membrane-associated complex of
LT23 mer and A20 mer was investigated by NMR spectroscopy. As a control, solution NMR studies of stoichiometric
mixtures of A20 mer and T20 mer were carried out. Base-pair
formation and stacking leads to characteristic changes of the
chemical shifts.[7] Slices of the 1H NOESY spectra are shown
(Figure 3). In the presence of LUVs, 1D spectra also
contained intense signals arising from lipids. In contrast, the
NOESY slices only showed the signals of the base and the
sugar moieties. Figure 3 a depicts the 1H NMR spectrum of
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Figure 3. Slices of 1H NOESY NMR spectra of free and membrane
bound oligonucleotides in the absence and presence of a complementary strand. a) T20 mer in buffer, b) A20 mer and T20 mer (1:1) in
buffer; c) LT23 mer associated with POPC membranes; d) LT23 mer
associated with POPC membranes in the presence of A20 mers. Lines
are drawn to guide the eye. Purple bars denote upfield and yellow bars
denote downfield shifts. A summary of the chemical shift changes (in
ppm) for the A and T moieties is given in the structural formula above.
Slight chemical shift changes of the H5’/H5’’ signals may indicate a
conformational change in the furanose upon double-strand formation.
the T20 mer. In the presence of the complementary A20 mer
strands, all the resonance signals were shifted in a specific
manner (Figure 3 b). The LT23 mer bound to POPC LUVs
(Figure 3 c) showed exactly identical 1H NMR chemical shifts
as isolated T20 mer (cf. Figure 3 a). After addition of the
complementary A20 mer, identical relative shifts compared to
the A20 mer–T20 mer double helix were observed (Figure 3 d). The chemical shift changes of A20 mer after binding
2006 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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Angewandte
Chemie
to either T20 mer or LT23 mer were also identical (not
shown). This result strongly suggests that very similar
structures are formed and neither the lipid-anchor nor the
membrane proximity interfere with the formation of the
DNA double helix through Watson–Crick base pairing.
It has been shown that GUVs prepared from a lipid
mixture of POPC, cholesterol, and sphingomyeline (1:1:1)
form liquid-disordered and liquid-ordered domains.[8]
Domains can be visualized through fluorescence microscopy
by employing fluorescent lipid analogues, which preferentially enrich in one of the two domains. The lipophilic
fluorophore Merocyanine 540 accumulates in the liquiddisordered domain[9] as verified by N-NBD-PE which is
another domain marker for liquid-disordered domains (data
not shown).[8b] GUVs of the above mentioned a lipid mixture
(1:1:1 (molar ratio)) were used with LT23 mer (about
1 mol %) and Merocyanine 540. Subsequently, complementary oligonucleotides (5’FITC-A20 mer) were added. In contrast to the homogeneous distribution found in pure POPC–
GUVs, the LT23 mer with the complementary strand 5’FITCA20 mer was recruited into the liquid-disordered domain as
identified by Merocyanine 540 staining (Figure 4). Such a
preferential recruitment can be explained by the bulky atocopherol anchor which could pack more easily into a liquiddisordered phase. Preferential partitioning of isoprenylated
proteins into liquid-disordered domains has also been
observed.[10]
In summary, we have described a simple strategy to
synthesize lipophilic nucleosides that can be used to produce
large quantities of lipid-anchored oligonucleotides with
variable length and primary structure. These molecules
insert into lipid membranes and expose the ssDNA to the
aqueous phase. Complementary strands are recognized and
Figure 4. DIC and fluorescence images indicating that LT23 mer binds
preferentially to liquid-disordered membrane domains. GUVs were
prepared from a mixture of POPC/sphingomyeline/cholesterol (1:1:1
molar ratio) containing about 1 mol % LT23 mer and incubated with
5’FITC-A20 mer in KCl buffer (50 mm), pH 8.0, at room temperature.
LT23 mer–5’FITC-A20 mer complexes are recruited to the liquid-disordered domain which was detected by Merocyanine 540 (M540). This
dye is known to enrich in liquid-disordered domains.
Angew. Chem. Int. Ed. 2006, 45, 4440 –4444
self-assemble into double stranded DNA with Watson–Crick
base pairing. Thus, molecules, vesicles, drugs, or biologically
active RNA can be associated with membrane or cellular
surfaces at will. An important observation is the recruitment
of lipid-anchored oligonucleotides to specific lipid domains.
This property could be used to tether double-stranded DNA
to specific domains of membrane surfaces and thus to
functionalize these domains. Given that lipid domains play
an important role in endocytosis,[11] enrichment of lipophilic
oligonucleotides to those domains could be explored to
enhance cellular uptake. Thus, nanostructures may be built
and various cell biological or medical applications can be
foreseen.
Experimental Section
1-Palmitoyl-2-oleoyl-sn-glycerophosphocholine (POPC), sphingomyeline (SM), and the fluorescent lipid N-(7-nitro-2,1,3-benzoxadiazol-4-yl)-1,2-dipalmitoyl-sn-glycero-3-phosphatidyl-ethanolamine
(N-NBD-PE) were obtained from Avanti Polar Lipids (Birmingham,
USA). Cholesterol and Merocyanine 540 were from Sigma (Deisenhof, Germany). DNA-oligonucleotides with or without a covalently
attached fluorophores (rhodamine (Rh) or fluorescein (fluoresceinisothiocyanate (FITC))) were synthesized by BioTez (Berlin-Buch,
Germany). All solutions used were buffered with 10 mm HEPES,
pH 8. The preparation of unilamellar vesicles was performed in
sucrose solution (80 to 250 mm). For washing and dilution of the
vesicles and preparation of multilamellar vesicles buffered KCl, KCl/
glucose, or glucose solutions of equal osmolarity were used (see
below).
For preparation of LUVs, the lipid mixture dissolved in chloroform was evaporated in a round-bottomed flask by a rotary
evaporator forming a thin lipid layer on the flask wall. The lipid
film was resuspended by manual swirling of the glass flask.
Subsequently, LUVs were formed by subjecting this suspension to
five freeze–thaw cycles and extrusion through a polycarbonate filter
(Nucleopore GmbH, TGbingen, Germany) of 100 nm diameter using
an extruder (Lipex Biomembranes, Vancouver).[12] For membrane
incorporation, LT23 mer was added to an aliquot of the LUVs and the
suspension was again subjected to five freeze–thaw cycles. Finally,
LUVs were ultracentrifuged (120 000 H g) and unbound LT23 mer and
any micelles remaining were removed (supernatant).
GUVs were generated at room temperature by the electroformation technique in a chamber made of indium tin oxide (ITO)
coated glass slides.[13] Lipids were dissolved in chloroform (0.25 mg
lipid mL 1) and 70 mL of solved lipids were deposited in small droplets
on each slide. The solvent was evaporated in a desiccator (10 mbar) at
room temperature for 60 min. Subsequently, the chamber was closed
by sealed with silicon paste. The sucrose solution (250 mm sucrose,
10 mm HEPES, 0.02 % NaN3) was injected with a syringe through a
micropore filter immediately before connecting the completely sealed
chamber to the generator. LT23 mer was added to the sucrose solution
to allow its incorporation during GUV preparation. The voltage of
the applied AC-field was increased in steps every 6 min from 20 mV
up to 1.1 V while continuously increasing the frequency from 4 Hz to
10 Hz within the first minute. The AC-field was applied for 3–12 h. To
complete the procedure, voltage was raised to 1.3 Vand the frequency
lowered to 4 Hz. The chamber was then stored in a refrigerator at
4 8C.
Fluorescence spectra and kinetics were recorded using an
Aminco Bowman spectrometer series 2 (SLM-Aminco, Rochester,
NY). N-NBD-PE was excited at 460 nm (slit width 4 nm) and
fluorescence spectra were recorded between 470 and 610 nm with a
scan rate of 1 nm s 1 whereas for kinetic measurements the emission
wavelength was set to 532 nm (slit width 4 nm). GUVs were examined
2006 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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4443
Communications
with an Olympus IX-81 inverted fluorescence microscope (Olympus,
Hamburg, Germany) equipped with a cooled CCD camera (SPOT
slider, Visitron Systems, Puchheim, Germany). Images were acquired
using 100 H Plan-APO oil immersion objective with the appropriate
differential interference contrast (DIC) optics and fluorescence filter
sets: BP 470-490, FT 505, and BP 510-550 (NBD, FITC); BP 530–550,
FT 580, and LP 590 (rhodamine, Merocyanine 540).
For NMR spectroscopy experiments, aliquots of LT23 mer were
added to extruded 100 nm POPC vesicle suspensions (20 mm in D2O
buffer (10 mm HEPES, 100 mm NaCl, pD 7.0)) at a LT23 mer to
POPC molar ratio of 1:100. NMR experiments were carried out on a
Bruker DRX600 or Avance 700 NMR spectrometer at a resonance
frequency of 600.13 MHz or 700.13 MHz, respectively, at 30 8C.
Spectra were acquired at a spectral width of 7 kHz with a 908 pulse
length of 10.3 ms. For phase sensitive NOESY experiments (mixing
time 300 ms) 480 complex data points were collected in the t1
dimensions with 32 or 64 scans per increment at a 4 s relaxation
delay. Subsequently, aliquots of A20 mer was added at a 1:1 molar
ratio with respect to LT23 mer and NMR spectra were collected again.
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Received: March 2, 2006
.
Keywords: FRET (fluorescence energy resonance transfer) ·
molecular recognition · nanostructures · NMR spectroscopy ·
nucleic acids
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