Biocatalytic self-assembly of 2D peptide-based nanostructures

Article abstract of DOI:10.1039/c1sm05981e

Peptide based 2D nanostructures of micronscale size in both X and Y dimensions are extremely rare because amino acid chirality favours helical structures, and nucleation-growth mechanisms usually favour uni-directional growth. We demonstrate the production of extended two-dimensional (2D) peptide nanostructures via the thermolysin catalysed condensation of Fmoc protected hydrophilic amino acid (serine, Fmoc-S) and a hydrophobic amino acid ester (phenylalanine, F-OMe). We propose that lateral self-assembly is enabled by the reversible nature of the system, favouring the thermodynamic product (extended sheets) over kinetically favoured 1 dimensional structures. Fmoc-SF-OMe forms extended arrays of β-sheet structures interlock via π-stacking between Fmoc groups. We propose that, due to its alternating hydrophilic/hydrophobic amino acid sequence, amphiphilic sheets presenting either phenyl or hydroxyl functionality are formed that assemble pair-wise, thereby shielding hydrophobic groups from the aqueous environment. Formation of these structures was supported by fluorescence emission spectroscopy, FTIR and XRD analysis and molecular mechanics minimization. At enhanced enzyme concentrations, hierarchical self-assembly was observed giving rise to spherulitic structures, with the number of spherulites dictated by enzyme concentration.

Full text of DOI:10.1039/c1sm05981e

View Article Online / Journal Homepage / Table of Contents for this issue
Dynamic Article Links <
C
Soft Matter
Cite this: Soft Matter, 2011, 7, 10032
www.rsc.org/softmatter
PAPER
Biocatalytic self-assembly of 2D peptide-based nanostructures†
Meghan Hughes,a Haixia Xu,b Pim W. J. M. Frederix,ad Andrew M. Smith,c Neil T. Hunt,d Tell Tuttle,a
b
Ian A. Kinloch and Rein V. Ulijn
a
*
Received 27th May 2011, Accepted 16th August 2011
DOI: 10.1039/c1sm05981e
Peptide based 2D nanostructures of micronscale size in both X and Y dimensions are extremely
rare because amino acid chirality favours helical structures, and nucleation-growth mechanisms
usually favour uni-directional growth. We demonstrate the production of extended two-dimensional
(2D) peptide nanostructures via the thermolysin catalysed condensation of Fmoc protected
hydrophilic amino acid (serine, Fmoc-S) and a hydrophobic amino acid ester (phenylalanine, F-
OMe). We propose that lateral self-assembly is enabled by the reversible nature of the system,
favouring the thermodynamic product (extended sheets) over kinetically favoured 1 dimensional
structures. Fmoc-SF-OMe forms extended arrays of b-sheet structures interlock via p-stacking
between Fmoc groups. We propose that, due to its alternating hydrophilic/hydrophobic amino acid
sequence, amphiphilic sheets presenting either phenyl or hydroxyl functionality are formed that
assemble pair-wise, thereby shielding hydrophobic groups from the aqueous environment.
Formation of these structures was supported by fluorescence emission spectroscopy, FTIR and
XRD analysis and molecular mechanics minimization. At enhanced enzyme concentrations,
hierarchical self-assembly was observed giving rise to spherulitic structures, with the number of
spherulites dictated by enzyme concentration.
These systems are 2D at the microscopic and macroscopic
Introduction
length scales but are ultimately composed of 1D fibrillar
nanostructures. Peptoids, which are non-chiral polymers with
peptide like side chains, were recently shown to form 2D
structures.16 Flat structures with limited lateral directions
(belts) were recently described by Cui et al.17 and Shao et al.18
Both systems were based on peptide amphiphiles with alter-
nating hydrophobic and hydrophilic amino acid residues.
These sequences are well known to have a predisposition to
form b-sheet like structures for oligo-peptides3,10,14 with
a hydrophobic and hydrophilic face, that have a tendency to
dimerise, thereby hiding the hydrophobic faces from water.
These further assemble to produce tapes and fibres, as dis-
cussed above.
There is a significant interest in the production of functional
materials by exploiting supramolecular chemistry.1,2 Peptides
are of particular interest as building blocks - they have a rich
chemical diversity and an inherent ability to effectively inter-
face with biological systems.3 Peptide based nanostructures
usually take the shape of zero- or one-dimensional structures,
such as micelles, tubes,4–6 fibers,7–9 or (twisted) ribbons.10
Chiral uni-directional structures are often favoured due to (i)
the inherent chirality11 in peptide building blocks and (ii) the
nucleation and growth mechanisms12 which generally involves
formation of nuclei that have a preference to extend uni-
directionally. Flat, sheet-like structures based on alternating
hydrophobic/hydrophilic polypeptides have already been
proposed in the mid-70s13 and oligopeptides composed of
interwoven fibrous networks14 as well as hybrid polysaccharide/
peptide amphiphile membranes15 have been demonstrated.
Although molecular self-assembly proposes that supramo-
lecular material properties can be fully encoded into molecular
building blocks, it is increasingly apparent that the self-
assembly pathway largely dictates the final structure.19 In the
biological world, self-assembly processes are commonly
coupled to catalysis which enables tight control over the self-
assembly pathway. The combination of (bio-) catalysis and
molecular self-assembly provides a relatively new concept in
laboratory based supramolecular chemistry. The approach
consists of converting non-assembling precursors into self-
assembly building blocks, usually by enzymatic removal of
a charged or steric group that prevents self-assembly,19–22 or by
condensation (reversed hydrolysis) of amino acid derivatives to
aWestCHEM, Department of Pure & Applied Chemistry, University of
Strathclyde, Glasgow, UK. E-mail: rein.ulijn@strath.ac.uk
bSchool of Materials, University of Manchester, Manchester, UK
cLehrstuhl Biomaterialien, University of Bayreuth, Germany
dSUPA, Department of Physics, University of Strathclyde, Glasgow, UK
† Electronic Supplementary Information (ESI) available: concentration
yield correlations, optical images of precipitates, supporting TEM and
SEM images, computer models, chemical synthesis of Fmoc-SF-OMe
and characterisation,
10.1039/c1sm05981e/
further
WAXS
data.
See
DOI:
10032 | Soft Matter, 2011, 7, 10032–10038
This journal is ª The Royal Society of Chemistry 2011

View Article Online
produce self-assembling peptide amphiphiles.23 Additionally,
the reversibility of the reaction will allow for the formation of
the most thermodynamically stable nanostructures.23
FTIR
FTIR spectra were acquired in a Bruker Vertex spectrometer
with a spectral resolution of 2 cmÀ1. The spectra were obtained
by averaging 64 interferograms for each sample. Measurements
were performed in a standard IR cuvette (Harrick Scientific), in
which the sample was contained between two CaF2 windows
(thickness, 2 mm) separated by a 25-mm PTFE spacer. All sample
manipulations were performed in a glove box to minimize
interference from atmospheric water vapour; D2O was used as
the solvent for all the IR measurements.
The self-assembly strategy is based on aromatic peptide
amphiphiles, i.e. short (generally less than five amino acids)
peptides appended with aromatic groups.6,26–31 These are thought
to self-assemble via a combination of hydrogen bonding between
peptide groups and p-stacking/hydrophobic interactions
between aromatic groups. Despite their relative simplicity, the
assembly rules are not fully understood with a few exceptions
which are composed of aromatic ligands such as N-fluorenyl-
9-methoxycarbonyl (Fmoc) or naphthalene derivatives linked to
hydrophobic peptides such as FF,6 LLL,32 AV30 giving rise to
chiral, 1D nanostructures.
WAXS
WAXS analysis were performed using a Philips X’Pert diffrac-
ꢀ
tometer with a wavelength of 1.5406 A. Fmoc-SF-OMe gel and
The peptide sequence used here involves a combination of an
Fmoc-protected hydrophilic (serine, S) and the methyl ester of
a hydrophobic (phenylalanine, F) residue. This combination was
found to give rise to stable self-assembling structures. It was
buffer solution were spread on two separate silica substrates as
film and allowed to air dry prior to data collection.
previously discovered by using
a dynamic combinatorial
TEM
library1,33,34 of precursor components, based on Fmoc-amino
acids (serine, S and threonine, T) and a range of amino acid
methyl esters all present simultaneously in a mixture containing
a non-specific protease which was able to catalyse condensation
of these residues and with the self-assembly thermodynamics
driving the condensation reactions. It was found that Fmoc-SF-
OMe was produced preferentially,35 which indicates it has
a strong tendency for molecular self-assembly.
Carbon-coated copper grids (200 mesh) were glow discharged in
air for 30 s. A 10 mL volume of gel was transferred onto the
support film and blotted down using filter paper. 20 ml of nega-
tive stain (1% aqueous methylamine vanadate obtained from
Nanovan; Nanoprobes) was applied and the mixture blotted
again using filter paper to remove excess. The dried specimens
were then imaged using a LEO 912 energy filtering transmission
electron microscope operating at 120kV fitted with 14bit/2 K
Proscan CCD camera.
Experimental
Fmoc-S (Bachem, UK) and F-OMe (Sigma-Aldrich, UK) were
weighed as a 20 : 80 mM ratio in a glass vial. The powder mixture
was dissolved in 2 mL of 0.1 M pH 8 potassium phosphate buffer
with the addition of the 1mg mlÀ1 lyophilised thermolysin powder
(bacillus Thermoproteolyticus rokko from Sigma-Aldrich, UK)
unless otherwise stated. The solution was vortexed and sonicated
to ensure dissolution. Samples were incubated at room
temperature.
CryoTEM
Gel samples were blotted onto a thin film grid of 100–200 nm.
The grid was then plunged into liquid ethane (temperatures
below À170 ꢀC) and transferred to a GATAN 626 cryoholder
and imaged using a JEOL 2100 transmission electron microscope
fitted with a GATAN 4 K Ultrascan camera. The cryoTEM
analysis was carried out at Unilever, Bedford.
Optical microscopy
HPLC
10 mL of sample was transferred onto a glass slide and covered
with a cover slip. The samples were then examined using a Zeiss
Axio Imager A1 optical microscope equipped with polarising
lenses at 0ꢀ using a bright lens and transmitted light. Homoge-
neity was ensured by repeating in triplicate.
ꢀ
A Dionex P680 system operating with a Macherey-Nagel 250 A,
4.6 Â 250 mm, C18 column was used for reversed phase HPLC.
10 mL of sample was injected. The mobile phase was comprised of
water and acetonitrile ramped from 20–80% over 35 min at a flow
rate of 1 mL minÀ1. Detection of the peptide amphiphiles was
carried out using a UVD170U UV-Vis detector at a 300 nm
wavelength.
Synthesis of Fmoc-SF-Ome
Fmoc-S (1 g, 2.8 mM), F-OMe (0.72g, 3.4 mM) and HBTU
(1.26g, 3.4 mM) were dissolved in 10 mL of DMF. 1.041 mL
(5.6 mM) of DIPEA was added and solution stirred for 24 h. The
solvent was removed by evaporation in vacuo. The resulting solid
was dissolved in ethyl acetate, washed in duplicate with equal
volumes of 1M sodium bicarbonate, brine and 1 M HCl. The
organic layer dried using sodium sulphate and filtered. Solvent
removed by evaporation in vacuo. The white solid then purified
over a silica column using 2.5%–7.5% methanol in chloroform.
Aliquots evaporated at room temperature and pressure and
white solid combined (21% yield, 98% purity).
Fluorescence
Fluorescence emission spectra were measured on a Jasco FP-
6500 spectrofluorometer with fluorescence measured orthogo-
nally to the excitation light with a scanning speed of 200 nm
minÀ1. Excitation light at 280 nm and emission data range
between 300 nm and 600 nm. The emission spectra were
measured with a bandwith of 3 nm with a medium response and
a 1 nm data pitch. Spectra were normalised to maximum
intensity.
This journal is ª The Royal Society of Chemistry 2011
Soft Matter, 2011, 7, 10032–10038 | 10033

View Article Online
Computer modelling
aromatic peptide amphiphiles,6,26–32 the self-assembly results in
a weak opaque suspension which remains stable over a number
of weeks (see Fig. 1(b)). It should be noted that the chemically
synthesised compound 3, in a pure state or as a 1 : 3 mixture of
compounds 3 and 2, forms amorphous precipitates in buffer with
no evidence of formation of ordered nanostructures (see ESI,†
Fig. S2). Initial investigation of the self-assembled structure was
carried out using TEM and cryoTEM (see Fig. 1(c)–(d)and
additional fields of view supplied in ESI,† Fig. S3). This revealed
sheet-like structures of many microns in both X and Y dimen-
sions, in addition to twisted nano-ribbons.
All structures were constructed in ArgusLab36 and subsequently
energy-minimized in vacuum using the conjugate gradient
method as implemented in NAMD.37 The minimization proce-
dure was carried out for 10,000 steps, at which point the fluc-
tuations in the energy were <10À4 kcal molÀ1 and the structure
was considered to be converged. The CHARMM c36a3 force
field38 was used to minimize the structure. VMD39 was used to
visualize the results.
Fluorescence emission spectroscopy was used to monitor
changes in fluorenyl environment as the reaction proceeded (see
Fig. 2(a)). It should be noted that a substantial reduction in
signal intensity was observed over time, this is, at least in part,
due to the sample becoming increasingly opaque. The spectra
were normalised to allow a more accurate determination of
peak formation and shifting; whilst original intensity changes
of each peak of interest are shown in Fig. 2(b). It is evident that
a number of structural changes occur. The starting mixture
shows two predominant features emitting at approximately 320
and 330 nm, representing monomeric and aggregated forms of
Fmoc-S precursors. Within 10 min a red shift accompanied by
an increase in intensity indicates a supramolecular transition
producing a more intensely emitting fluorenyl species. The
formation of a red-shifted peak at 365 nm accompanied by
system quenching (Fig. 2(b)) could be associated with
formation of extended aggregates through p- stacked
systems as previously observed, e.g. for Fmoc-LLL32 and
Fmoc-FF.6a,d However, it should be noted that the red-shift is
much less pronounced for the system under study, suggesting
that fluorenyl face to face p-stacking may not be optimal in
these systems. After 24 h, the free fluorenyl peak has nearly
disappeared, suggesting that few Fmoc-S or Fmoc-SF-OMe
species are in the un-assembled state, which is consistent with
the condensation yield observed.
Results and discussion
Condensation of Fmoc-S, 1, and F-OMe, 2 in a 1 : 4 ratio by
thermolysin from Bacillus thermoproteolyticus rokko resulted in
formation of Fmoc-SF-OMe, 3, (Fig. 1(a)) at a constant
conversion of 98% after 24 h with an initial reactant concentra-
tion of 20/80 mM.35 The system exhibits typical equilibrium
behaviour, with higher yields of 3 formed at corresponding
increased concentrations of starting materials (see ESI,† Fig. S1).
Rather than forming translucent hydrogels typically observed for
The hydrogen bonding interactions of peptide moieties of 3
were then monitored using FTIR spectroscopy focusing on
carbonyl stretching in the amide I region. Fig. 2(c) illustrates the
time-dependent correlation peaks at 1640 cmÀ1, 1680 cmÀ1 and
1650 cmÀ1 peak intensities. As they all appear at the same point
over the reaction coordinate it can be assumed that they
represent the molecular organisation of nanostructural sheets.
The pair of peaks at 1640 cmÀ1 and 1680 cmÀ1 are characteristic
of the formation of a b-sheet arrangement of the peptide
moieties (see Fig. 2(d)).6,32,40 This structure becomes evident
from 5 h and also remains in this arrangement over 72 h. It is
noticeable that the peaks gradually shift to lower frequencies
during this time, consistent with hydrogen bonding strength
increasing. This is accompanied by the occurrence of a peak due
to the carbonyl stretch associated with the methyl ester moiety
at 1728 cmÀ1 in addition to the peak at 1745 cmÀ1 attributable to
the carbonyl of free F-OMe. However, a prominent peak at
1650 cmÀ1, indicative of random order of peptides40 is also
present, which indicates that although the structure is organised
as an extended anti-parallel b-sheet arrangement, there is also
a disordered component. In contrast to the similar dynamics of
the FTIR peaks located at 1640 and 1680 cmÀ1 (beta sheet) and
1650 cmÀ1 (pseudo random coil), it is interesting to note the
Fig. 1 (a) reaction scheme of Fmoc-S, 1, coupled with F-OMe, 2, to
form Fmoc-SF-OMe, 3, via condensation using protease enzyme ther-
molysin. (b) 3 self-assembles to form nano-sheets within a self-supporting
opaque suspension. (c) and (d) TEM and Cryo-TEM images of nano-
sheets of 3 obtained after 24 h. (e) The schematic representation of the
molecular assembly of 3 to form bilayer nano-sheets.
10034 | Soft Matter, 2011, 7, 10032–10038
This journal is ª The Royal Society of Chemistry 2011

View Article Online
Fig. 2 (a) Fluorescence emission spectra monitored over 72 h. (b) Relative intensity of 320 nm, 330 nm and 365 nm peaks within fluorescence spectra
monitored with time (intensity normalised). (c) Relative intensity of 1640 cmÀ1, 1680 cmÀ1 and 1650 cmÀ1 bands within FTIR spectra monitored with
time. (d) Amide I region of FTIR absorbance spectra monitored over 72 h, representing carbonyl stretching mode of amino acid residues. (e) WAXS
data of dried sample at 24 h.
appearance of the peak at 1620 cmÀ1, which occurs only after
24 h. The assignment of this peak is currently unclear but the
timescales are consistent with higher order structure formation
of spherulites as discussed below.‡
(p-b) molecular association results in fibrillar network struc-
tures,6a,32 but has also been observed to produce belts when
Fmoc-dipeptides with hydrophilic and (modified) hydrophobic
residues were used.18 We propose a model that is consistent
with these reflections obtained for the WAXS data and the
observed planar sheets (see Fig. 3). The Fmoc-groups form
stacked pairs rather than a continuous p-stack proposed for
Fmoc-FF.6a,d In that system, the C-terminal phenyl group
intercalated between the Fmoc groups to create a continuous
p- stack. In the case of Fmoc-SF-OMe the methyl ester
prevents the molecules assuming this configuration, which
would also explain the absence of a 450 nm peak observed in
fluorescence by Smith et al.6a and Xu et al.32 The phenyl
moieties of phenylalanine residues can interact with those from
a neighbouring array to form the p-b bilayer as illustrated in
Fig. 3. This molecular association also explains the formation
In order to confirm the proposed structure, WAXS diffrac-
tion patterns were obtained (see Fig. 2(e)). A prominent peak
with 5 higher order reflections indicates a repeating pattern of
ꢀ
16 A which we have correlated to the repeating unit along the
ꢀ
peptide backbone, and additional peaks corresponding to 4.8 A
ꢀ
and 3.7 A which relate to b- sheet spacing and p-stack spacing
respectively.32 Typically, this type of p-interlocked b-sheet
‡ The assignment of this peak is not clear though it is consistent with
intermolecular beta sheet formation observed in proteins and may
indicate a secondary interaction of beta sheet structures as the time
scales are consistent with higher order structure formation of spherulites.
This journal is ª The Royal Society of Chemistry 2011
Soft Matter, 2011, 7, 10032–10038 | 10035

View Article Online
hydroxyl group, which may explain the observed shift in the
methyl ester peak (see Fig. 2(d)).
In previous systems,17,18 growth has always been predomi-
nantly along one axis unless the chirality of the peptides has
been removed, and the nanostructure is comprised solely of
non-natural peptides forming crystalline sheets.16 In certain
cases, for example in b-hairpin assemble, lateral growth is
unfavoured because all peptides assembling in register, in which
case the width is dictated by peptide length.8 In cases where
lateral association through formation of ‘sticky ends’ is possible,
it remains unknown as to what causes such growth restric-
tions.17,18 It is most likely related to the size of initial nuclei
which have a preference to grow uni-directionally, i.e. a kinetic
effect. A key difference in the system described here, is that the
building blocks themselves can be reversibly formed and
hydrolysed, giving rise to a system which will be less prone to
kinetic locking.
TEM images suggest a mechanism for sheet formation, where
twisted tapes are predominantly observed at the early stages (see
ESI,† Fig. S5). 24 h after the reaction has been initiated,
branching of the twisted nanoribbons and nucleation of ribbons
from the edge of sheets is observed. The images suggest
a progression through several phases before reaching the final
sheet structures, which we believe to be related to constant
enzymatic structure correction at the edges of existing structures.
During the latter stages of self-assembly (24–72 h), a remark-
able further hierarchical self-assembly was observed which is
tunable by enzyme concentration. SEM analysis reveals that
macroscopic spherulitic structures are formed of several hundred
microns in diameter (see Fig. 4). These spherulites are sur-
rounded by non-spherulitic areas of material being constructed
of layered lamellar structures, thought to represent multilayers of
the 2D sheets discussed, which could feasibly interact via
hydrogen bonding of the exposed hydroxyl groups of the serine
residues. Higher magnification SEM images of the spherulitic
structures shows that these structures are composed of sheets,
with no obvious nucleation points at their centres. Peptide based
spherulitic structures have been obtained previously by solvent
induced phase transitions42 or by hierarchical assembly from
aromatic peptide amphiphiles.43
Fig. 3 Molecular association models. (a) 8-mer anti-parallel b-sheet
arrangement. (b) 8-mer side view. (c) 16-mer side view of two separate 8-
mers where their hydrophobic residues interact to from a bilayer.
of extended 2D structures rather than 1D fibers or tubes, since
next to the p-stacking of fluorenyl groups and anti- parallel b-
sheets there is a driving force that extends the structure in
a biaxial direction, yet keeping its linearity. Overall, this
assembly results in the hydrophobic phenylalanines being
completely buried within the bilayer, while the hydrophilic
serine residues are orientated outwards and can hydrogen bond
to the aqueous environment or with serine residues of
a neighbouring self-assembled bilayer system to produce multi-
bilayers, similar to Shao et al.18 and to well-documented
bilayered tapes in larger b-sheet self-assembling peptides.10,41
From the structure in Fig. 3 a 64-mer was built of 8 Â 4 Â 2
molecules (see ESI,† Fig. S4). The proposed structure proved
to be stable under minimization using molecular mechanics,
It has previously been demonstrated that self-assembly is
favoured in the direct vicinity of enzymes23–25 and therefore it is
tempting to propose enzymatic control of spherulite formation.
To investigate a possible link between enzyme concentration and
spherulite formation, a range of enzyme concentrations (0.1, 1,
10 mg mLÀ1) were analysed using polarising optical microscopy.
The spherulites display birefringence when viewed under cross
polarisers (see Fig. 5). Each of the samples appear as opaque
suspensions at 24 h. However, the rate at which each becomes
self-supporting varies; 0.1 mg mLÀ1 thermolysin after approxi-
mately 24 h, 1 mg mLÀ1 within a matter of hours, and
10 mg mLÀ1 after only a few minutes of the reaction being
initiated; which is to be expected. At a low enzyme concentration
of 0.1 mg mLÀ1, no spherulitic structures form. This has been
further confirmed by the formation of solely the layered lamellar
structures visible using SEM (see ESI,† Fig. S6). Additionally,
the use of a higher enzyme concentration (10 mg mLÀ1) displayed
a dramatic increase in the number of these spherulitic structures
are formed (see Fig. 5). This signifies that the enzyme plays a key
ꢀ
and exhibits repeating units with average distances of 16.2 A
ꢀ
(Fmoc-column to Fmoc-column), 4.9 A (peptide backbone to
ꢀ
backbone), and 3.7 A (Fmoc- to Fmoc- within the p-stacked
columns), which are in excellent agreement with the reflections
ꢀ
observed in WAXS. Additionally, 10.1 A can be observed for
Fmoc-pair to Fmoc-pair in the stacking direction, which also
corresponds to a peak on the WAXS data (see Fig. 2(d)). The
proposed model is also consistent with the other spectroscopic
results such as the formation of stable b-sheets observed from
FTIR. The minimized model also shows that the methyl ester
may form an intramolecular hydrogen bond with the serine
10036 | Soft Matter, 2011, 7, 10032–10038
This journal is ª The Royal Society of Chemistry 2011

View Article Online
Fig. 4 SEM imaging of self-assembled Fmoc-SF-OMe. (a) Spherical structures and ‘halved’ spherical structures can been seen to lie on a layer of the
nano-sheet material. (b) Spherical structures encased in nano-sheet material. (c,d) Spherical structures have crystalline spherulitic centres with no
obvious nucleation point and are surrounded by a layer of sheets. (d) Lamellar structure of non-spherulitic areas.
role in nucleation process to form these spherulites, however it
does not affect the ultimate size of the crystals (diameter of just
over 100 mM).
Conclusions
In summary, we have demonstrated a route towards the
production of two-dimensional nanostructures by enzymatic
condensation of chiral amino acid derivatives. We propose that
lateral self-assembly is enabled by the reversible nature of the
system, favouring the thermodynamic product (extended sheets)
over kinetically favoured 1 dimensional structures. The combi-
nation of hydrophilic and hydrophobic amino acids, which are
interlocked via p-stacking interactions of Fmoc groups, produce
amphiphilic bilayers, with the hydrophobic faces hidden from
water and further stabilised via p-stacking of phenyl rings. The
production of nano-sheets with well-defined chemical composi-
tion is of interest in production of surfaces with precisely pre-
sented bioactivity, and may also be used in templating,44,45 and
by combination with biocatalysis, dynamic template structures
may be feasible.46,47 The remarkable structural,3–6 biological,7–9
Fig. 5 Optical and polarised images of macrostructure. Birefringence or
crystalline structures is observed when viewed under cross-polarisers. A
and electronic18,32 properties of peptide based nanostructures,
relationship between enzyme concentration and number of spherulitic
structures is observed.
combined with the low cost of materials based on dipeptides or
This journal is ª The Royal Society of Chemistry 2011
Soft Matter, 2011, 7, 10032–10038 | 10037

View Article Online
22 C. Wang, Q. Chen, Z. Wang and X. Zhang, Angew. Chem., Int. Ed.,
2010, 49, 8612.
23 (a) S. Toledano, R. J. Williams, V. Jayawarna and R. V. Ulijn, J. Am.
Chem. Soc., 2006, 128, 1070; (b) R. J. Williams, A. M. Smith,
R. Collins, N. Hodson, A. K. Das and R. V. Ulijn, Nat.
Nanotechnol., 2009, 4, 19.
24 R. de la Rica, K. I. Fabijanic, A. Baldi and H. Matsui, Angew. Chem.
Int. Ed., 2010, 49, 1447.
25 H. Wang, Z. Wang, D. Song, J. Wang, J. Gao, L. Wang, D. Kong and
Z. Yang, Nanotechnology, 2010, 21, 155602.
26 R. Vegners, I. Shestakova, I. Kalvinsh, R. M. Ezzell and
P. A. Janmey, J. Pept. Sci., 1995, 1, 371.
27 D. M. Ryan, T. M. Doran, S. B. Anderson and B. L. Nilsson,
Langmuir, 2011, 27, 4029.
28 S. Debnath, A. Shome, D. Das and P. K. Das, J. Phys. Chem. B, 2010,
114, 4407.
amino acid derivatives, suggests future applications in material
science, biomedicine and nanotechnology.
References
1 J. M. Lehn, Supramolecular Chemistry - Concepts and Perspectives,
VCH, Weinheim, 1995.
2 G. M. Whitesides and B. Grzybowski, Science, 2002, 295, 2418.
3 (a) X. B. Zhao, F. Pan, H. Xu, M. Yaseen, H. H. Shan,
C. A. E. Hauser, S. G. Zhang and J. R. Lu, Chem. Soc. Rev., 2010,
39, 3480; (b) I. W. Hamley, Soft Matter, 2011, 7, 4122; (c)
M. Zelzer and R. V. Ulijn, Chem. Soc. Rev., 2010, 39, 3351.
4 M. R. Ghadiri, J. R. Granja, R. A. Milligan, D. E. McRee and
N. Khazanovich, Nature, 1993, 366, 324.
29 (a) L. Chen, K. Morris, A. Laybourn, D. Elias, M. R. Hicks,
A. Rodger, L. Serpell and D. J. Adams, Langmuir, 2010, 26, 5232;
(b) E. K. Johnson, D. J. Adams and P. J. Cameron, J. Mater.
Chem., 2011, 21, 2024.
30 G. Cheng, V. Castelletto, C. M. Moulton, G. E. Newby and
I. W. Hamley, Langmuir, 2010, 26, 4990.
31 D. Bardelang, F. Camerel, J. C. Margeson, D. M. Leek, M. Schutz,
B. Zaman, K. Yu, D. V. Soldatov, R. Ziessel, C. I. Ratcliffe and
J. A. Ripmeester, J. Am. Chem. Soc., 2008, 130,
3313.
32 H. Xu, A. K. Das, M. Horie, M. S. Shaik, A. M. Smith, Y. Luo,
X. Lu, R. Collins, S. Y. Liem, S. Song, P. L. A. Popelier,
M. L. Turner, P. Xiao, I. A. Kinloch and R. V. Ulijn, Nanoscale,
2010, 2, 960.
33 N. Sreenivasachary and J. M. Lehn, Proc. Natl. Acad. Sci. U. S. A.,
2005, 102, 5938.
34 S. Otto, R. L. E. Furlan and J. K. M. Sanders, Science, 2002, 297,
590.
35 A. K. Das, A. R. Hirst and R. V. Ulijn, Faraday Discuss., 2009, 143,
293.
36 M. A. Thompson, 2004, ArgusLab 4.0.1, Seattle, WA Planaria
Software LLC.
5 M. Reches and E. Gazit, Science, 2003, 300, 625.
6 (a) A. M. Smith, R. J. Williams, C. Tang, P. Coppo, R. F. Collins,
M. L. Turner, A. Saiani and R. V. Ulijn, Adv. Mater., 2008, 20, 37;
(b) V. Jayawarna, M. Ali, T. A. Jowitt, A. F. Miller, A. Saiani,
J. E. Gough and R. V. Ulijn, Adv. Mater., 2006, 18, 611–614; (c)
A. Mahler, M. Reches, M. Rechter, S. Cohen and E. Gazit, Adv.
Mater., 2006, 18, 1365; (d) C. Tang, A. M. Smith, R. F. Collins,
R. V. Ulijn and A. Saiani, Langmuir, 2009, 25, 9447.
7 G. A. Silva, C. Czeisler, K. L. Niece, E. Beniash, D. A. Harrington,
J. A. Kessler and S. I. Stupp, Science, 2004, 303, 1352.
8 L. Haines-Butterick, K. Rajagopal, M. Branco, D. Salick,
R. Rughani, M. Pilarz, M. S. Lamm, D. J. Pochan and
J. P. Schneider, Proc. Natl. Acad. Sci. U. S. A., 2007, 104, 7791.
9 E. F. Banwell, E. S. Abelardo, D. J. Adams, M. A. Birchall,
A. Corrigan, A. M. Donald, M. Kirkland, L. C. Serpell,
M. F. Butler and D. N. Woolfson, Nat. Mater., 2009, 8, 596.
10 A. Aggeli, M. Bell, N. Boden, J. N. Keen, P. F. Knowles,
T. C. B. McLeish, M. Pitkeathly and S. E. Radford, Nature, 1997,
386, 259.
11 P. Jonkheijm, P. van der Schoot, A. P. H. J. Schenning and
E. W. Meijer, Science, 2006, 313, 80.
12 J. Cornelissen, A. E. Rowan, R. J. M. Nolte and N. Sommerdijk,
Chem. Rev., 2001, 101, 4039.
13 A. Brack and L. E. Orgel, Nature, 1975, 256, 383.
14 S. G. Zhang, T. Holmes, C. Lockshin and A. Rich, Proc. Natl. Acad.
Sci. U. S. A., 1993, 90, 3334.
15 R. M. Capito, H. S. Azevedo, Y. S. Velichko, A. Mata and S. I. Stupp,
Science, 2008, 319, 1812.
37 J. C. Phillips, R. Braun, W. Wang, J. Gumbart, E. Tajkhorshid,
E. Villa, C. Chipot, R. D. Skeel, L. Kale and K. Schulten, J.
Comput. Chem., 2005, 26, 1781.
38 B. R. Brooks, et al., J. Comput. Chem., 2009, 30, 1545.
39 W. Humphrey, A. Dalke and K. Schulten, J. Mol. Graphics, 1996, 14,
33.
40 A. Barth and C. Zscherp, Q. Rev. Biophys., 2002, 35, 369.
41 H. Yokoi, T. Kinoshita and S. Zhang, Proc. Natl. Acad. Sci. U. S. A.,
2005, 102, 8414.
16 K. T. Nam, S. A. Shelby, P. H. Choi, A. B. Marciel, R. Chen, L. Tan,
T. K. Chu, R. A. Mesch, B. C. Lee, M. D. Connolly, C. Kisielowski
and R. N. Zuckermann, Nat. Mater., 2010, 9, 454.
42 P. Zhu, X. Yan, Y. Su, Y. Yang and J. Li, Chem.–Eur. J., 2010, 16,
3176.
17 H. Cui, T. Muraoka, A. G. Cheetham and S. I. Stupp, Nano Lett.,
2009, 9, 945.
43 W. Wang and Y. Chau, Soft Matter, 2009, 5, 4893.
44 N. Sharma, A. Top, K. L. Kiick and D. J. Pochan, Angew. Chem., Int.
Ed., 2009, 48, 7078.
45 M. S. Lamm, N. Sharma, K. Rajagopal, F. L. Beyer, J. P. Schneider
and D. J. Pochan, Adv. Mater., 2008, 20, 447.
46 F. Patolsky, Y. Weizmann and I. Willner, Nat. Mater., 2004, 3,
692.
18 H. Shao and J. Parquette, Chem. Commun., 2010, 46, 4285.
19 A. R. Hirst, S. Roy, M. Arora, A. K. Das, N. Hodson, P. Murray,
N. Javid, J. Sefcik, J. Boekhoven, J. H. van Esch, S. Santabarbara,
N. T. Hunt and R. V. Ulijn, Nat. Chem., 2010, 2, 1089–1094.
20 (a) Z. M. Yang, H. W. Gu, D. G. Fu, P. Gao, J. K. Lam and B. Xu,
Adv. Mater., 2004, 16, 1440; (b) Y. Zhang, H. W. Gu, Z. M. Yang and
B. Xu, J. Am. Chem. Soc., 2003, 125, 13680.
47 N. Ostrov and E. Gazit, Angew. Chem., Int. Ed., 2010, 49,
3018.
21 S. Winkler, D. Wilson and D. L. Kaplan, Biochemistry, 2000, 39(12),
739.
10038 | Soft Matter, 2011, 7, 10032–10038
This journal is ª The Royal Society of Chemistry 2011