Lipid-bilayer-spanning DNA nanopores with a bifunctional porphyrin anchor

Article abstract of DOI:10.1002/anie.201305765

Holding tight: An artificial membrane nanopore assembled from DNA oligonucleotides carries porphyrin tags (red), which anchor the nanostructure into the lipid bilayer. The porphyrin moieties also act as fluorescent dyes to aid the microscopic visualization of the DNA nanopore. Copyright

Full text of DOI:10.1002/anie.201305765

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Chemie
DOI: 10.1002/anie.201305765
DNA Nanotechnology
Hot Paper
Lipid-Bilayer-Spanning DNA Nanopores with a Bifunctional Porphyrin
Anchor**
Jonathan R. Burns, Kerstin Gçpfrich, James W. Wood, Vivek V. Thacker, Eugen Stulz,
Ulrich F. Keyser, and Stefan Howorka*
Chemistry is in a powerful position to enhance the capabilities
of DNA nanotechnology.[1,2] DNA origami offers rationally
designed nanoscale structural frameworks,[2,3] yet chemistry
can advance them by adding tailored functionality through
the selective modification of nucleic acids with chemical tags.
This synergistic approach has helped create new nanoscale
devices, ranging from affinity nanoarrays capable of binding
proteins[4] and quantum dots,[5] to nanoplatforms for single-
molecule photochemistry,[6] and to molecular display agents
for biosensing.[7,8] Herein, we present a unique chemical
strategy for enlarging and enriching the emerging class of
membrane-spanning nanopores composed of folded
DNA.[9,10] We show that solely two porphyrin-based hydro-
phobic tags achieve the otherwise energetically unfavorable
anchoring of the highly negatively charged DNA nano-
structure into the hydrophobic core of lipid bilayers. This very
small number of porphyrin tags considerably simplifies the
currently available chemical strategies for bilayer anchoring
of nanopores. The aromatic porphyrin tags are also fluores-
cent, and hence facilitate the microscopic visualization of
DNA-based membrane channels. Our generic route for dual-
functional chemical tags can likely be applied to many other
DNA designs, and will help broaden experimental access to
versatile DNA-origami pores.
biotechnological interest, because they are able to replicate
the transport of water-soluble molecules across bilayers[13] for
applications in research or biosensing.[14] Inspired by nano-
funnels[15] and porous nanoplates,[16] DNA-origami pores have
been designed to insert into lipid bilayers.[9,10] In these studies,
hydrophobic chemical tags were deliberately positioned to
anchor the strongly hydrophilic DNA structures into bilayers.
The tags previously described were either cholesterol-based
lipid anchors covalently attached to DNA strands,[9] or ethyl-
modified phosphorothioate groups that replace the negatively
charged backbone phosphate to form a hydrophobic belt to
mimic natural protein pores.[10] The former tag was placed at
up to 26 strategic positions of the pore, whereas the latter
group was introduced 72 times into a DNA-origami structure.
With the intent to simplify nanopore design, and move
towards minimal chemical intervention, the present study
explores whether other chemical tags of greater hydropho-
bicity can achieve membrane anchoring using a very small
number of tag copies.
We surmised that a porphyrin derivative would satisfy the
criterion of strong hydrophobicity because of its large
aromatic core. Porphyrin has a van der Waals surface area
approximately 12-times higher than ethane, and the area can
be further increased with additional aromatic substituents. In
addition, most porphyrins are chromophores with a fluores-
cence emission at 656 nm and can thereby act as powerful
visualization tags. Moreover, inserting porphyrins into lipid
bilayers leads to a characteristic shift in their fluorescence
spectrum, which offers an additional experimental handle to
confirm membrane anchoring.[17,18]
Membrane-spanning nanopores composed of folded and
structurally defined DNA are the most recent and striking
example of a long series of artificial or synthetic membrane
channels,[11] including those made of porphyrins.[12] In general,
biomimetic and engineered nanopores are of scientific and
For the creation of membrane-spanning DNA nanopores,
we selected the tetraphenylporphyrin (TPP) tag (Fig-
ure 1a)[19,20] which matches the requirements in terms of
hydrophobicity and fluorescence emission and can be easily
coupled to DNA.[21] Acetylene-TPP was attached to deoxy-
uridine through a Sonogashira coupling to achieve a rigid
linkage. The site-specific insertion of the modified nucleoside
into oligo-deoxyribonucleotides was accomplished using
standard phosphoramidite chemistry as previously described
(Figure 1a; see also the Supporting Information, Figures S1–
S4).[19,21]
In our nanopore design (Figure 1b), a total of six DNA
oligonucleotides were folded into six DNA duplexes, which
are interconnected by crossovers to add structural stability.
The threading of the oligonucleotides through the duplexes is
indicated by the green lines in Figure 1b (see also Figure S5).
The resulting six-helix bundle has a width of 5.5 nm, a height
of 14 nm, and an inner channel diameter of approximately
2 nm. The overall dimensions are close to those of our
[*] Dr. J. R. Burns, Dr. S. Howorka
Department of Chemistry, University College London
20 Gordon Street, London WC1H OAJ (UK)
E-mail: s.howorka@ucl.ac.uk
K. Gçpfrich, V. V. Thacker, Dr. U. F. Keyser
Cavendish Laboratory, University of Cambridge
Cambridge CB3 0HE (UK)
J. W. Wood, Dr. E. Stulz
School of Chemistry, University of Southampton
Southampton SO17 1BJ (UK)
[**] Funded by the Leverhulme Trust (RPG-170), UCL Chemistry, the
EPSRC (Institutional Sponsorship Award), and the NPL. K.G. and
U.F.K. acknowledge funding from his ERC starting grant. V.V.T.
acknowledges funding from the Cambridge Commonwealth Trust,
the Jawaharlal Nehru Memorial Trust, and the Emmy Noether
program of the Deutsche Forschungsgemeinschaft. We thank Silvia
Hernꢀndez-Ainsa for assistance in acquiring DLS data and Hugh
Martin for preparing the ToC graphic.
Supporting information for this article is available on the WWW
under http://dx.doi.org/10.1002/anie.201305765.
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Figure 1. DNA-nanopore-carrying porphyrin-based lipid anchors.
a) Deoxyuridine bonded to tetraphenylporphyrin (TPP) through an
acetylene linkage at the 5 position of the nucleobase. b) A DNA
nanopore composed of six interconnected duplexes, represented as
cylinders. The six-component DNA oligonucleotides are shown in
green. The magenta porphyrin tags anchor the DNA nanopore into the
lipid bilayer. For reasons of visual clarity, only the porphyrin core
without the phenyl groups is shown. The TPP tags are not drawn to
scale.
Figure 2. Characterization of porphyrin-modified nanobarrels assem-
bled from DNA oligonucleotides. a) Agarose gel electrophoresis. Lane
1, 100 bp marker; lane 2, DNA nanopore; lane 3, TPP-DNA nanopore.
b) UV melting profile of a TPP nanopore. c) Dynamic light scattering
trace of TPP nanopore. d) AFM micrograph of individual nanopores
(arrows) of elongated nanopore assemblies, which are stabilized by
inter-barrel base stacking; scale bar=50 nm. Abs.=absorbance,
T=temperature.
previously published nanopore,[10] but the current design is
considerably simpler, as six rather than 14 strands are used.
Crucially, two porphyrin tags are positioned at the end of the
nanobarrel to facilitate its insertion into the bilayer in
a directional manner (Figure 1b). To facilitate the compar-
ison to our previous nanopore work, we used a design with
two DNA strands that contain phosphorothiate groups in
place of the phosphates in a number of positions in the
backbone. The thioate groups behave like the native neg-
atively charged phosphate groups at our experimental
conditions of pH 8.0 (Table S1).[10]
The DNA nanopore was assembled by heating and
cooling an equimolar mixture of four regular and two TPP-
modified DNA strands (for sequences, see Table S1). The
assembly mixture was characterized to confirm the correct
and successful formation of the DNA nanopore. Native gel
electrophoresis yielded a band, which migrated to the same
height as a control nanopore without the porphyrin anchor
(Figure 2a, lanes 2 and 3, respectively; main band co-migrat-
ing at the 550 bp marker). The tailing of the band for the
porphyrin DNA pore does not indicate unfolding, but is
rather caused by the very hydrophobic tag.[9,19,20] The
concerted assembly into the nanobarrels was also confirmed
by a single defined transition in the UV melting profiles,
(Figure 2b; 53.4 Æ 1.08C; n = 3); the opposite and unexpected
independent hybridization of the multicomponent DNA
duplexes of different melting temperatures would have led
to a very broad transition. Furthermore, dynamic light
scattering (DLS) established the monomeric nature of the
nanobarrels, as only a single peak with a hydrodynamic radius
of 5.5 Æ 0.1 nm was observed (Figure 2c). The radius is larger
than the calculated value of 4.9 nm,[22,23] but in line with the
accuracy of DLS measurements for related DNA nano-
structures.[23,24] The detailed dimensions of the nanobarrel
were established with atomic force microscopy (AFM)
analysis (Figure 2d). The apparent height of (2.20 Æ
0.25) nm was expected for tip-compressed hollow DNA
nanostructures.[8,25] Similarly, the AFM-derived length and
width of (20.4 Æ 4.5) nm and (9.7 Æ 2) nm (full width at half
maximum), respectively, were in good agreement with the
theoretical dimensions (14 nm and 5.5 nm) after correcting
for tip-deconvolution.[10,26]
Having completed their structural characterization, we
investigated whether the porphyrin-tagged DNA nanopore
can be stably anchored into lipid bilayers. In this examination
we took advantage of the fluorescence properties of the
porphyrin tag. Once incubated with giant unilamellar vesicles,
the tagged nanopores could be visualized using fluorescence
microscopy (Figure 3a, bottom). A bright-field image docu-
mented the shape of the vesicle (Figure 3a, top). The bright
fluorescence spots in the ring-shaped bilayer may represent
individual DNA nanopores, given that the imaging conditions
of our custom-built set-up are powerful enough to detect
single fluorophores.[27] However, as the DNA pores rapidly
diffuse out of the focal plane (Supporting Information,
Movie SM1) and exhibit surprising photostability, it is hard
to detect individual pores by single-fluorophore photobleach-
ing. The anchoring of the nanopore into the bilayer was
confirmed by independent spectroscopic analysis based on
Figure 3. Porphyrin tags anchor DNA nanopores into lipid bilayers.
a) Microscopic bright-field (top) and fluorescence (bottom;
lexc =532 nm) images of a DPhPC vesicle containing DNA nanopores;
scale bars=5 mm. b) The fluorescence emission spectrum for the TPP
nanopore (c) is shifted upon insertion into vesicle bilayers (a;
lexc =424 nm). F=fluorescence intensity.
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the known fluorescence shift of porphyrin. As illustrated in
Figure 3b, the fluorescence emission maxima at 605 and
653 nm shifted upon bilayer insertion, by 2 nm and 1 nm,
respectively. These changes are in line with previous studies
on the membrane binding of individual porphyrin-carrying
DNA duplexes[17] and were enhanced by the use of zinc-
porphyin tags (Figure S6). Membrane-anchoring also led to
an increase of the thermal stability of the DNA nanopore by
almost 68C to 59.2 Æ 1.28C, as determined by UV-based
melting profiles (data not shown). The fluorescence measure-
ments do not, however, rule out that a proportion of the DNA
nanobarrels anchor with one TPP group and lie parallel to the
membrane plane.
To provide evidence that the porphyrin nanopore is
capable of spanning the membrane, we carried out current
recordings to measure the ion flow across the lipid bilayers.
Lipid vesicles were first incubated with DNA nanopores, then
suctioned onto a nanocapillary, and, after application of
a voltage pulse, analyzed by recording the ionic current
caused by the transmembrane potential. A representative
current trace for a single pore at standard electrolyte
conditions and potentials (Figure 4a) demonstrates that the
inserted pores punctured the bilayer with nanoscale holes of
steady ionic current. Results from multiple nanopore record-
ings yielded a distribution of conductances with a maximum
of 250 pS (Figure 4b and 4d). This value is slightly lower than
the average obtained from nanopores with a similar six-helix
core structure,[10] which could be due to the divergent designs,
as the present nanopore inserts at its terminus, whereas the
previously reported one crosses the bilayer at the middle
section. In combination with the lateral membrane pressure
of the capillary-held vesicles, slightly different inner-channel
cross-sections could result. Alternatively, the pore might
insert into the membrane in an angled rather than the
assumed perpendicular orientation, thus leading to partial
obstruction of the ion conduction path. These factors might
also contribute to the observed broad distribution of con-
ductances (Figure 4b), which could additionally be due to
inferring of single-pore conductances from multistep closures
(Figure 4d) with a potentially different minimal residual
current, or to some structural variability of the assembled
DNA network, as the RMS current noise changed between
channels (not shown). However, under additional analysis,
the porphyrin nanopores exhibited almost ohmic behavior in
current–voltage curves (Figure 4c), which implies overall
structural integrity within the DNA nanoarchitecture and
stable insertion into the lipid bilayer.
In summary, our overarching research objective to
synergistically combine chemistry with DNA nanotechnology
has led to the creation of membrane-spanning DNA nano-
pores that carry two bifunctional tags. Exemplified by
a relatively simple nanostructure, our strategy of minimal
chemical intervention drastically increases flexibility in nano-
pore design and will enable more complex architectures that
can perform higher-level functions for applications in bio-
mimetic research and biosensing.
Experimental Section
Design of the nanopore structure: The nanobarrel was designed using
the caDNAno software.[28] Several suggested scaffold and staple
strands were terminally linked to form a more stable structure
composed of only six DNA strands (Figure S5). Using a molecular
model, which was generated with Macromodel in combination with
caDNAno, the positions for attaching porphyrins were selected to be
on two opposite duplexes.
Synthesis and purification of porphyrin-DNA, and nanopore
assembly: Tetraphenylporphyrin-tagged deoxyuridine was synthe-
sized and incorporated into DNA oligonucleotides as described
(Figures S1 and S2).[19] The purity of the strands was confirmed by
PAGE, whereas the yield was determined by UV/Vis spectroscopy
(Figures S3 and S4). Nanopores were assembled by heating an
equimolar mixture of the six strands (1 mm each), dissolved in
buffer A (KCl (1m), tris(hydroxymethylaminomethane) (Tris; 50 mm,
pH 8.0); total volume = 1000 mL) at 958C for 5 min, followed by
cooling to 168C at a rate of 0.258CminÀ1 in a Varian Cary 300 Bio
UV/Vis spectrophotometer, equipped with a Peltier cooling element.
Nanobarrel characterization was accomplished with native gel
electrophoresis, UV/Vis spectroscopy, dynamic light scattering, and
atomic force microscopy. The assembled DNA barrels were analyzed
using agarose (0.8%) gel electrophoresis in a standard TBE buffer,
supplemented with MgCl2 (11 mm) and running conditions of 80 V,
80 min, 88C, followed by ethidium bromide staining. The melting
point analysis was performed on samples (0.2 mm) dissolved in
buffer A, using a heating rate of 0.58CminÀ1. DLS experiments
were conducted on a Zetasizer Nano S from Malvern,[29] using DNA
samples (0.25 mm) in buffer A. AFM analysis was carried out by first
Figure 4. Porphyrin-DNA nanopores span lipid bilayers. a) Current
trace of membrane-inserted TPP nanopores in 1m KCl, 50 mm Tris,
pH 8.0, acquired at a filtering and sampling frequency of 2 kHz and
10 kHz, respectively. A prolonged trace is shown in Figure S7. The
RMS noise increases from 1.30 pA to 1.43 pA from 0 mV to 100 mV.
b) Histogram of single-pore conductances, as determined from single-
channel recordings or from current steps of multichannel traces (panel
D). c) Voltage–current curve, obtained from 8 independent recordings.
d) Current trace of multiple pores, with the stepwise reduction
interpreted as separate pore closures.
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adsorbing DNA barrels onto mica, following a modified version of
a published procedure.[30] Freshly cleaned mica was incubated with
a solution of MgCl2 (3 mm) for 5 min. The surface was then incubated
with a 20 nm solution of the DNA-barrel solution. AFM topo-
graphical images were acquired in situ at RTwith a Multimode atomic
force microscope as described.[10]
Lipid vesicle formation: Giant unilamellar vesicles (GUVs) for
imaging and current recordings were prepared using an electro-
formation unit (Vesicle Prep Pro, Nanion Technologies, Germany) as
described.[31] The GUVs (1–30 mm) were stored at 48C for up to
a week.
DOI: 10.1021/ar400014r; G. W. Gokel, S. Negin, Acc. Chem. Res.
2013, DOI:10.1021/ar400026x; J. Montenegro, M. R. Ghadiri,
J. R. Granja, Acc. Chem. Res. 2013, DOI: 10.1021/ar400061d; Y.
Zhao, H. Cho, L. Widanapathirana, S. Zhang, Acc. Chem. Res.
2013, DOI: 10.1021/ar300337f.
[12] A. Satake, M. Yamamura, M. Oda, Y. Kobuke, J. Am. Chem.
Soc. 2008, 130, 6314 – 6315; M. Boccalon, E. Iengo, P. Tecilla, J.
Am. Chem. Soc. 2012, 134, 20310 – 20313; U. Devi, J. R. Brown,
A. Almond, S. J. Webb, Langmuir 2011, 27, 1448 – 1456.
[13] J. M. Pagꢁs, C. E. James, M. Winterhalter, Nat. Rev. Microbiol.
2008, 6, 893 – 903.
Ionic current recordings: The recordings were performed using
a nanobilayer setup as previously described.[32] On this setup, the
bilayers were formed reproducibly by bursting GUVs on the tip of
a nanocapillary (200 nm diameter). GUVs were incubated with DNA
nanopores (30 nm) for 1 h at RT in buffer A. Bilayers that held DNA
nanopores were identified by their lowered seal resistances. Nano-
pore incorporation was triggered by applying a voltage pulse. Ionic
current data were acquired using an Axopatch 200B amplifier and
analyzed as described.[32,33]
Fluorescence microscopic imaging: DNA nanopores (3 nm) were
incubated with GUVs (0.5 mL) in buffer A for 1 h at RT. The
suspension (20 mL) was then imaged using a previously described
microscope setup[34] with a laser (l = 532 nm, 2 mW) and an EMCCD
camera (acquisition time = 5 ms). Data acquisition and image pro-
cessing were performed as described.[34]
[14] S. Howorka, Z. Siwy, Chem. Soc. Rev. 2009, 38, 2360 – 2384;
G. M. Cherf, K. R. Lieberman, H. Rashid, C. E. Lam, K.
Karplus, M. Akeson, Nat. Biotechnol. 2012, 30, 344 – 348; J.
Clarke, H. C. Wu, L. Jayasinghe, A. Patel, S. Reid, H. Bayley,
Nat. Nanotechnol. 2009, 4, 265 – 270; L. Movileanu, Trends
Biotechnol. 2009, 27, 333 – 341.
[15] N. A. Bell, C. R. Engst, M. Ablay, G. Divitini, C. Ducati, T. Liedl,
U. F. Keyser, Nano Lett. 2012, 12, 512 – 517.
[16] R. Wei, T. G. Martin, U. Rant, H. Dietz, Angew. Chem. 2012, 124,
4948 – 4951; Angew. Chem. Int. Ed. 2012, 51, 4864 – 4867.
[17] K. Bçrjesson, J. Wiberg, A. H. El-Sagheer, T. Ljungdahl, J.
Mꢂrtensson, T. Brown, B. Nordꢃn, B. Albinsson, ACS Nano
2010, 4, 5037 – 5046.
[18] J. G. Woller, K. Bçrjesson, S. Svedhem, B. Albinsson, Langmuir
2012, 28, 1944 – 1953.
[19] L.-A. Fendt, I. Bouamaied, S. Thoeni, N. Amiot, E. Stulz, J. Am.
Chem. Soc. 2007, 129, 15319 – 15329; T. Nguyen, A. Brewer, E.
Stulz, Angew. Chem. 2009, 121, 2008 – 2011; Angew. Chem. Int.
Ed. 2009, 48, 1974 – 1977.
Received: July 3, 2013
Published online: && &&, &&&&
[20] A. Brewer, G. Siligardi, C. Neylon, E. Stulz, Org. Biomol. Chem.
2011, 9, 777 – 782.
[21] J. R. Burns, S. Preus, D. G. Singleton, E. Stulz, Chem. Commun.
2012, 48, 11088 – 11090.
[22] A. Ortega, D. Amoros, J. G. de La Torre, Biophys. J. 2011, 101,
892 – 898.
[23] Y. G. Ke, J. Sharma, M. H. Liu, K. Jahn, Y. Liu, H. Yan, Nano
Lett. 2009, 9, 2445 – 2447.
[24] A. Kuzuya, M. Komiyama, Chem. Commun. 2009, 4182 – 4184.
[25] N. Mitchell, R. Schlapak, M. Kastner, D. Armitage, W. Chrza-
nowski, J. Riener, P. Hinterdorfer, A. Ebner, S. Howorka,
Angew. Chem. 2009, 121, 533 – 535; Angew. Chem. Int. Ed. 2009,
48, 525 – 527.
[26] P. Klapetek, I. Ohlꢄdal, Ultramicroscopy 2003, 94, 19 – 29.
[27] S. Hernꢅndez-Ainsa, N. A. Bell, V. V. Thacker, K. Gçpfrich, K.
Misiunas, M. E. Fuentes-Perez, F. Moreno-Herrero, U. F.
Keyser, ACS Nano 2013, 7, 6024 – 6030.
[28] S. M. Douglas, A. H. Marblestone, S. Teerapittayanon, A.
Vazquez, G. M. Church, W. M. Shih, Nucleic Acids Res. 2009,
37, 5001 – 5006.
[29] L. A. Clifton, M. R. Sanders, V. Castelletto, S. E. Rogers, R. K.
Heenan, C. Neylon, R. A. Frazier, R. J. Green, Phys. Chem.
Chem. Phys. 2011, 13, 8881 – 8888.
[30] N. Mitchell, A. Ebner, P. Hinterdorfer, R. Tampe, S. Howorka,
Small 2010, 6, 1732 – 1735.
[31] M. Kreir, C. Farre, M. Beckler, M. George, N. Fertig, Lab Chip
2008, 8, 587 – 595; O. Gassmann, M. Kreir, C. Ambrosi, J.
Pranskevich, A. Oshima, C. Roling, G. Sosinsky, N. Fertig, C.
Steinem, J. Struct. Biol. 2009, 168, 168 – 176.
[32] K. Gçpfrich, C. V. Kulkarni, O. J. Pambos, U. F. Keyser, Lang-
muir 2013, 29, 355 – 364.
[33] J. L. Gornall, K. R. Mahendran, O. J. Pambos, L. J. Steinbock, O.
Otto, C. Chimerel, M. Winterhalter, U. F. Keyser, Nano Lett.
2011, 11, 3334 – 3340.
Keywords: DNA structures · membranes · nanopores ·
porphyrins · single-molecule studies
.
[1] E. Stulz, Chem. Eur. J. 2012, 18, 4456 – 4469; H. Yang, C. K.
McLaughlin, F. A. Aldaye, G. D. Hamblin, A. Z. Rys, I. Rouiller,
H. F. Sleiman, Nat. Chem. 2009, 1, 390 – 396.
[2] B. Saccꢀ, C. M. Niemeyer, Angew. Chem. 2012, 124, 60 – 69;
Angew. Chem. Int. Ed. 2012, 51, 58 – 66.
[3] B. Ding, N. C. Seeman, Science 2006, 314, 1583 – 1585; C. Lin, H.
Yan, Nat. Nanotechnol. 2009, 4, 249 – 254; P. W. Rothemund,
Nature 2006, 440, 297 – 302; R. P. Goodman, I. A. Schaap, C. F.
Tardin, C. M. Erben, R. M. Berry, C. F. Schmidt, A. J. Turber-
field, Science 2005, 310, 1661 – 1665.
[4] S. H. Park, P. Yin, Y. Liu, J. H. Reif, T. H. LaBean, H. Yan, Nano
Lett. 2005, 5, 729 – 733; B. Saccꢀ, R. Meyer, M. Erkelenz, K.
Kiko, A. Arndt, H. Schroeder, K. S. Rabe, C. M. Niemeyer,
Angew. Chem. 2010, 122, 9568 – 9573; Angew. Chem. Int. Ed.
2010, 49, 9378 – 9383.
[5] J. Sharma, Y. G. Ke, C. X. Lin, R. Chhabra, Q. B. Wang, J.
Nangreave, Y. Liu, H. Yan, Angew. Chem. 2008, 120, 5235 – 5237;
Angew. Chem. Int. Ed. 2008, 47, 5157 – 5159.
[6] S. Helmig, A. Rotaru, D. Arian, L. Kovbasyuk, J. Arnbjerg, P. R.
Ogilby, J. Kjems, A. Mokhir, F. Besenbacher, K. V. Gothelf, ACS
Nano 2010, 4, 7475 – 7480.
[7] R. Schlapak, J. Danzberger, D. Armitage, D. Morgan, A. Ebner,
P. Hinterdorfer, P. Pollheimer, H. J. Gruber, F. Schaffler, S.
Howorka, Small 2012, 8, 89 – 97.
[8] M. Leitner, N. Mitchell, M. Kastner, R. Schlapak, H. J. Gruber,
P. Hinterdorfer, S. Howorka, A. Ebner, ACS Nano 2011, 5,
7048 – 7054.
[9] M. Langecker, V. Arnaut, T. G. Martin, J. List, S. Renner, M.
Mayer, H. Dietz, F. C. Simmel, Science 2012, 338, 932 – 936.
[10] J. Burns, E. Stulz, S. Howorka, Nano Lett. 2013, 13, 2351 – 2356.
[11] H. Bayley, L. Jayasinghe, Mol. Membr. Biol. 2004, 21, 209 – 220;
N. Sakai, S. Matile, Langmuir 2013, 29, 9031 – 9040; A. Vargas
Jentzsch, A. Hennig, J. Mareda, S. Matile, Acc. Chem. Res. 2013,
[34] V. V. Thacker, S. Ghosal, S. Hernꢅndez-Ainsa, N. A. Bell, U. F.
Keyser, Appl. Phys. Lett. 2012, 101, 223704.
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Communications
DNA Nanotechnology
Holding tight: An artificial membrane
nanopore assembled from DNA oligonu-
cleotides carries porphyrin tags (red),
which anchor the nanostructure into the
lipid bilayer. The porphyrin moieties also
act as fluorescent dyes to aid the micro-
scopic visualization of the DNA nano-
pore.
J. R. Burns, K. Gçpfrich, J. W. Wood,
V. V. Thacker, E. Stulz, U. F. Keyser,
S. Howorka*
&&&& — &&&&
Lipid-Bilayer-Spanning DNA Nanopores
with a Bifunctional Porphyrin Anchor
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ꢀ 2013 Wiley-VCH Verlag GmbH & Co. KGaA, Weinheim
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