Synthesis and self-assembly of DNA-chromophore hybrid amphiphiles

Article abstract of DOI:10.1039/c6ob00681g

DNA based spherical nanostructures are one of the promising nanostructures for several biomedical and biotechnological applications due to their excellent biocompatibility and DNA-directed surface addressability. Herein, we report the synthesis and amphiphilicity-driven self-assembly of two classes of DNA (hydrophilic)-chromophore (hydrophobic) hybrid amphiphiles into spherical nanostructures. A solid-phase "click" chemistry based modular approach is demonstrated for the synthesis of DNA-chromophore amphiphiles. Various spectroscopic and microscopic analyses reveal the self-assembly of the amphiphiles into vesicular and micellar assemblies with the corona made of hydrophilic DNA and the hydrophobic chromophoric unit as the core of the spherical nanostructures.

Full text of DOI:10.1039/c6ob00681g

View Article Online
View Journal
Organic &
Biomolecular
Chemistry
Accepted Manuscript
This article can be cited before page numbers have been issued, to do this please use: S. K. Albert, M.
Golla, H. V. P. Thelu, N. Krishnan, P. Deepak and R. Varghese, Org. Biomol. Chem., 2016, DOI:
10.1039/C6OB00681G.
This is an Accepted Manuscript, which has been through the
Royal Society of Chemistry peer review process and has been
accepted for publication.
Accepted Manuscripts are published online shortly after
acceptance, before technical editing, formatting and proof reading.
Using this free service, authors can make their results available
to the community, in citable form, before we publish the edited
article. We will replace this Accepted Manuscript with the edited
and formatted Advance Article as soon as it is available.
You can find more information about Accepted Manuscripts in the
Information for Authors.
Please note that technical editing may introduce minor changes
to the text and/or graphics, which may alter content. The journal’s
standard Terms & Conditions and the Ethical guidelines still
apply. In no event shall the Royal Society of Chemistry be held
responsible for any errors or omissions in this Accepted Manuscript
or any consequences arising from the use of any information it
contains.
www.rsc.org/obc

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 1 of 9
View Article Online
DOI: 10.1039/C6OB00681G
Journal Name
ARTICLE
Synthesis and self-assembly of DNA-chromophore hybrid
amphiphiles
Received 00th January 20xx,
Accepted 00th January 20xx
Shine K. Albert, Murali Golla, Hari Veera Prasad Thelu, Nithiyanandan Krishnan, Perapaka Deepak,
and Reji Varghese*
DOI: 10.1039/x0xx00000x
DNA based spherical nanostructures is one of the promising nanostructures for several biomedical and biotechnological
applications due to their excellent biocompatibility and DNA-directed surface addressability. Herein, we report the
synthesis and amphiphilicity-driven self-assembly of two classes of DNA (hydrophilic)-chromophore (hydrophobic) hybrid
amphiphiles into spherical nanostructures. A solid-phase “click” chemistry based modular approach is demonstrated for
the synthesis of DNA-chromophore amphiphiles. Various spectroscopic and microscopic analyses reveal the self-assembly
of the amphiphiles into vesicular and micellar assemblies with corona made of hydrophilic DNA and the hydrophobic
chromophoric unit as the core of the spherical nanostructures.
www.rsc.org/
structural scaffold for the helical organization of chromophores
either covalently or non-covalently.12 Very recently, DNA has also
been applied as the hydrophilic segment in the design of DNA based
amphiphilic systems.13 The most attractive feature of the self-
Introduction
assembly of DNA based amphiphiles in aqueous medium is the
formation of nanostructures with corona made of hydrophilic DNA.
Hence, such nanostructures potentially allow the reversible
integration of other functional molecules onto their surface through
the sequence specific DNA hybridization, which has been exploited
for many technological and biomedical applications.14-20
Design and synthesis of biocompatible chromophoric assemblies
with well-defined morphology and unique optical properties is
challenging.1 Bottom-up self-assembly using non-covalent
interactions is an efficient approach for the creation of functional
nanostructures.2 Numerous examples of chromophoric assemblies
that exploit hydrogen bonding,3 π-π stacking,4 donor-acceptor,5
host-guest,6 and metal-ligand7 interactions are reported.
Amphiphilicity-driven self-assembly is an alternative bottom-up
approach for the creation of nanostructures, and is particularly
attractive because of the inherent self-assembling tendency of
amphiphilic systems.8 Amphiphiles consist of hydrophobic and
hydrophilic moieties joined together either covalently or non-
covalently, and are capable of spontaneously assembles into
diverse nanostructures such as micelles, vesicles, sheets or tubes
mainly through hydrophobic interactions. A unique feature of
amphiphilicity-driven self-assembly is the morphology tunability of
the nanostructures, which is routinely achieved by controlling the
volume ratio between hydrophilic and hydrophobic segments. This
approach has been successfully applied in many chromophoric
systems for the crafting of nanoarchitectures with interesting
optical and electronic properties.9
The replacement of flexible polymeric segment of conventional
DNA amphiphile with a rigid π-conjugated chromophore for the
design of DNA amphiphiles has several advantages when compared
with the conventional DNA amphiphiles. These include: (i) self-
assembly propensity of the amphiphiles is greatly enhanced
through the strong π-π-stacking interaction of the chromophoric
moiety; (ii) this would also helps in effective encapsulation of the
hydrophobic guest molecule through the strong π-stacking as well
as hydrophobic interaction of the guest molecules with the
chromophoric segment; and (iii) chromophoric rigid block could
potentially serve as an optical reporter owing to the modulation of
their optical properties upon assembly/disassembly process.
However, covalent ligation of hydrophilic DNA and hydrophobic
segment is always challenging.21 The rapid emergence of Cu(I)-
catalyzed alkyne-azide cycloaddition (CuAAC) reaction, commonly
known as the “click” reaction, have shown ample opportunities in
covalent modification of DNA with other functional molecules.22
Very recently, we have reported a solid-phase “click” chemistry
approach for the synthesis of a series of DNA–rigid π-conjugated
chromophoric amphiphiles and showed their reversible self-
assembly into DNA based surface-engineered vesicles with
enhanced emission.23 Furthermore, we have demonstrated that the
DNA-directed surface addressability of the vesicles can be used for
the reversible organization of Au-NPs and other fluorophores onto
the surface of the vesicle through DNA hybridization. Herein, we
report the solid-phase “click chemistry” approach for the synthesis
The characteristic structural features of DNA such as predictable
secondary structure, nanoscale dimension, ease of synthesis and
molecular recognition properties have greatly attracted researchers
to use DNA as a building block in the design of DNA based
nanostructures. This gave birth to the emerging field of DNA
nanotechnology.10,11 Furthermore, DNA has proven to be an ideal
This journal is © The Royal Society of Chemistry 20xx
J. Name., 2013, 00, 1-3 | 1
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 2 of 9
View Article Online
DOI: 10.1039/C6OB00681G
ARTICLE
Journal Name
furnished the hydroxypyridone derivative 8. Subsequent treatment
of 8 with N,N-diphenylformamidine (DPFA) in acetic anhydride
afforded the corresponding enaminone derivative, which was
treated in-situ with the azide functionalized pyridinium salt 9 in the
presence of potassium acetate (KOAc) as base furnished the azide
modified merocyanine derivative 10. Synthesis of DNA-
chromophore amphiphiles (DNA1 and DNA2) was achieved by the
solid-phase “click” reaction between the CPG bound 5’-alkyne
hydrophilic segment
DNA
hydrophobic segment
alkyl chain
chromophore
DNAꢀchromophore amphiphile
amphiphilicityꢀdriven
selfꢀassembly
modified
DNA
(3’-ATGTCGATATGAACTTGCX-5’)
and
the
corresponding azide functionalized chromophore derivative (3 or
10) in a mixture of DMSO and t-BuOH (3:1) in the presence of
DNA shell
Cu(I)Br
as
catalyst
with tris[(1-benzyl-1H-1,2,3-triazol-4-
yl)methyl]amine (TBTA) as the ligand at 60 °C for 24 h.
Subsequently, the DNA was deprotected from the solid support
using ammonium hydroxide solution (28 %) provided the DNA-
chromophore amphiphiles DNA1 and DNA2, which was
subsequently purified and characterized through ESI-MS analysis
(Scheme 1b).
or
functional core
micelle
vesicle
DNA based surface engineered nanostructures
Optical Studies
Fig. 1 Schematic representation depicting the self-assembly of DNA-
chromophore hybrid amphiphiles into DNA based surface
engineered micelle or vesicle.
Self-assembly of the amphiphiles DNA1 and DNA2 was performed
by heating the respective amphiphile (1 µM) in Tris buffer (50 mM,
pH 7.4) at 90 °C for 5 min and then allowing to cool to room
temperature. UV-Vis absorption spectra of DNA1 and DNA2 show
the characteristic absorption of DNA and the corresponding
of two classes of DNA amphiphiles, DNA-porphyrin (DNA1) and
DNA-merocyanine (DNA2) based systems. These chromophores
were chosen in our study due to their promising optical properties
and potential applications in various fields.24 We also report the
chromophoric
segments.
Temperature-dependent
UV-Vis
absorption spectra of DNA1 and DNA2 provided better
understanding into the self-assembly of these amphiphiles.
Absorption spectrum of DNA1 shows the characteristic absorption
peaks of aggregated porphyrin at 418 nm (Soret-band) and 549 nm
(Q-band).27 Temperature dependent absorption spectrum of DNA1
shows only a slight decrease in the absorption bands at 418 nm and
549 nm with no change in the absorption maximum (Fig. 2a).
Similarly, absorption spectrum of DNA2 shows the characteristic
absorption of merocyanine and DNA at 480 nm and 260 nm,
respectively (Fig. 2b).28 In this case, temperature dependent
absorption spectrum shows a gradual red-shift of 7 nm in the
absorption maximum of merocyanine from 480 nm to 487 nm with
the increase in temperature from 20 °C to 70 °C. In accordance with
this, the temperature dependent excitation spectrum also shows a
red-shift of 7 nm with the increase in the temperature (Fig. 2b
inset). Furthermore, DNA1 and DNA2 aggregates display induced
circular dichroism (ICD) signals in the absorption regions of the
respective chromophores, indicating the transfer of molecular
chirality of DNA to the chromophoric stacks of the aggregates of
DNA1 and DNA2 (Figure 2c, d). No change is observed in the
intensity of the ICD signals of DNA1 and DNA2 aggregates with the
increase in temperature. Fluorescence spectrum of DNA1 displays
the characteristic emission of self-assembled porphyrin with
maximum centered at 645 nm (λex = 420 nm). The quantum yield of
DNA1 is 0.01. Notably, no significant change, except a slight
decrease in the emission intensity is observed in the temperature-
dependent fluorescence spectrum of DNA1 (Fig. 3a), which is in
accordance with the temperature dependent absorption spectral
studies. Emission spectrum of DNA2 shows the characteristic
emission of aggregated merocyanine at 698 nm (λex = 480 nm),
which also shows a gradual decrease in emission intensity upon
increasing the temperature from 20 °C to 80 °C (Fig. 3b). In order to
understand whether the observed photophysical changes of DNA1
and DNA2 aggregates with temperature is indeed due to the
disassembly of
amphiphilicity-driven
amphiphiles into vesicular and micellar nanostructures (Fig. 1).
self-assembly
of
DNA-chromophore
Results and discussion
Synthesis of DNA-chromophore amphiphiles
For the synthesis of DNA-chromophore amphiphiles, we adopted a
solid phase CuAAC as reported recently by us.23 For this purpose,
alkyne moiety was attached to the 5’-end of a random DNA (3’-
ATGTCGATATGAACTTGC-5’) using the commercially available C8-
alkyne-dT-CE phosphoramidite (X) following standard protocols.
The
CPG
bound
alkyne
functionalized
DNA
(3’-
ATGTCGATATGAACTTGCX-5’) thus obtained was directly used for
the solid phase “click” reaction without further purification. On the
other hand, azide modified hydrophobic chromophoric derivatives 3
and 10 were synthesized in multistep synthesis as shown in Scheme
1a. Necessary hydrophobicity to the chromophoric segments was
achieved by attaching long hydrocarbon chains (-C18H37 or -C12H25
)
to the chromophoric backbone. For the synthesis of azide
functionalized porphyrin derivative 3, bis(octadecyloxy) substituted
ester derivative of porphyrin 1 was synthesized from commercially
available starting materials.25 Lithium aluminium hydride (LiAlH4)
reduction of the ester
1 followed by the treatment of the
corresponding alcohol 2 with 1,8-Diazabicyclo[5.4.0]undec-7-ene
(DBU) and diphenylphosphoryl azide (DPPA) furnished the azide
functionalized porphyrin derivative 3. Synthesis of azide modified
merocyanine derivative 10 was started from the commercially
available N-(3,5-dihydroxyphenyl)acetamide, which was converted
into 3,5-bis(dodecyloxy)aniline (4) through multi step synthesis
following
cyanoacetic
a
reported procedure.26 Reaction between
acid (5) in the presence of
4
and
N,N'-
dicyclohexylcarbodiimide (DCC) gave the bis(dodecyloxy) amide
derivative 6, which upon condensation with ethyl acetoacetate (7)
2 | J. Name., 2012, 00, 1-3
This journal is © The Royal Society of Chemistry 20xx
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 3 of 9
View Article Online
DOI: 10.1039/C6OB00681G
Journal Name
ARTICLE
Scheme 1. (a) Synthetic scheme for azide functionalized chromophores 3 and 10. (b) Synthesis of DNA-chromophore amphiphiles
(DNA1 and DNA2) through solid-phase CuAAC reaction between CPG bound 5’-alkyne modified DNA (3’-ATGTCGATATGAACTTGCX-5’)
and azide functionalized chromophores (3 or 10).
conformation for ssDNA segment of DNAs were assumed as shown
the aggregates, as expected or the mere effect of temperature on
in Fig. 7 for the calculation of bilayer distance for DNA1 and DNA2.
photophysical properties, variable temperature optical studies were
Accordingly, the calculated bilayer distances for DNA1 and DNA2
carried out on 3 and 10. Azide derivatives 3 and 10 are the
are ∼22 nm, ∼21 nm, respectively. For the micellar assemblies the
precursor chromophores for DNA1 and DNA2, respectively. The
particle size would be equal to the bilayer packing distance,
effect of temperature on the photophysical properties of these
whereas, in the case of vesicular assemblies the particle size would
chromophores was studied by doing variable temperature optical
be significantly larger than the bilayer packing distance, because
studies of the monomeric species of 3 and 10. Experiments were
bilayer distance in the case of vesicles represents only the wall
performed in chloroform as the solvent, where the precursor
thickness of the vesicles.29 The DLS analysis of DNA1 at 20 °C shows
chromophores 3 and 10 exist as monomeric species. Interestingly,
unimodal distribution of spherical particles with size ranging from
temperature-dependent absorption and emission studies of 3 and
120 nm to 530 nm (σ = 0.296) with an average diameter of 272 nm
10 revealed similar photophysical changes as that of DNA1 and
(Fig. 4a). The diameter of the smallest particle (120 nm) is
DNA2 aggregates, respectively (Fig. S1-4). These observations
considerably larger than the calculated bilayer distance of DNA1
suggest that the observed photophysical changes for DNA1 and
(∼22 nm), clearly revealing the vesicular nature for DNA1 particles.
DNA2 aggregates with temperature could be the mere effect of
Interestingly, DLS analysis at 70 °C reveals that vesicles are not
temperature on the photophysical properties of DNA1 and DNA2
dissociating into the monomers even at high temperature (Fig. 4a).
aggregates, and not due to the temperature induced disassociation
This is in accordance with the temperature dependent optical
of the aggregates.
studies of DNA1, and confirms that the vesicles are thermally
Dynamic Light Scattering (DLS) Studies
stable, at least up to 70 °C. On the other hand, a narrow unimodal
distribution of spherical aggregates with particle size ranging from
25 nm to 95 nm (σ = 0.273) with an average diameter of 47 nm is
observed for DNA2 aggregates (Fig. 4b). In this case, it is important
to note that the smallest particle has a diameter of 25 nm, which is
approximately equal to the calculated bilayer distance of DNA2
(∼21 nm). This clearly implies that the spherical aggregates of DNA2
are micellar in nature, and not vesicular. The larger spherical
particles (>25 nm) observed for DNA2 assemblies could be the
aggregated micelles in solution. In this case also, in accordance with
the temperature dependent optical studies, DLS analysis at 70 °C
To get more insight into the morphology of the aggregated species
of DNA1 and DNA2 in solution, detailed DLS analyses were carried
out. The DLS analyses of DNA1 and DNA2 show the formation of
equilibrated nanometer sized spherical particles in solution. The
micellar or vesicular nature of the spherical particles observed in
DLS can easily be distinguished by comparing the experimental
particle size and the calculated bilayer packing distance, which is
equal to twice the molecular length of the amphiphile. It is to be
noted that in our case, interdigitation of alkyl chains and a helical
This journal is © The Royal Society of Chemistry 20xx
J. Name., 2013, 00, 1-3 | 3
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 4 of 9
View Article Online
DOI: 10.1039/C6OB00681G
ARTICLE
Journal Name
a)
a)
b)
c)
d)
b)
Fig. 2 Temperature dependent absorption spectra of (a) DNA1 and
(b) DNA2. Inset shows variable temperature excitation spectrum
(normalized) of DNA2 (monitored at λem = 698 nm). CD spectra of
(c) DNA1 and (d) DNA2 at 20 °C. All experiments were performed
with 1 μM DNA in 50 mM Tris buffer, pH 7.4, path length of the cell
= 10 mm.
Fig. 3 Normalized temperature dependent emission spectra of (a)
DNA1 (λex = 420 nm) and (b) DNA2 (λex = 480 nm). All experiments
were performed with 1 μM DNA in 50 mM Tris buffer, pH 7.4, path
length of the cell = 10 mm.
shows that micelles are thermally stable, and hence not dissociating
into the corresponding monomers with the increase in temperature
(Fig. 4b). These results clearly suggest that the amphiphiles DNA1
and DNA2 undergoes amphiphilicity-driven self-assembly in
aqueous medium into thermally stable vesicles and micelles,
respectively. Furthermore, zeta potential measurements
respectively show −4.58 mV, and −4.46 mV values for DNA1 and
DNA2 (Fig. S7), indicating the negatively charged surfaces of DNA1
vesicle and DNA2 micelle as expected.
average diameter of the particles obtained from DLS analysis. The
cross-sectional analysis of the vesicles reveals that the average
height of the vesicle is only ∼11 nm, much smaller than their
respective average diameter. The low average height of DNA1
vesicles can be attributed to the considerable flattening of the
vesicles on the mica surface, which is a characteristic feature of
a)
Dye Encapsulation and Microscopic Studies
Better insights into the morphology of these aggregates were
provided by the dye encapsulation and microscopic studies. Clear
evidence for the vesicular nature for the spherical aggregates of
DNA1 was obtained from the encapsulation studies with Calcein.
Calcein is a fluorescent hydrophilic dye, and hence would be
encapsulated in the hydrophilic interior of the vesicle.30
A
significant quenching is observed for the emission intensity of
Calcein encapsulated inside the DNA1 vesicle when compared with
the emission of an absorption matched solution of Calcein in
vesicle-free solution (λex = 470 nm, Fig. 5a). This can be attributed
to the self-quenching of the dye emission due to the high
encapsulation of Calcein in the hydrophilic cavity of DNA1 vesicle.
Laser Scanning Confocal Microscope (LSCM), Atomic Force
Microscope (AFM), and Transmission Electron Microscope (TEM)
were also used to study the self-assembled nanostructures of DNA1
and DNA2 amphiphiles. In accordance with the fluorescence
properties of DNA1, LSCM images show the formation of
fluorescent vesicles for DNA1 (λex = 405 nm, Fig. 5b). The height
images of self-assemblies of DNA1 obtained by tapping-mode AFM
show the formation of nanometer sized spherical assemblies (Fig.
6a). The average diameter of the spheres of DNA1 assemblies,
which were estimated from the fitted histograms of the size
distribution curves after subtracting the tip-broadening factor, is
∼250 nm. This is in good agreement with the
b)
Fig. 4 Size distribution graphs from the DLS measurements of (a)
DNA1, and (b) DNA2 at 20 °C (solid line) and 70 °C (dotted line). All
experiments were performed with 1 μM DNA in 50 mM Tris buffer,
pH 7.4.
4 | J. Name., 2012, 00, 1-3
This journal is © The Royal Society of Chemistry 20xx
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 5 of 9
View Article Online
DOI: 10.1039/C6OB00681G
Journal Name
ARTICLE
a)
b)
a)
~22 nm
O
O
N
N
N
DNA1
O
N
N
N
O
N
N
N
Zn
Zn
N
N
N
Zn
Zn
N
O
N
N
O
O
N
O
~20 nm
100 nm
0.5 ꢁm
c)
d)
~21 nm
O
O
CN
CN
H3
H3
C
O
O
O
O
C
N
O
O
N
O
N
N
N
N
O
O
O
N
O
N
O
O
b)
O
CH3
CH3
DNA2
CN
CN
~20 nm
20 nm
100 nm
Fig. 7 Zoom-out TEM images of (a) DNA1 and (c) DNA2, and the
corresponding zoom-in TEM images of (b) DNA1 and (d) DNA2 on a
carbon coated grid. Schematic representations of the proposed
bilayer packing of the amphiphiles are also shown.
10 ꢁm
“soft” vesicles.31 On the other hand, AFM analysis of DNA2
assemblies shows the formation of spherical particles with size
ranging from ∼20 nm to ∼90 nm (Fig. 6b). The average height of the
particle is ∼7 nm as revealed from the cross-sectional analysis. The
micellar nature of DNA2 assemblies is clear from the approximate
matching of the diameter of the smallest particle (∼20 nm) with the
calculated bilayer packing distance for DNA2 (∼21 nm). The
morphology of the spherical aggregates of DNA1 and DNA2
deduced from various spectroscopic and microscopic studies were
unequivocally confirmed by the TEM analyses of the
nanostructures. For DNA1, spherical particles of diameters in the
range of 100–400 nm with clear contrast difference between the
periphery and the inner part of the spheres are observed in the
TEM analysis (Fig. 7a, b), which is a conclusive evidence for the
vesicular nature for the spheres of DNA1. More importantly, the
wall thickness of the vesicle calculated from the TEM images (∼20
nm) is approximately equal to the calculated wall thickness (∼22
nm). However, 2 nm decrease in the experimentally observed wall
thickness compared with the calculated value can be attributed to
the flexible nature of the alkyl chains, the linker, and the ssDNA
segments of DNA1. On the other hand, TEM images of DNA2
aggregates show the formation of spherical particles with almost
uniform size of ∼20 nm, which is approximately equal to the bilayer
packing distance for DNA2 (∼21 nm, Fig. 7c, d). This confirms that
the spherical aggregates of DNA2 are micellar in nature as revealed
by other spectroscopic and microscopic techniques. It is worth
noting that no change is observed in vesicular morphology of DNA1
and micellar morphology of DNA2 even after hybridization of ssDNA
on their surface with the corresponding complementary DNA strand
(5’-TACAGCTATACTTGAACG-3’) (Fig. S8). We have also carried out
detailed morphological studies of the aggregates of 3 and 10, which
were obtained by the addition of water into the THF solution of 3
and 10, in order to compare their self-assembling properties with
the corresponding DNA based amphiphiles. Interestingly, fibrous
morphology was observed for the aggregates of 3 and 10 from the
respective AFM analyses (Fig. S9 and S10). Optical and
Fig. 5 (a) Fluorescence spectra of Calcein encapsulated in DNA1
vesicle and Calcein in vesicle-free solution (λex = 470 nm). (b) LSCM
image of DNA1 vesicle (λex = 405 nm).
a)
1ꢁm
b)
100 nm
Fig. 6 AFM height images of (a) DNA1 (z-scale = 25 nm) and (b)
DNA2 (z-scale = 50 nm) with the corresponding the particle size
distributions. A representative cross-section analysis of a single
particle is also shown. All experiments were performed on a freshly
cleaved mica surface.
This journal is © The Royal Society of Chemistry 20xx
J. Name., 2013, 00, 1-3 | 5
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 6 of 9
View Article Online
DOI: 10.1039/C6OB00681G
ARTICLE
Journal Name
morphological studies clearly reveal that aggregates of 3 and 10 and Samples were allowed to adsorb on grid for 2 min and excess
the corresponding amphiphiles (DNA1 and DNA2) exhibit dissimilar sample was wicked with filter paper. Grid was washed twice by
optical and morphological properties, and hence it can be touching grid with a drop of water and then removing excess water
concluded that DNA induced amphiphilicity-driven self-assembly using filter paper and was stained with 0.7% uranyl formate
leads to unique functional nanostructures.
solution. Absorption spectra were recorded on a peltier attached
Shimadzu UV-3600 Vis-NIR Spectrophotometer in a quartz cuvette
of 10 mm path length. Steady state fluorescence and excitation
spectra were recorded on
a Horiba Jobin Vyon Fluorimeter
Conclusions
equipped with peltier cell holder. Temperature dependent emission
experiments were carried out by heating samples from 20 °C to 70
°C at an interval of 5 °C equilibrating samples for 5 minutes at each
temperature before recording the spectra and were given
correction for solvents. DLS analyses were done on Malvern
Zetasizer Nano Zs equipped with 655 nm laser. Experiments were
performed at 25 °C at a back scattering angle of 173°. HR-MS
analysis was done on thermo extractive orbitrap mass
spectrometry. ESI-MS was recorded on Waters Xevo G2 QTof and
GC-MS on Shimadzu GCMS-QP2010. Confocal microscopic analysis
was done on inverted Leica SP5-DMRX Laser Confocal Microscope.
Sample was prepared by drop casting the solution on a cleaned
glass slide and was dried under vacuum. ProLong Gold (antifade
reagent) was used to mount the cover slip and imaging was done in
63X magnification in presence of immersion oil.
In summary, we have reported a solid-phase “click” chemistry
based modular approach for the synthesis of DNA (hydrophilic)-
chromophore (hydrophobic) hybrid amphiphiles, and demonstrated
their amphiphilicity driven self-assembly into DNA based surface
engineered spherical nanostructures. The generality of the
synthetic approach was demonstrated through the synthesis of two
different classes of DNA-chromophore amphiphiles. Temperature-
dependent optical and light scattering analyses have revealed that
the amphiphiles are self-assembling into thermally stable spherical
nanostructures. Detailed microscopic analyses have shown that the
DNA1 self-assemble into vesicular nanostructures, whereas DNA2
self-assemble into micellar nanostructures. One of the most striking
features of this class of nanostructures is the dense display of DNA
their surface, which makes this kind of nanostructures
biocompatible and surface addressable, and hence would be an
ideal candidate for diverse biomedical and biotechnological
applications. We hope the “click” chemistry based modular
synthesis and unique structural features of this kind nanostructures
may encourage researchers to design other classes of DNA-
chromophores hybrid amphiphiles for addressing the challenges in
drug delivery and nanoelectronics.
Synthesis of 2: To a solution of 1 (0.100 g, 0.0892 mmol) in freshly
distilled THF (2 mL), LiAlH4 (2M, 6 mL) was added and stirred at 0 °C
for 1 h. Then the temperature was slowly increased to room
temperature and was stirred 4 h. After completion of the reaction,
reaction mixture was poured into ice-cold water and was extracted
with ethyl acetate. Organic layer was dried over anhydrous Na2SO4
and the solvent was removed. The crude product was purified using
column chromatography using petroleum ether:ethyl acetate
(80:20) as eluent to get desired product as dark purple solid (90%).
1
M.P. 76 °C; TLC (petroleum ether:ethyl acetate, 80:20) Rf = 0.27; H
Experimental
NMR (500 MHz, CDCl3), δ (ppm) = 0.78 (t, J = 7.1 Hz, 6H), 1.14–1.20
(m, 52H), 1.25–1.29 (m, 4H), 1.40–1.44 (m, 4H), 1.50–1.80 (m, 4H),
1.77–1.80 (m, 4H), 4.06 (t, J = 5 Hz, 4H), 4.74 (s, 2H), 6.83 (t, J = 2.3
Hz, 1H), 7.35 (d, J = 2.2 Hz, 2H), 7.55 (d, J = 7.7 Hz, 2H), 8.12 (d, J =
7.7 Hz, 2H), 9.01 (d, J = 4.35 Hz, 2H), 9.18 (d, J = 4.45 Hz, 2H), 9.32–
9.34 (m, 4H), 10.20 (s, 2H); 13C NMR (125 MHz, CDCl3), δ (ppm) =
14.11, 22.68, 26.14, 29.35, 29.43, 29.46, 29.61, 29.64, 29.68, 31.92,
65.25, 68.44, 100.97, 106.18, 114.55, 119.23, 120.30, 125.19,
131.69, 131.72, 132.36, 132.66, 133.71, 134.77, 142.06, 144.36,
149.44, 149.59, 149.99, 150.03, 158.35. HR-MS (m/z); [M+H]+ calcd.
for [C69H95N4O3Zn]+: 1091.6617; found: 1091.6668.
Compounds 1,25 and 426 were obtained through multistep synthesis
as per the reported procedures. All chemicals used for the organic
synthesis were purchased from Sigma Aldrich and were used as
received. Solvents were dried using the standard procedures
wherever required. Column chromatography was done on 200-400
mesh silica gel. TLC analyses were performed on aluminium plates
coated with silica gel 60 F254. Melting points were recorded on
SMP30 melting point apparatus and are uncorrected. 1H and 13C
spectra were recorded on Bruker Avance-III 500 MHz NMR
Synthesis of 3: To a solution of 2 (0.057 g, 0.05 mmol) in freshly
dried toluene (2 mL), 1,8-diazabicyclo[5.4.0]undec-7-ene (DBU) (10
µL, 0.07 mmol) and diphenylphosphoryl azide (DPPA) (13.5 µL, 0.06
mmol) were added and the reaction mixture was stirred at room
temperature for 24 h. After completion of the reaction saturated
NH4Cl was added and was extracted with ethyl acetate. Organic
layer was dried over anhydrous Na2SO4 and solvent was removed.
Crude product was further purified using column chromatography
using petroleum ether:ethyl acetate (80:20) as eluent to get the
desired product as dark purple solid (90%). M.P. 70 °C; TLC
(petroleum ether:ethyl acetate) Rf = 0.6; IR (KBr): 2927, 2854, 2100,
1597, 1259, 1097, 920, 763 cm-1. 1H NMR (500 MHz, CDCl3), δ (ppm)
= 0.77 (t, J = 7.0 Hz, 6H), 1.14–1.21 (m, 52H), 1.28–1.32 (m, 6H),
1.38–1.40 (m, 4H), 1.42–1.43 (m, 4H), 1.79–1.83 (m, 4H), 4.08 (t, J =
2.4 Hz, 4H), 4.61 (s, 2H), 6.84 (t, J = 2.0 Hz, 1H), 7.35 (d, J = 2.5 Hz,
2H), 7.44 (d, J = 6.4 Hz, 2H), 8.16 (d, J = 8.0 Hz, 2H), 8.99 (d, J = 4.50
Hz, 2H), 9.18 (d, J = 4.50 Hz, 2H), 9.36 (t, J = 4.0 Hz, 4H), 10.24 (s,
2H); 13C NMR (125 MHz, CDCl3), δ (ppm) = 14.06, 22.65, 26.13,
Spectrometer.
Deuterated
solvents
containing
1,1,1,1-
tetramethylsilane as internal standard was used for recording NMR.
Shimadzu IR prestige-21 FT-IR was used for recording IR spectra
where solid samples were pelletized along with KBr. Water used for
all the experiments was de-ionised Milli Q (18.2 MΩ.cm). All
phosphoramidites for the oligonucleotide synthesis were purchased
from Glen Research and were used as received. Alkyne modified
oligonucleotides were synthesized on H-8 K&A DNA/RNA
synthesizer in 1 μmole scale. AFM imaging was done on Multimode
SPM (Veeco Nanoscope V). Samples were prepared by depositing 2
μL of aggregated samples on freshly cleaved mica surface and were
dried under air. Images were recorded under ambient conditions in
tapping mode. Probe used for imaging was antimony doped silicon
cantilever with a resonant frequency of 300 kHz and spring constant
of 40 Nm-1. TEM analyses were carried out on FEI Tecnai 30 G2 (300
kV) High Resolution-TEM. Sample preparation was done by drop-
casting 2 μL of sample on negatively glow discharged (PELCO
easiGlow) 400-mesh carbon coated copper grid (Ted Pella, Inc.).
6 | J. Name., 2012, 00, 1-3
This journal is © The Royal Society of Chemistry 20xx
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 7 of 9
View Article Online
DOI: 10.1039/C6OB00681G
Journal Name
ARTICLE
29.31, 29.45, 29.59, 29.60, 29.64, 31.89, 54.85, 68.44, 101.01, the solution of Cu(1)Br (100 μL) and TBTA (50 μL) in a 5 mL two-
106.25, 114.55, 119.19, 120.14, 126.43, 131.75, 131.79, 132.25, neck glass flask. The mixture was degassed by three cycles of
132.71, 134.58, 134.93, 142.76, 144.27, 149.47, 149.64, 149.88, freeze-pump-thaw method. CPG bound alkyne modified DNA (1
150.07, 158.37. HR-MS (m/z); [M+H]+ calcd. for [C67H94N7O2Zn]+: μmol) was added to the degassed solution under argon purging and
1116.6682; found: 1116.6728.
stirred at 60 °C for 24 h. Afterward, the beads were washed
Synthesis of 6: A solution of 4 (5 g, 10.84 mmol) and cyanoacetic repeatedly with THF, DCM and finally with acetonitrile for several
acid (1.19 g, 14.09 mmol) in freshly dried and distilled THF was times (minimum 10 times) and then dried under vacuum. Beads
stirred at 70 °C for 1 h and DCC (2.90 g, 14.09 mmol) was added to were transferred into a 2 mL Eppendorf vials, 1 mL of 28 %
the hot reaction mixture. It was then cooled to room temperature, ammonia solution was added and vortexed for 24 h at room
solvent was removed and crude product was purified by column temperature. Beads were removed by filtering using centrifugal
chromatography using dichloromethane as eluent to get desired filters (0.45 μm filter size, Millipore Ultrafree MC). After removing
product as white solid (75%). M.P. 62.0 °C; TLC (dichloromethane) Rf the ammonia using speed vacuum, sodium acetate solution (0.3 M,
= 0.37; 1H NMR (500 MHz, CDCl3) δ (ppm) = 0.80 (t, J = 7 Hz, 6H), 100 μL) was added to this and was again vortexed at room
1.19–1.25 (m, 34H), 1.32–1.36 (m, 4H), 1.60–1.70 (m, 4H), 3.44 (s, temperature for another 1 h. Solution was then filtered using
2H), 3.83 (t, J = 6.5 Hz, 4H), 6.20 (s, 1H), 6.60 (d, J = 1.9, 2H), 7.55 (s, centrifugal filter (0.45 μm filter size, Millipore Ultrafree MC).
1H); 13C NMR (125 MHz, CDCl3) δ (ppm) = 14.09, 22.67, 24.65, Unconjugated DNAs were then removed through Amicon
25.20, 26.01, 29.33, 29.60, 30.83, 31.90, 32.35, 40.84, 50.61, 68.19, centrifugal filters. Purification was done by centrifuging the DNA
98.37, 99.05, 114.66, 138.58, 152.76, 159.48, 160.57. HR-MS (m/z); solution 5 times at 8800 rpm for 3 minutes using 3K filters (Milipore
[M+H]+ calcd. for [C33H57N2O3]+: 529.4291; found: 529.4359.
UFC50003BK) followed by centrifuging 20 times using 30K filters
Synthesis of 8: To a solution of 6 (1.0 g, 1.89 mmol) and (Milipore UFC5030BK) at 5000 rpm for 3 minutes. Purity of the
ethylacetocetate (0.254 g, 1.89 mmol) in acetonitrile (5 mL), DNAs was confirmed by gel electrophoresis analysis (20%
piperidine (0.46 mL) was added and heated at 90 °C for 20 h. After denaturing PAGE), and were characterized through ESI-MS analyses.
the reaction, solvent was removed and crude product was filtered Mass calcd. for DNA1: 7002.61; found 7001.98; mass calcd. for
on silica neutralized with triethylamine using dichloromethane as DNA2: 6681.48; found 6682.37.
eluent to get desired product as dark brown semi solid (43%). TLC Calcein encapsulation studies: Stock solution of Calcein was
(dichloromethane) Rf = 0.31; 1H NMR (500 MHz, CDCl3) δ (ppm) = prepared in methanol. Stock solution was taken in a vial and solvent
0.80 (t, J = 7 Hz, 6H), 1.06–1.32 (m, 34H), 1.62–1.66 (m, 4H), 2.13 (s, was evaporated. Solution of DNA1 (1 μM) in water was added into
3H), 3.80 (s, 4H), 5.50 (s, 1H), 6.22(s, 2H), 6.33 (s, 1H), 10.48 (s, 1H); the Calcein solution so that final concentration of Calcein was 0.5
13C NMR (125 MHz, DMSO-d6) δ (ppm) = 8.66, 13.89, 20.38, 22.05, μM. Solution was sonicated for 1 minute and was then heated to 90
25.48, 28.61, 28.66, 28.69, 28.94, 28.97, 28.99, 31.26, 45.79, 54.82, ⁰C and was maintained at 90 ⁰C for 5 minutes on a thermal shaker.
67.48, 75.58, 98.83, 99.60, 108.12, 121.56, 140.45, 151.98, 159.45, It was then switched off and solution was allowed to cool to room
163.76, 164.25. HR-MS (m/z); [M+H]+ calcd. for [C37H59N2O4]+: temperature. Unencapsulated Calcein was removed by using 3 KDa
595.4397; found: 595.4818.
molecular weight cut-off centrifugal filters from Amicon. Filtration
Synthesis of 10: A solution of 8 (0.4g, 0.67 mmol) and N,N’- was done by centrifuging the sample at 5800 rpm for 3 minutes.
diphenylformamidine (0.67 mmol) in acetic anhydride (0.67 mL) Centrifuging was repeated for nearly 20 times and afterwards
emission was recorded by exciting the sample at 470 nm. Control
experiment was done by dissolving Calcein in water in such a way
that concentration of Calcein in control as well as experimental
solution was same.
was stirred (∼15 min) at room temperature till the reaction mixture
become solid and was then stirred at 90 °C for 15 min and cooled to
room temperature. To this reaction mixture 9 (0.15 g, 0.67 mmol)
and potassium acetate (0.06 g, 0.67 mmol) was added and stirred at
90 °C for 2 h. After reaction acetic anhydride was removed under
reduced pressure and the product was purified by column
chromatography using dichloromethane:methanol as eluent (98:2)
to get desired product as dark brown solid (67%). M.P. 174.0 °C; TLC
(dichloromethane) Rf = 0.37; IR (KBr): 3057, 2922, 2852, 2193, 2094,
Acknowledgements
We thank the financial support from DBT (No.
BT/PR7030/NNT/28/636/2012). R.V. is grateful to the DST for a
Ramanujan Fellowship. S.K.A. thanks CSIR, and H.V.P.T. and
N.K. acknowledge UGC for research fellowships. We are
thankful to University of Madras for TEM experiments. We
acknowledge CSIR-NIIST for HR-MS and M.G. University for ESI-
MS analyses.
1
1658, 1593, 1564, 1490, 1390, 1282, 1161, 833, 744 cm-1. H NMR
(500 MHz, CD2Cl2), δ (ppm) = 0.81 (t, J = 7 Hz, 6H), 1.18–1.45 (m,
43H), 1.50–1.52 (m, 2H), 1.64–1.68 (m, 4H), 1.85–1.90 (m, 2H), 2.44
(s, 3H), 3.31 (t, J = 6.5 Hz, 2H), 3.83 (t, J = 6.5 Hz, 4H), 4.01 (t, J = 4
Hz, 2H), 6.19 (s, 2H), 6.37 (t, J = 2 Hz, 1H), 7.30 (d, J = 6.5 Hz, 2H),
7.58 (d, J = 7.5 Hz, 2H), 7.71 (d, J = 2 Hz, 2H); 13C NMR (125 MHz,
CD2Cl2) δ (ppm) = 13.89, 18.70, 22.69, 25.48, 26.04, 28.34, 29.27,
29.35, 29.43, 29.62, 29.64, 29.65, 29.68, 31.92, 50.74, 58.72, 68.29,
87.91, 100.59, 106.33, 107.55, 113.80, 119.32, 120.51, 139.35,
139.53, 140.60, 156.86, 156.99, 160.58, 163.20, 163.53. HR-MS
(m/z); [M+H]+ calcd. for [C48H71N6O4]+: 795.5459; found: 795.5530.
General procedure for the synthesis of DNA1 and DNA2: Azide
modified chromophore derivative (3 or 10, 20 μmol) was dissolved
in 200 μL of dry THF in a glass vial. Cu(1)Br (1.5 mg, 10 μmol) was
dissolved in 100 μL of freshly prepared DMSO:t-butanol (3:1)
mixture. Tris[(1-benzyl-1H-1,2,3-triazol-4-yl)methyl]amine (TBTA,
5.3 mg, 10 μmol) was dissolved in 50 μL of DMSO:t-butanol (3:1)
mixture. Subsequently, solution of 3 or 10 (200 μL) was added to
Notes and references
1
(a) J.-M. Lehn, Science, 2002, 295, 2400–2403; (b) S. I. Stupp,
Nano Lett., 2010, 10, 4783–4786; (c) T. Aida, E. W. Meijer
and S. I. Stupp, Science, 2012, 335, 813–817.
2
(a) L. J. Prins, D. N. Reinhoudt and P. Timmerman, Angew.
Chem., Int. Ed., 2001, 40, 2382–2426; (b) L. Brunsveld, B. J. B.
Folmer, E. W. Meijer and R. P. Sijbesma, Chem. Rev., 2001,
101, 4071–4097; (c) F. Würthner, Chem. Commun., 2004,
1564–1579; (d) A. C. Grimsdale and K. Müllen, Angew.
This journal is © The Royal Society of Chemistry 20xx
J. Name., 2013, 00, 1-3 | 7
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 8 of 9
View Article Online
DOI: 10.1039/C6OB00681G
ARTICLE
Journal Name
Chem., Int. Ed., 2005, 44, 5592–5629; (e) F. J. M. Hoeben, P.
Jonkheijm, E. W. Meijer and A. P. H. J. Schenning, Chem.
Rev., 2005, 105, 1491–1546; (f) S. S. Babu, V. K. Praveen and 13 (a) M. P. Thompson, M.-P. Chien, T.-H. Ku, A. M. Rush and N.
9040–9046; (h) V. L. Malinovskii, D. Wenger and R. Häner,
Chem. Soc. Rev., 2010, 39, 410–422.
A. Ajayaghosh, Chem. Rev., 2014, 114, 1973–2129.
C. Gianneschi, Nano Lett., 2010, 10, 2690–2693; (b) K. M. M.
Carneiro, F. A. Aldaye and H. F. Sleiman, J. Am. Chem. Soc.,
2010, 132, 679–685; (c) M. Kwak and A. Herrmann, Chem.
Soc. Rev., 2011, 40, 5745–5755; (d) Z. Zhao, L.Wang, Y. Liu, Z.
Yang, Y.-M. He, Z. Li, Q.-H. Fan and D. Liu, Chem. Commun.,
2012, 48, 9753–9755; (e) X. Xiong, H. Liu, Z. Zhao, M. B.
Altman, D. Lopez-Colon, C. J. Yang, L.-J. Chang, C. Liu and W.
Tan, Angew. Chem., Int. Ed., 2013, 52, 1472–1476; (f) M.
3
(a) C. Thalacker and F. Würthner, Adv. Funct. Mater., 2002,
12, 209–218; (b) K. Sugiyasu, N. Fujita and S. Shinkai, Angew.
Chem., Int. Ed., 2004, 43, 1229–1233; (c) A. Ajayaghosh, R.
Varghese, S. J. George and C. Vijayakumar, Angew. Chem.,
Int. Ed., 2006, 45, 1141–1144; (d) E.-K. Schillinger, E. Mena-
Osteritz, J. Hentschel, H. G. Börner and P. Bäuerle, Adv.
Mater., 2009, 21, 1562–1567; (e) S. Yagai, Y. Goto, X. Lin, T.
Karatsu, A. Kitamura, D. Kuzuhara, H. Yamada, Y. Kikkawa, A.
Saeki and S. Seki, Angew. Chem., Int. Ed., 2012, 51, 6643–
6647.
Humenik, M. Drechsler and T. Scheibel, Nano Lett., 2014, 14
,
3999–4004; C. Zhou, D. Wang, Y. Dong, L. Xin, Y. Sun, Z. Yang
and D. Liu, Small, 2015, 11, 1161–1164 (h) Y. Vyborna, M.
Vybornyi, A. V. Rudnev and R. Häner, Angew. Chem., Int. Ed.,
2015, 54, 7934–7938.
4
(a) A. Ajayaghosh, R. Varghese, V. K. Praveen and S. Mahesh,
Angew. Chem., Int. Ed., 2006, 45, 3261–3264; (b) Y. Tang, Z.
Zhou, K. Ogawa, G. P. Lopez, K. S. Schanze and D. G. Whitten, 14 L. Wang, Y. Feng, Y. Sun, Z. Li, Z. Yang, Y.-M. He, Q.-H. Fan
Langmuir, 2009, 25, 21–25; (c) C. Kulkarni and S. J. George, and D. Liu, Soft Matter, 2011, , 7187–7190.
Chem. Eur. J., 2014, 20, 4537–4541; (d) C. Rest, M. J. 15 F. E. Alemdaroglu, K. Ding, R. Berger and A. Herrmann,
Mayoral, K. Fucke, J. Schellheimer, V. Stepanenko and G. Angew. Chem., Int. Ed., 2006, 45, 4206–4210.
Fernández, Angew. Chem., Int. Ed., 2014, 53, 700–705; (e) R. 16 M.-P. Chien and N. C. Gianneschi, Small, 2011,
T. Cheriya, A. R. Mallia and M. Hariharan, Energy Environ. 17 M. Kwak, J. Gao, D. K. Prusty, A. J. Musser, V. A. Markov, N.
7
7, 2041–2046.
Sci., 2014,
(a) J. J. Reczek, K. R. Villazor, V. Lynch, T. M. Swager and B. L.
Iverson, J. Am. Chem. Soc., 2006, 128, 7995–8002; (b) K. Liu,
C. Wang, Z. Li and X. Zhang, Angew. Chem., Int. Ed., 2011, 50,
4952–4956; (c) K. Jalani, M. Kumar and S. J. George, Chem.
Commun., 2013, 49, 5174–5176.
(a) A. Harada, Y. Takashima and H. Yamaguchi, Chem. Soc.
Rev., 2009, 38, 875–882; (b) D. Jiao, J. Geng, X. J. Loh, D. Das, 20 F. E. Alemdaroglu, N. C. Alemdaroglu, P. Langguth and A.
T.-C. Lee and O. A. Scherman, Angew. Chem., Int. Ed., 2012,
51, 9633–9637.
7
, 1661–1669.
Tombros, M. C. A. Stuart, W. R. Browne, E. J. Boekema, G.
Brinke, H. T. Jonkman, B. J. van Wees, M. A. Loi and A.
Herrmann, Angew. Chem., Int. Ed., 2011, 50, 3206– 3210.
18 M.-P. Chien, M. P. Thompson and N. C. Gianneschi, Chem.
Commun., 2011, 47, 167–169.
5
19 T. G. W. Edwardson, K. M. M. Carneiro, C. K. McLaughlin, C. J.
Serpell and H. F. Sleiman, Nat. Chem., 2013, 5, 868–875.
6
7
Herrmann, Adv. Mater., 2008, 20, 899–902.
21 (a) S. E. Averick, S. K. Dey, D. Grahacharya, K. Matyjaszewski
and S. R. Das, Angew. Chem., Int. Ed., 2014, 53, 2739–2744;
(b) T. G. W. Edwardson, K. M. M. Carneiro, C. J. Serpell and H.
F. Sleiman, Angew. Chem., Int. Ed., 2014, 53, 4567–4571.
(a) C. Tan, M. R. Pinto and K. S. Schanze, Chem. Commun.,
2002, 446–447; (b) V. Stepanenko, M. Stocker, P. Müller, M.
Büchner and F. Würthner, J. Mater. Chem., 2009, 19, 6816–
6826; (c) G. R. Whittell, M. D. Hager, U. S. Schubert and I. 22 (a) J. Gierlich, G. A. Burley, P. M. E. Gramlich, D. M.
Manners, Nat. Mater., 2011, 10, 176–188.
Hammond and T. Carell, Org. Lett., 2006, 8, 3639–3642; (b)
8
9
(a) Y.-b. Lim, K.-S. Moon and M. Lee, Chem. Soc. Rev., 2009,
38, 925–934; (b) C. Wang, Z. Wang and X. Zhang, Acc. Chem.
Res., 2012, 45, 608–618.
G. Godeau, C. Staedel, P. Barthélémy, J. Med. Chem., 2008,
51, 4374–4376; (c) A. H. El-Sagheera and T. Brown, Chem.
Soc. Rev., 2010, 39, 1388–1405; (d) C. Beyer and H.-A.
Wagenknecht, Chem. Commun., 2010, 46, 2230–2231; (e) S.
Pal, R. Varghese, Z. Deng, Z. Zhao, A. Kumar, H. Yan and Y.
Liu, Angew. Chem., Int. Ed., 2011, 50, 4176–4179; (f) S. E.
Averick, E. Paredes, S. K. Dey, K. M. Snyder, N. Tapinos, K.
(a) J. P. Hill, W. Jin, A. Kosaka, T. Fukushima, H. Ichihara, T.
Shimomura, K. Ito, T. Hashizume, N. Ishii and T. Aida, Science,
2004, 304, 1481–1483; (b) J. F. Hulvat, M. Sofos, K. Tajima
and S. I. Stupp, J. Am. Chem. Soc., 2005, 127, 366–372; (c) B.
Narayan, S. P. Senanayak, A. Jain, K. S. Narayan and S. J.
George, Adv. Funct. Mater., 2013, 23, 3053–3060; (d) S.
Yagai, S. Okamura, Y. Nakano, M. Yamauchi, K. Kishikawa, T.
Karatsu, A. Kitamura, A. Ueno, D. Kuzuhara, H. Yamada, T.
Matyjaszewski and S. R. Das, J. Am. Chem. Soc., 2013, 135
,
12508−12511; (g) A. Aimé, N. Beztsinna, A. Patwa, A.
Pokolenko, I. Bestel, P. Barthélémy, Bioconjugate Chem.,
2013, 24, 1345–1355; (h) M. Humenik and T. Scheibel, ACS
Seki and H. Ito, Nat. Commun., 2014,
Martin, V. Stepanenko, N. K. Allampally, D. Schmidt and G.
Fernández, Chem. Commun., 2014, 50, 13366–13369.
10 N. C. Seeman, J. Theor. Biol., 1982, 99, 237–247.
11 For reviews, see: (a) K. V. Gothelf and T. H. LaBean, Org.
Biomol. Chem., 2005, 3, 4023–4037; (b) N. C. Seeman, Annu.
5
, 1–10; (e) C. Rest, A.
Nano, 2014, 8, 1342–1349; (i) R. V. Thaner, I. Eryazici, O. K.
Farha, C. A. Mirkin and S. T. Nguyen, Chem. Sci., 2014, 5,
1091–1096.
23 S. K. Albert, H. V. P. Thelu, M. Golla, N. Krishnan, S.
Chaudhary and R. Varghese, Angew. Chem., Int. Ed., 2014,
53, 8352–8357.
Rev. Biochem., 2010, 79, 65–87; (c) F. A. Aldaye, A. L. Palmer 24 (a) A. C. Grimsdale and K. Müllen, Angew. Chem., Int. Ed.,
2005, 44, 5592–5629; (b) Z. Chen, A. Lohr, C. R. Saha-Möller
and F. Würthner, Chem. Soc. Rev., 2009, 38, 564–584.
and H. F. Sleiman, Science, 2008, 321, 1795–1799; (d) A. V.
Pinheiro, D. Han, W. M. Shih and H. Yan, Nat. Nanotech.,
2011,
Yan and K. V. Gothelf, Chem. Soc. Rev., 2011, 40, 5636–5646.
12 (a) J. Gao, C. Strässler, D. Tahmassebi and E. T. Kool, J. Am. 26 F. Würthner, S. Yao, T. Debaerdemaeker and R. Wortmann, J.
Chem. Soc., 2002, 124, 11590–11591; (b) H. Kashida, H. Am. Chem. Soc., 2002, 124, 9431–9447.
Asanuma and M. Komiyama, Angew. Chem., Int. Ed., 2004, 27 L.-A. Fendt, I. Bouamaied, S. Thöni, N. Amiot and E. Stulz, J.
43, 6522–6525; (c) E. Mayer-Enthart and H.-A. Wagenknecht, Am. Chem. Soc., 2007, 129, 15319–15329.
Angew. Chem., Int. Ed., 2006, 45, 3372–3375; (d) V. L. 28 U. Rösch, S. Yao, R. Wortmann and F. Würthner, Angew.
Malinovskii, F. Samain and R. Häner, Angew. Chem., Int. Ed., Chem., Int. Ed., 2006, 45, 7026–7030.
2007, 46 4464–4467; (f) R. Varghese and H.-A. 29 B.-S. Kim, D.-J. Hong, J. Bae and M. Lee, J. Am. Chem. Soc.,
Wagenknecht, Chem. Commun., 2009, 2615–2624; (g) R. 2005, 127, 16333–16337.
Varghese and H.-A. Wagenknecht, Chem. Eur. J., 2010, 16
6, 763–772; (e) T. Tørring, N. V. Voigt, J. Nangreave, H. 25 D. Kim, J. Heo, S. Ham, H. Yoo, C.-H. Lee, H. Yoon, D. Ryu, D.
Kim and W.-D. Jang, Chem. Commun., 2011, 47, 2405–2407.
,
,
8 | J. Name., 2012, 00, 1-3
This journal is © The Royal Society of Chemistry 20xx
Please do not adjust margins

Please do not adjust margins
Organic & Biomolecular Chemistry
Page 9 of 9
View Article Online
DOI: 10.1039/C6OB00681G
Journal Name
ARTICLE
30 E. N. Savariar, S. V. Aathimanikandan, and S. Thayumanavan,
J. Am. Chem. Soc., 2006, 128, 16224–16230.
31 M. Yang, W. Wang, F. Yuan, X. Zhang, J. Li, F. Liang, B. He, B.
Minch and G. Wegner, J. Am. Chem. Soc., 2005, 127, 15107–
15111.
This journal is © The Royal Society of Chemistry 20xx
J. Name., 2013, 00, 1-3 | 9
Please do not adjust margins