[System] polypeptide self-assembly literature information extraction

[User]
Dipeptide-conjugates can be efficient low molecular weight hydrogelators.
However, the effective design of a gelator for a specific application is
compromised by the lack of a clear understanding of the design rules that govern
assembly and hence gelation. Here, we report a library of
naphthalene-dipeptides, their physical chemical properties, and gelation
ability. We have varied both the amino acids in the dipeptides and the
substitution on the naphthalene ring to allow variation of the structure
throughout the molecule. We have examined the effects of these permutations on
the critical micelle concentration and air-water partition coefficient at high
pH and the apparent pK(a). We show that there is a clear link between these
properties and the predicted hydrophobicity of the overall conjugates, rather
than the properties varying with, for example, the dipeptide sequence. The
majority of these dipeptide-conjugates are effective hydrogelators, although
there is no apparent link between the solution properties and whether or not a
conjugate is a hydrogelator. Nevertheless, where gelation occurs, the link
between hydrophobicity and apparent pK(a) allows the prediction of the pH at
which a gel will be formed and hence informed choice of gelator for specific
applications.

Materials. All chemicals and solvents were purchased from
Sigma-Aldrich and used as received. Millipore water (resistivity
= 18.2 mΩ/cm) was used throughout. Full synthesis and char-
acterization details for the all dipeptides and intermediates are
available in the Supporting Information. The Fmoc-dipeptides
were synthesized as described elsewhere.34
Transmission Electron Microscopy. Samples for examina-
tion by TEM were prepared on Formvar/carbon film coated 400-
mesh copper grids (Agar scientific). The required amount of GdL
was added to a solution of dipeptide derivative (0.5 wt % solution
at pH 10.7) and allowed to gelate. The gel was sampled by
removing 10 μL, placing on a grid, and allowing adsorption for
1 min followed by a 1 min wash and two 1 min negative stains
using 2% w/v uranyl acetate. Where necessary samples for TEM
were prepared in situ by immediately placing grids inverted onto
10 μL droplets of the gelation solution. In a humid chamber at
room temperature, material was allowed to adsorb onto the grids
and removed after 120 min followed by 5 min of drying and two
stains as above. Negatively stained grids were allowed to dry and
examined on a Hitachi-7100 TEM operated at 100 kV. Images
were acquired digitally using an axially mounted (2000 (cid:2) 2000
pixels) Gatan Ultrascan 1000 CCD camera (Gatan, Oxford, UK).
Surface Tension Measurements. The surface tension mea-
surements were performed on the high throughput Delta-8
system (Kibron Inc.), and the data were analyzed by the
Delta-8 Manger software. The 200 μL concentrated dipeptide
samples were transferred onto the first column of a standard
Nunclon 96 well plate. A series of concentrations were prepared
by sequential dilution with pH 9-10 (tested by pH paper) water
across the plate, with the concentration decreased by a dilution
factor for each column using a transfer-and-mix protocol.
Final 50 μL samples were transferred to the detection plate
for measurements.
The adsorption isotherm curves of different samples were
obtained and the apparent air-water partition coefficient (Kaw),
cross-sectional areas (As), and critical micelle concentration (cmc)
calculated based on the Gibbs adsorption equation
Determination of the Apparent pKa. A FC200 pH probe
(HANNA instruments) with a (6 mm (cid:2) 10 mm) conical tip was
employed for all pH measurements. The stated accuracy of the pH
measurements is (0.1. The pKa values of dipeptide solutions were
determined by titration via the addition of aliquots of a 0.1 M HCl
solution. pH values were recorded until reaching a stable value
after each addition during the titration process. To prevent the
formation of gel, the solutions were gently stirred, thus keeping
the solution liquid during the whole “titration” process. For select
samples, heating between aliquot addition and recooling was
carried out to check whether mixing and equilibration had
occurred.22 No deviation between pKa measurements with or
without heating was observed, and hence the values reported
are for samples without heating.
Gelation Studies. The dipeptide derivative (25.0 mg) was
suspended in deionized water (5.00 mL). An equimolar quantity
of NaOH (0.1 M, aq) was added and the solution gently stirred for
30 min until a clear solution was formed. The pH of this solution
was measured to be 10.7. To prepare hydrogels, solutions were
added to measured quantities of glucono-δ-lactone (GdL) accord-
ing to the requirements of final pH and the samples left to stand
for 24 h. To obtain the highest pH of gelation, a series of dipeptide
samples were prepared with varying concentrations of GdL such
that different final pHs of between pH 8 and pH 3 were achieved.
After 24 h, the samples were examined by the vial inversion test.
The highest pH of gelation was defined by the pH under which the
stable gels were formed, and no flow of liquid was found in the
Determination of the Minimum Gelation Concentration.
Stock solutions of dipeptide-conjugates at a concentration of
5 mg/mL (0.5 wt %) were prepared. These were then diluted with
pH 10 water to a number of concentrations. To adjust the pH, for
each dipeptide-conjugate solution, a number of samples were
prepared with different quantities of GdL to ensure the prepara-
tion of samples with a final pH between 3.2 and 3.6. All the
samples were left to stand at least 24 h. The minimum gelation
concentrations were determined by the lowest concentration at
which a self-supporting gel was formed as determined by the vial
inversion test.
Rheology. Dynamic rheological experiments were performed
on an Anton Paar Physica MCR101 rheometer. For the oscilla-
tory shear measurements, a sandblasted parallel top plate with a
50 mm diameter and 1.0 mm gap distance were used. Gels for
rheological experiments were prepared on the bottom plate of the
rheometer by loading a 2.0 mL solution of the gelator immediately
after GdL addition. At this point, the sample is still a free-flowing
liquid. Hence, sample uniformity and reproducibility are high.
Evaporation of water from the hydrogel was minimized by
covering the sides of the plate with low-viscosity mineral oil.
The measurements of the shear modulus (storage modulus G0 and
loss modulus G00) with gelation were made as a function of time at
a frequency of 1.59 Hz (10 rad/s) and at a constant strain of 0.5%
for a period of 24 h. To ensure the measurements were made in the
linear viscoelastic regime, an amplitude sweep was performed.
The results showed no variation in G0 and G00 at a strain of 0.5% as
used above (see Figures S1-S3 for example data, Supporting
Information).
where Γ is the surface concentration, γ is the surface tension, and
C is the concentration of dipeptide-conjugate in solution. The
area/molecule at the air/water interface is given by 1/Γ. The slope
can be obtained from the surface tension measurements plotted

[Assistant] [{"peptide_sequence": "AA ", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Napthalene", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Conjugate", "conjugate_partner": "Napthalene", "solution": "100 H2O", "peptide_concentration(mg/ml)": "5.0", "PH": "3.4", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "AA", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Napthalene", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Conjugate", "conjugate_partner": "Napthalene", "solution": "100 H2O", "peptide_concentration(mg/ml)": "5.0", "PH": "3.2", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "AA", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Napthalene", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Conjugate", "conjugate_partner": "Napthalene", "solution": "100 H2O", "peptide_concentration(mg/ml)": "5.0", "PH": "3.6", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
Self-assembling short peptides can offer an opportunity to make useful
nano-/microstructures that find potential application in drug delivery. We
report here the formation of multivesicular structures from self-assembling
water-soluble synthetic amphiphilic dipeptides containing a glutamic acid
residue at the C-terminus. These vesicular structures are stable over a wide
range of pH (pH 2-12). However, they are sensitive towards calcium ions. This
causes the rupturing of these vesicles. Interestingly, these vesicles can not
only encapsulate an anticancer drug and a fluorescent dye, but also can release
them in the presence of calcium ions. Moreover, these multivesicular structures
have the potential to carry biologically important molecules like cyclic
adenosine monophosphate (cAMP) within the cells keeping their biological
functions intact. A MTT cell-survival assay suggests the almost nontoxic nature
of these vesicles. Thus, these peptide vesicles can be used as biocompatible
delivery vehicles for carrying drugs and other bioactive molecules.

Self-assembling dipeptide-based nontoxic vesicles as carriers for drugs and
other biologically important molecules†
Self-assembling short peptides can offer an opportunity to make useful nano-/microstructures that ﬁnd
potential application in drug delivery. We report here the formation of multivesicular structures from
self-assembling water-soluble synthetic amphiphilic dipeptides containing a glutamic acid residue at the
C-terminus. These vesicular structures are stable over a wide range of pH (pH 2–12). However, they are
sensitive towards calcium ions. This causes the rupturing of these vesicles. Interestingly, these vesicles
can not only encapsulate an anticancer drug and a ﬂuorescent dye, but also can release them in the
presence of calcium ions. Moreover, these multivesicular structures have the potential to carry
biologically important molecules like cyclic adenosine monophosphate (cAMP) within the cells keeping
their biological functions intact. A MTT cell-survival assay suggests the almost nontoxic nature of
these vesicles. Thus, these peptide vesicles can be used as biocompatible delivery vehicles for carrying
drugs and other bioactive molecules.
Molecular self-assembly offers a wonderful strategy to make var-
ious supramolecular structures including nanotubes,1 nanorods,2
nanoﬁbers,3 nanovesicles4 and others.5 Vesicles (enclosed spherical
assemblies) seem to be a fascinating structure, because of their
structural resemblance with primitive biological cells.6 Vesicles
are of technological interest for various applications including
encapsulation and sustained release of drugs,7 chemical sensing,8a–b
biosensing,8c nanoscale reactors9 and enzyme encapsulation.10
Antonietti et al. have reported that the vesicle formation is a two-
step self-assembly process, an amphiphile ﬁrst forms a bilayer,
which then in a second step closes to form a vesicle.7c Previous
reports include that vesicles are obtained from various building
blocks including block copolymers,11 oligonucleotides,12
based molecules,4a,4h–i,7c–d polysaccrides,13 polypeptides11b–e,14 and a
few synthetic organic molecules like calixarenes,15 cyclodextrins,16
and curcubit[6]urils.17 Peptide assemblies are biocompatible, as is
evidenced by their successful applications in biological and medi-
cal ﬁelds including stem cell differentiation, tissue engineering, in
vivo nerve regeneration and others.18 Vesicles obtained by the self-
assembly of the polypeptides are termed as “peptosomes” and they
have been studied extensively by different research groups.11b–e,14
Deming and his coworkers made a signiﬁcant contribution in
constructing vesicles from self-assembling charged amphiphilic
block copolypeptides and they have used these vesicles as an
intracellular delivery system.11b–e Formation of vesicles depends
on various factors including the proper choice of self-assembling
molecular scaffold,4d,4j,19 pH,20 concentration21 and some other
factors.22 However, examples of short peptide-based vesicles are
relatively fewer than polypeptide-based vesicles. Raches and Gazit
have reported that diphenylglycine, the most generic form of an
aromatic dipeptide, diphenylalanine, can form spherical nano-
metric assemblies and these hollow nanospheres have remarkable
stability, like nanotubes formed by diphenylalanine.23 In a recent
report, it is demonstrated that nanovesicles are formed from self-
assembling dipeptides with a conformationally constrained a,b-
dihydrophenylalanine residue (DPhe). These vesicles can encap-
sulate small drugs and other biologically important molecules
including riboﬂavin, vitamin B12, bioactive peptides and even a
protein. They are resistant to the proteolytic enzyme, proteinase
There are many reports of self-assembled biomolecule based
vesicular structures. However, vesicular assemblies which contain
small vesicles within their interior compartments, similar to the
discrete organelles within eukaryotic cells, are very rare in the
literature25 and the structural control of such assembly has not yet
been achieved properly. There is a need for nano- or microvesicles
composed of biological building blocks, which can deliver the
encapsulated molecules at the target site in response to a stimulus
and these vesicles should exhibit minimum or no toxicity, so that
they can be used as good delivery vehicles for carrying drugs
and other biologically important molecules. Peptides 1, 2 and 3
(Fig. 1) have been selected in such a way that each peptide contains
a C-terminally located hydrophilic glutamic acid residue and the
A ﬁeld emission scanning electron microscopic (FE-SEM) study
has been performed to investigate the morphological features of
these self-assembled peptides. Each of these peptides (0.06 mmol)
was dissolved in 1 mL water of pH 6.96 and allowed to age for 24 h.
The FE-SEM study clearly demonstrates (Fig. 2) the formation of
spherical aggregates (nanospheres) having diameters within the
range of 90–100 nm for peptide 1, 90–100 nm for the peptide 2
and 80–90 nm for the peptide 3, respectively.
Transmission electron microscopy (TEM) experiments were
carried out to examine the internal structure of these spherical
aggregates, which were obtained from self-assembling peptides 1,
2 and 3. No clear and credible images with distinct structural
features was obtained in the TEM experiments (Fig. S15, ESI†) at
0.06 mmol mL-1 peptide concentration (i.e. under the conditions
of the FE-SEM experiment).
So, all TEM experiments were performed at a lower con-
centration of peptides (0.02 mmol of each peptide in 1 mL of
water). These TEM images vividly illustrate the architectural
arrangement of multivesicular assemblies for these self-assembled
peptides, in which one or more vesicles have been enclosed within
a relatively larger one (Fig. 3 and Fig. S19, ESI†). These peptide
vesicles are stable within a wide range of pH values (pH 2 to
12). It is interesting to note that the vesicles within the vesicles
exist throughout the experimental conditions of different pH
values (Fig. 3, 4, 5). This is probably due to the fact that a
change of pH (2–12) cannot change the amphiphilic nature of
the molecule and the self-assembly pattern in the formation
of the vesicular structure. In some cases, these assemblies are
similar to the organization of biological eukaryotic cells and
their subcellular organelles (Fig. 3a,b, 4c, 5a,c).25 The sizes of
these vesicles obtained from FE-SEM and TEM experiments are
different as the peptide concentrations that were used for these two
experiments were not the same. TEM experiments were carried
Fig. 3 Transmission electron microscopy (TEM) images of (a) peptide 1,
(b) peptide 2 and (c) peptide 3 at pH 6.96.
N-terminally positioned hydrophobic residue. The reason behind
that is all these amphiphilic peptide molecules (1, 2 and 3) are
anionic at slightly acidic to neutral pH as well as in basic pH
and they are sensitive to bivalent cations. In this study, we present
the formation of multivesicular architectures (vesicles containing
small vesicles within their interior compartments) from the self-
assembling water-soluble synthetic dipeptides containing glutamic
acid as the C-terminal residue. To the best of our knowledge, this
is the ﬁrst report of multivesicular architectures obtained from
water-soluble synthetic anionic dipeptides. In this report, it has
been demonstrated that these peptide-based vesicles are stable
over a wide pH range (pH 2–12), and responsive to Ca+2 ions.
Moreover, a ﬂuorescent dye (eosin B) and an anticancer drug
(doxorubicin) can be encapsulated inside these vesicles and these
encapsulated materials can be released in the presence of calcium
ions. The nontoxic nature of these dye/drug-loaded vesicles has
also been demonstrated using MTT (3,4-dimethylthiazol-2-yl)-
2,5-diphenyltetrazolium bromide) cell survival assay. A cellular
uptake study using vesicles loaded with biologically important
molecules like cyclic adenosine monophosphate (cAMP) demon-
strates that these vesicles not only enter into the cells but also
can successfully deliver the entrapped molecules within the cells
keeping their biological functions intact. These functional peptide-
based vesicles are endowed with fascinating properties and this
opens up an opportunity to use these vesicles as vehicles for
delivering drugs and other biologically important molecules inside
the cytosolic counterpart of cells.

[Assistant] [{"peptide_sequence": "VG", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "10.45", "PH": "6.96", "temperature(celsius)": "37.0", "heating_cooling": "No", "incubation_period(min)": "2160.0"}, {"peptide_sequence": "LG", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "11.29", "PH": "6.96", "temperature(celsius)": "37.0", "heating_cooling": "No", "incubation_period(min)": "2160.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
Supramolecular chemistry enables the creation of a diversity of nanostructures
and materials. Many of these have been explored for applications as biomaterials
and therapeutics. Among them, self-assembling peptides have been broadly
applied. The structural diversity afforded from the library of amino acid
building blocks has enabled control of emergent properties across length-scales.
Here, we report on a family of amphiphilic tripeptides with sequence-controlled
nanostructure. By altering one amino acid in these peptides, we can produce a
diversity of nanostructures with different aspect-ratio and geometry. Peptides
that produce high aspect-ratio structures can physically entangle to form
hydrogels, which support cell viability in culture. Importantly, in comparison
to many other short self-assembling peptide biomaterials, those reported here
form filamentous nanostructures in the absence of typical secondary structures
(i.e., β-sheet). Thus, we have illustrated a facile way to obtain versatile
biomaterials with different nanostructural morphology from short and defined
peptide sequences.

Fig. 3 Live/dead ﬂuorescent imaging of 3T3 cells grown suspended in
tripeptide hydrogels of FVD, FID, and FLD. Live cells are indicated by
green ﬂuorescence (calcein-AM) and dead cells are indicated with red
ﬂuorescence (ethidium homodimer).

[Assistant] [{"peptide_sequence": "FGD", "self-assembly_phase": "none", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "6.75", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FAD", "self-assembly_phase": "none", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "7.03", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FVD", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "7.59", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FID", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "7.87", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FLD", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 H2O", "peptide_concentration(mg/ml)": "7.87", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FVD", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 PBS", "peptide_concentration(mg/ml)": "7.59", "PH": "7.4", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FLD", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 PBS", "peptide_concentration(mg/ml)": "7.87", "PH": "7.4", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "FID", "self-assembly_phase": "hydrogel", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "100 PBS", "peptide_concentration(mg/ml)": "7.87", "PH": "7.4", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
The grafting of a short nucleic acid strand to ditryptophan dipeptide (WW)
results in a peptide-DNA hybrid, which assembles into fibrils under controlled
aggregation conditions as evidenced by label free optical sensing owing to the
intrinsic fluorescence of the dipeptide.

Label-free, optical sensing of the supramolecular
assembly into fibrils of a ditryptophan–DNA
The synthesis of amino acid sequences and hybrids thereof is an
elegant and versatile approach to assemble nanostructures. More-
over, the functional residues along the amino acid chains infer
responsiveness to a variety of physicochemical stimuli. For instance,
sensitivity to ionic strength or temperature might induce size
variation;1 whereas pH changes trigger the release of an encapsu-
lated compound,2 biological stimulus responsiveness induces
morphological transitions.3 Along these lines, the self-assembly
of WW-based hybrids has been reported previously.4 The result-
ing structures were demonstrated to be responsive to various
stimuli such as sunlight,5 the presence of potassium ions4b and
pH variation.4a Of major interest though is the intrinsic fluores-
cence of WW and the hybrids thereof, which could be used to
monitor changes in the solvent polarity6 as well as specific
biotin–avidin and ferrocene–cyclodextrin ligand interactions.7
We recently described the eﬃcient coupling of a short single
stranded synthetic nucleotide sequence to the diphenylalanine
(FF) dipeptide to induce the formation of pH sensitive vesicular
structures.2 In comparison to phenylalanine, tryptophan reveals
a University of Geneva, Faculty of Sciences, Department of Inorganic and Analytical
Chemistry, 1211 Geneva 4, Switzerland. E-mail: corinne.vebert@unige.ch;
Fax: +41 22 379 60 69; Tel: +41 22 379 64 22
b Center for Cellular Imaging and NanoAnalytics (C-CINA), Biozentrum,
University Basel, Mattenstrasse 26, CH-4058 Basel, Switzerland.
E-mail: mohamed.chami@unibas.ch; Fax: +41-61-387 39 86;
Tel: +41-61-387 32 24
c Department of Chemistry, Centre for Environmental Sciences and Engineering,
DST Thematic Unit of Excellence on Soft Nanofabrication, Indian Institute of
Technology Kanpur, 208016 (UP), India. E-mail: sverma@iitk.ac.in;
Tel: +91-512-259-7643
† Electronic supplementary information (ESI) available: Chemical synthesis and
characterization light scattering data and electron microscopy imaging, pH
stability. See DOI: 10.1039/c4cc02631d
‡ The two authors contributed equally to this research work.
Grafting by conventional solid phase synthesis (see ESI†) of
the WW dipeptide to a model linear (50-CTC TCT CTC TTT-30
50-(CT)5T2-30) synthetic single stranded nucleotide sequence
(modified at the 50 end by an amino group) results in an
amphiphilic, rod-coil peptide–DNA hybrid (Scheme 1).
Chemical characterization by HPLC (Fig. S1, ESI†) and ESI-MS
(Fig. S2, ESI†) evidences the formation of the peptide–DNA hybrid in
accordance with its chemical structure. Owing to both chemical and
physical incompatibility between the rigid hydrophobic dipeptide
fragment and the water soluble flexible nucleotide sequence, the
amphiphilic molecule organizes into spherical structures in dilute
aqueous solution (Fig. 1a). Depending on concentration and
incubation time, fibrils of a thickness of 0.5–1 mm and length of
several micrometres developed (Fig. 1b). As could be observed
Fig. 1 Morphological characterization by scanning electron microscopy of
the structures assembled by WW–((CT)5T2): (a) spherical structures assembled
at low concentration (below 1 mM) whereas (b) fibres developed at high
concentration under controlled aggregation (time and temperature).
by transmission and scanning electron microscopy (TEM and SEM),
direct dissolution in aqueous solution induced the spontaneous
formation of spherical structures of about 300 nm size (Fig. 1a and
Fig. S3, ESI†). An apparent hydrodynamic radius (230 nm) of the
spherical structures assembled in the low concentration regime was
determined by dynamic light scattering (Fig. S4, ESI†), which was in
agreement with the size estimated by electron microscopy. The
dipeptide itself is not soluble in pure water but spherical structures
assembled in an aqueous solution of 1,1,1,3,3,3-hexafluoro-2-
propanol (HFIP) (1 : 50 v/v HFIP/H2O ratio) (Fig. 2). Removal of
the organic solvent induces the precipitation of the dipeptide.
Structure formation in the aqueous solution of HFIP occurs
owing to the polar property of the indole ring of tryptophan,
which acts as a quadruple as demonstrated in the formation of
vesicular structures from N-alkylindoles.9
The peptide–DNA hybrid might therefore form the observed
fibrils by both p–p stacking and hydrogen bonding according to
a nucleation polymerization process comparable to the one that
induces the formation of polypeptide fibrils.11 The 230 nm size
spherical structures would then act as seeds for this mecha-
nism to take place. As described by Hamley, polypeptide fibril
formation occurs by the interaction between the common main
chain polypeptide backbone whereas folding is due to specific
interactions of the side chains.11 At low concentration, spherical
structures formed by WW–DNA is induced by the chemical and
physical incompatibility between the dipeptide and nucleic acid
strands and stabilized by hydrogen bonding between the nucleo-
tide sequences engaged in structure formation. These spherical
structures at higher concentrations and under controlled aggre-
gation (time and temperature) might develop into fibres by
nucleation dependent polymerization.
We could thus resort to label-free fluorescence spectroscopy
amenable by the WW dipeptide intrinsic fluorescence to evidence
this mechanism of structure formation and the putative role of the
interactions involved. This peculiar property of WW is sensitive to
the polarity of the environment. Since the supramolecular struc-
tures are held by non-covalent interactions, they disassemble below
a critical aggregation concentration (cac), which was quantified by
fluorescence spectroscopy (Fig. 3). Fig. 4 shows the corresponding
concentration dependent fluorescence spectra. The dilution course
of the fluorescence intensity shows that below 0.1 mM the struc-
tures started to disassemble as further evidenced by SEM (Fig. S5,
ESI†). The role of hydrogen bonding in stabilization of the spherical
structures was further supported by incubation with a chaotropic
agent, urea, which also induced their disassembly as imaged by
SEM (data not shown) and fluorescence spectroscopy (Fig. S6, ESI†).
These experimental results demonstrate that the spherical struc-
tures are stabilized by hydrogen bonding, which occurs between the
nucleotide sequences engaged in structure formation. This was
further supported by the disassembly of the spherical struc-
tures by the addition of synthetic nucleic acid strands (either
complementary or non-complementary) in the surrounding
medium (Fig. S7, ESI†).
Fig. 2 Spherical structures of WW dipeptide in aqueous solution of HFIP
as observed by (a) scanning electron microscopy and (b) transmission
electron microscopy.
Fig. 4 Fluorescence emission spectra for decreasing concentrations of the
WW–DNA hybrid. (a) 2 mM, (b) 1 mM, (c) 0.5 mM, (d) 0.25 mM, (e) 0.13 mM,
and (f) 0.06 mM.
sequences in the medium (Fig. S8, ESI†) aﬀects the organiza-
tion into spherical structures of the WW–DNA. With increasing
concentration of the nucleic acid strands, disassembly occurs
(Fig. S9, ESI†). Moreover, as can be observed in Fig. 4, the
fluorescence intensity maximum (lmax at 370 nm) increases with
increasing concentration and the maximum is reached at a concen-
tration of 0.5 mM. On further increasing the concentration (1 mM),
the intensity of this peak decreases and a red-shifted signal appears
at 476 nm. On further increasing the concentration to 2 mM, this
feature becomes more prominent. This red shifted peak is an
indication of the presence of an extensive J-type aggregate, which
results from the p–p stacking of aromatic rings. These types of
aggregates were indeed observed previously in other systems.12 This
is further evidenced by SEM (Fig. 1). WW–DNA hybrids organize
into spherical structures at concentrations between 0.05 mM and
0.5 mM. On further increasing the concentration (1 mM), they self-
assemble to long fibres. These experimental results indicate that
structures formed at high concentrations are supported by the
interaction of the aromatic WW dipeptide in agreement with reports
published previously by Reches and Gazit13 as well as Verma and
coworkers.4a We unfortunately could not resort to CD spectroscopy
to evidence further p–p stacking. Both the nucleic acid strand and
WW–DNA hybrid induce a negative shift at 218 nm (Fig. S10, ESI†).
In addition to p–p stacking of WW residues, the nucleic acid
strands might also undergo hydrogen bonding in the fibrillar
structure. To assess the role of these interactions, we grafted the
WW dipeptide to a water soluble polymer, poly(ethylene glycol)
(PEG, MW = 2000). Unlike a nucleotide sequence, PEG does not
engage in specific hydrogen bonding. Long needle shaped fibres of
about 50 nm width were observed by TEM (Fig. S11, ESI†). Unlike
WW–DNA, no spheres were assembled by WW–PEG at low concen-
tration. Similar results were previously reported for PEG-FFFF.12a
The authors associated the finite length of the PEG–tetrapeptide
hybrid with the end cap energy due to the hydrophobicity of the
peptide. WW–PEG thus assembles through b-sheet formation at
high concentration due to the p–p stacking of the amino acids
which is in agreement with J-type aggregation, as observed for the
WW–DNA hybrids through fluorescence spectroscopy (Fig. 4).
It further shows that hydrogen bonding plays a key role in the
formation of the long WW–DNA fibres. Besides, it has to be
emphasized that fibrils could not be assembled by the WW–DNA
hybrid in presence of the complementary nucleotide sequence of
the one involved in structure formation (data not shown). More-
over, the fibres could not be disassembled by the addition of the
complementary sequence subsequent to fibril formation which
strongly supports the involvement of hydrogen bonding in WW–
DNA fibrils formation in support of p–p stacking. The WW–DNA
hybrid organizes according to a mechanism analogous to that of
the nucleation polymerization of polypeptide fibrils as further
evidenced by the monitoring of the time course of the dipeptide
fluorescence (Fig. 5). The peak assigned to the formation of J-type
aggregates appears after 6 hours of incubation.
In conclusion, we report here on the successful coupling of
the ditryptophan dipeptide to a synthetic nucleotide sequence.
The resulting hybrid forms spherical structures in dilute aqueous
solution. At high concentration and under controlled aggregation
Fig. 5 Time course of the fluorescence intensity monitored upon assembly
of WW–DNA (0.5 mM) at time intervals 0 (t0), 6 (t6) and 24 hours (t24).
(time and temperature), the formation of fibrils has been
observed. Of high interest is the intrinsic fluorescence of the
dipeptide which enables the quantification of a critical aggregation
concentration and the indirect evidence in a label-free optical
mode that the WW–DNA hybrid organization is analogous to that
of the nucleation polymerization of polypeptide fibrils, i.e. sup-
ported by both main chains p–p stacking of the dipeptide and side
chains hydrogen bonding between the nucleic acid strands. This is
supported by concentration and time dependent fluorescence
spectroscopy and monitoring of the fluorescence spectra of the
structures assembled by the WW–DNA hybrid in presence of either
the complementary or the non-complementary sequence of the
nucleic acid strands involved in the self-assembly.

[Assistant] [{"peptide_sequence": "WW", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "0.35", "PH": "7.5", "temperature(celsius)": "37.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}, {"peptide_sequence": "WW", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "0.31", "PH": "7.5", "temperature(celsius)": "37.0", "heating_cooling": "No", "incubation_period(min)": "1440.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
Copyright © 2014 European Peptide Society and John Wiley & Sons, Ltd.

FF-peptide nanotubes (PNT) and LL–PNT were prepared by
dissolving the L-diphenylalanine peptide (Bachem, Bubendorf,
Switzerland) and L-dileucine peptide (Bachem, Bubendorf, Switzerland)
in 1,1,1,3,3,3-hexaﬂuoro-2-propanol (Sigma-Aldrich, Rehovot, Israel) at
an initial concentration of 100 mg/ml, respectively, and then
further diluted in deionized water.
For environmental
scanning electron microscope (ESEM)
measurements, drops of the above solutions at a ﬁnal concentra-
tion of 2 mg/ml were placed onto clean silicon substrates and dried
at room temperature. The thermally induced LL PNT samples were
heated in an oven for 90 min at 180 °C. The heated and native PNT
samples were coated with palladium–gold and scanned using a
JSM JEO 6300 scanning electron microscope operating at
10–15 kV. For FF PNT, drops of the above solutions at a ﬁnal
concentration of 2 mg/ml were placed onto clean silicon sub-
strates and dried at room temperature. The thermally induced
PNT samples were heated in an oven for 180 min at 180 °C.
The PL spectra of the FF and LL nanotubes were measured by
different techniques: The LL nanotubes were self-assembled in
water (at a concentration of 20 mg/ml) and placed onto glass
cover slips and dried at room temperature. After heating for 1 h
for each temperature step, the LL nanotubes were inserted into
a quartz cuvette ﬁlled with deionized water. The PL of the LL
nanotubes that were immersed into aqueous solution produces
a Raman peak that is associated with water. As LL nanotubes,
the FF nanotubes were self-assembled in water (at concentration
of 20 mg/ml) and placed onto a quartz plate and dried at room
temperature. After heating for 1 h for each temperature step,
the PL spectra of the solid-state FF-PNTs sample were measured.
The PL was measured by Horiba Jobin Yvon FL3-11 Spectroﬂuo-
rometer, with a 2-nm slit.
For CD measurements, a solution with a ﬁnal concentration of
12.6 mg/ml was made. Drops containing 200 μl of the above
solutions were placed onto glass cover slips and dried at room
temperature. The dried samples were heated 1 h for each tem-
perature step,and then were re-dissolved in 5 ml deionized water.
One milliliter from the new solution was then inserted into a
quartz cuvette. CD spectra were obtained by Chirascan CD Spec-
trometer (Applied Photophysics, UK). The wavelengths were
scanned every 4 s, from 180 to 250 nm.
For ESEM measurements, drops of the above solutions at a ﬁnal
concentration of 2 mg/ml were placed onto clean silicon
substrates and dried at room temperature. The thermally induced
PNT samples were heated in an oven for 90 min at 180 °C. All
samples were measured by PHI Model 2100 TRIFT II system.
Figure 1. ESEM images of (a) native FF-PNT and (b) native LL-PNT. The
cross-sections of both hollow tubes have pronounced regular shapes,
allowing to relate them to deﬁnite crystallographic classes, where FF-PNT
possess nanocrystalline hexagonal symmetry and LL-PNT possess orthorhom-
bic symmetry (inset images of individual FF and LL nanotubes were taken from
Ref. [17]) - Reproduced by permission of the PCCP Owner Societies, http://
pubs.rsc.org/en/Content/ArticleLanding/2012/CP/c2cp40157f#!divAbstract
The insets at Figure 1 show the individual native FF and LL
nanotubes. The cross-sections of the tubes have pronounced reg-
ular shapes, allowing to relate them to deﬁnite crystallographic
classes, where FF-PNT have nanocrystalline hexagonal symmetry
P61 [17–19] and LL-PNT possesses orthorhombic symmetry of
P212121 [20,21]. However, despite of similar hydrophobic origin,
leading to wrapping of FF and LL peptides into similar native
nanotubes, at this initial phase, crystallization of both dipeptides
goes at the nanoscale into different space group structures. We
assume that the self-assembly process of FF-native PNT and LL-
native PNT is determined by their various molecular recognition
mechanisms, aromatic and non-aromatic. Both bioorganic nano-
structures are related to the structures without center of symme-
try, which is the basis for observation of a wide spectrum of
ferroelectric and related phenomena, such as piezoelectric effect,
electrical spontaneous polarization [18,22] and second order
nonlinear response [12].
We found that native FF and LL nanotubes undergo irreversible
phase transition. In the same temperature region (T = 140–180 °
C), both FF-PNT and LL-PNT change drastically their nanotube
morphology and adopt a shape of ultrathin ﬁber-like nanostruc-
ture, as evident from Figure 2. FF nanotubes demonstrate gradual
growing needle-like thin wire-ﬁbrils. Our previous data [17,19]
showed for a single FF nanotube that heated open-end FF-PNT
is gradually close its central hole and transformed into ﬁber or
into a bundle of ﬁbers. The same transformation is also observed
in LL nanotubes when they split into very thin nanoﬁbrils. Found
similar morphological reconﬁrmation into nanoﬁbrils in both FF-PNT
and LL-PNT can be classiﬁed as reconstructive phase transition [17].
We revealed that both newly assembled FF and LL ﬁbrillar structures
during heating (Figure 2) changed their native asymmetric crystal
structure into centrosymmetric. These results are based on our
previous investigations, which showed that the piezoelectric and
nonlinear optical properties of FF-PNT and LL-PNT disappear after
heating at temperature above 120–140 °C [17].
Figure 2. ESEM images of (a) heated FF-PNF to 180 °C and (b) heated LL-
PNF to180 °C. FF nanotubes demonstrate gradual growing needle-like
thin wire-ﬁbrils. The same transformation is also observed in LL nano-
structures when they split into very thin nanoﬁbrils.
Figure 4. CD spectra of (a) FF-PNT and (b) LL-PNT during heating. CD
spectra show that as the temperature increases, the sign of the ellipticity
of FF and LL nanotubes changes and becomes negative, thus suggesting
conformation in both
nanostructures.
Figure 3. ToF-SIMS of (a) native FF-PNT, (b) heated FF-PNF, (c) native LL-PNT,
and (d) heated LL PNF. The molecular weight of both FF and LL was reduced
after heating by 18 g/mol, corresponding to water loss during linear-to-cyclic
molecular transformation. (ToF-SIMS data was taken from Ref. [17] - Reproduced
by permission of the PCCP Owner Societies, http://pubs.rsc.org/en/Content/
ArticleLanding/2012/CP/c2cp40157f#!divAbstract)
the 225–235 nm regions arises from the contributions of aromatic
residues in phenylalanine [32]. Previous studies that demonstrated
positive CD spectrum of FF-based nanostructures ascribed them to
a β-turn conformation [28,30]. The negative CD spectrum recorded
in the range of 210–220 nm was observed in our thermally induced
FF-PNF after high temperature treatment (Figure 4a) and is
indicated on a typical β-sheet structure signature [24].
To the best of our knowledge, the origin of the positive CD
band of native LL-PNT at room temperature and its relation to
any secondary structure was not studied. However, the negative
CD spectrum that was recorded in the thermally induced LL
nanoﬁbrils at high temperature is similar to the CD spectra of
other aliphatic [33,34] and aromatic peptides [28] and can be re-
lated to the β-sheet arrangements of the LL dipeptide molecules.
The formation of a β-sheet structure of the LL molecules is in
agreement with the tendency of proteins with high content of
leucine amino acid to form α/β arrangement [35].
The deep variation of the CD spectra of FF and LL nanostruc-
tures during heating indicates that the secondary arrangement
of FF and LL nanoﬁbrils strongly depends on temperature. Envi-
ronmental conditions can affect the arrangements of molecules,
which are expressed in different CD bands of the same molecule
[28,36]. Heat treatment could also alter the CD bands [37] and even
reverse the samples’ chirality [38]. In our case, thermal treatment of
FF-PNT and LL-PNT induces a β-sheet conformation, accompanied
with a change in the crystal structure symmetry [17]. Thus, although
native FF-PNT and LL-PNT are self-assembled by dissimilar
intermolecular interactions because of their molecular composition
of different origin, both dipeptide nanostructures rearrange into
a similar β-sheet secondary structure, accompanied by similar ﬁ-
bril morphology. It should also be noted that the morphology
and β-sheet secondary structure of both aromatic and aliphatic
peptides are similar to amyloid nanoﬁbrils [5,39].
In contrast to the crystal-related physical properties of FF-PNT
and LL-PNT (piezoelectric and second order nonlinear optical re-
sponse), linear optical properties (such as optical absorption and
PL) are deﬁned by the electronic structure of the self-assembled
peptide nanotubular structures. In our recent studies [7,17], we
found an emission peak at 290 nm from the aromatic FF-PNT,
excited at 265 nm. We found that this PL peak does not change
with temperature, and we believe that it stems from the aromatic
residues of FF molecules [40]. Recent publications showed that
the addition or loss of water molecules affects the FF molecule’s
atomic vibrations and thus inﬂuences their optical characteristics
only in the UV region [41]. Aliphatic native LL-PNT does not pos-
sess any PL properties at all and, as we found in this work, does
not exhibit any peak in the optical absorption.
During thermal treatment, both FF and LL native nanotubes
change their morphology into a ﬁber-like structure (Figure 2).
This morphological transition is accompanied by deep transfor-
mation of their optical properties. As the temperature increases
to T ~120–140 °C, a blue PL peak in the region 420–460 nm ap-
pears both in FF and LL nanostructures and gradually grows until
T ~180 °C (depicted in Figure 5). The narrow sharp peak observed
in the LL ﬁber structure, located at λ = 411 nm, does not depend
on temperature and is associated with the Raman scattering in
water, where its peak position is located exactly at wavenumber
3600 cm(cid:1)1 lower than the excitation wavenumber [42]. Because
the sample was measured within an aqueous solution, a water-
mark was expected.
Several models have been proposed to explain the origin of
the blue light photon emission from the FF-PNF during heating,
such as quantum conﬁnement
[8] and cyclization process
[19,43]. However, we can conclude that blue PL peak observed
in FF and LL ultrashort peptide nanoﬁbrils can be ascribed to
their similar secondary peptide nanostructures and most likely
is related to a β-sheet secondary structure as was found by us
in this study (Figure 4). This phenomenon should be also valid
to other peptides, which are natively rich in β-sheets, without
the need of thermal treatment. Recently, it was observed that
Fmoc-FF hydrogels, which possess β-sheet conformation, exhibit
a blue PL peak in 460 nm without any thermal treatment [27].
These ﬁndings support our assumption about the origin of blue
PL peak in our dipeptide FF and LL nanoﬁbrils.
As we noted before, the same blue PL peak was recently found
in amyloid ﬁbrils having similar nanoﬁbrilar structure [5,6]. Origi-
nal amyloid ﬁbrils are known to have a β-sheet structure that is
stabilized by hydrogen bonds between the components of the ﬁ-
ber. Such hydrogen bonds are also formed in the thermally
Figure 5. PL and PLE spectra in the blue-visible spectral region of (a) FF-PNF
and (b) LL-PNF during heating. As the temperature increases to T ~120–140 °C,
a blue PL peak in the region 420–460 nm appears both in the FF and LL
nanostructures and gradually grows until T ~180 °C. (The peak at 411 nm that
appears in the PL spectra of LL nanotubes is associated with Raman peak as-
cribed to water because of the experimental conditions).
Recently, several studies reported on the observation of a blue
light emission in large crystalline proteins with lack of aromatic-
ity, such as gamma-II crystalline [47], diseased-related human
peptides, lysozyme [5] and other amyloid-like ﬁbrils formed by
peptides [48,49]. The common feature to all these ﬁbrils was that
they are all rich in β-sheets secondary structures [5]. As such, the
most accede speculation to the appearance of the blue visible
ﬂuorescence is that hydrogen bonds in β-sheet structure can
cause low energy electronic transitions [5,47], and hence the
emission of a low-wavelength light in the blue visible region is
possible. In addition, it was suggested that the amyloid state of

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[System] polypeptide self-assembly literature information extraction

[User]
The diphenylalanine peptide, the core recognition motif of the beta-amyloid
polypeptide, efficiently self-assembles into discrete, well-ordered nanotubes.
Here, we describe the notable thermal and chemical stability of these tubular
structures both in aqueous solution and under dry conditions. Scanning and
transmission electron microscopy (SEM and TEM) as well as atomic force
microscopy (AFM) revealed the stability of the nanotubes in aqueous solution at
temperatures above the boiling point of water upon autoclave treatment. The
nanotubes preserved their secondary structure at temperatures up to 90 degrees
C, as shown by circular dichroism (CD) spectra. Cold field emission gun (CFEG)
high-resolution scanning electron microscope (HRSEM) and thermogravimetric
analysis (TGA) of the peptide nanotubes after dry heat revealed durability at
higher temperature. It was shown that the thermal stability of diphenylalanine
peptide nanotubes is significantly higher than that of a nonassembling
dipeptide, dialanine. In addition to thermal stability, the peptide nanotubes
were chemically stable in organic solvents such as ethanol, methanol,
2-propanol, acetone, and acetonitrile, as shown by SEM analysis. Moreover, the
acetone environment enabled AFM imaging of the nanotubes in solution. The
significant thermal and chemical stability of the peptide nanotubes demonstrated
here points toward their possible use in conventional microelectronic and
microelectromechanics processes and fabrication into functional
nanotechnological devices.

Preparation of Initial Solutions of Peptides. Peptides were
purchased from Bachem (Bubendorf, Switzerland). Fresh stock
solutions were prepared by dissolving the lyophilized form of the
peptides in 1,1,1,3,3,3-hexafluoro-2-propanol (Sigma Aldrich) at a
concentration of 100 mg/mL. To avoid any preaggregation, fresh
stock solutions were prepared for each experiment.
Transmission electron microscopy (TEM) samples were viewed
using a JEOL 1200EX electron microscope (JEOL, Tokyo, Japan)
operating at 80 kV. Scanning electron microscopy (SEM) images
were prepared using a JSM JEOL 6300 SEM (JEOL, Tokyo, Japan)
operating at 5 kV. A Cold field emission gun (CFEG) high-resolution
scanning electron microscope (HRSEM) (JEOL JSM-6700 field
emission scanning electron microscope equipped with a cold field
emission gun operating at 1 kV (JEOL, Tokyo, Japan) was used to
view samples prepared under dry heating conditions. CD spectra at
the far-UV region (200-250 nm) were recorded with an AVIV 202
spectropolarimeter (Aviv Instruments, Lakewood, NJ) equipped with
a temperature-controlled cell using a cell of path length 0.5 cm; the
bandwidth was 1 nm, and the averaging time was 4s for each
measurement. TGA measurements were recorded on a TGA 2950
at a heating rate of 10 (cid:176) C/min.
Thermal Stability of Autoclaved Nanotubes. Peptide stock
solution was diluted in double distilled (dd) H2O to a final
concentration of 2 mg/mL, autoclaved at 121 (cid:176) C, 1.2 atm, and aliquots
(10 (cid:237)L) were prepared for further analysis. For TEM, an aliquot (10
(cid:237)L) of the autoclaved peptide nanotubes solution was placed onto
a 400 mesh copper grid (SPI Supplies, West Chester, PA) covered
by carbon stabilized Formvar film. Excess fluid was removed after
1 min, and the grid was negatively stained with 2% uranyl acetate
in water. After 2 min, excess fluid was again removed from the grid
and the samples were viewed. For SEM, an aliquot (10 (cid:237)L) of the
autoclaved peptide nanotubes solution was dried at room temperature
on a microscope glass cover slip and viewed after coating with gold.
For AFM, an aliquot (10 (cid:237)L) of the autoclaved peptide nanotubes
solution was diluted to a final concentration of 0.5 mg/mL and then
dropped onto a freshly cleaved mica and dried at room temperature.
All imaging was performed in air in tapping mode using a “J” type
scanner with heating stage using silicon nitride tapping probes
(supplied by Veeco Metrology Group) mounted on cantilevers with
spring constants of 34.4-37.2 N/m and a resonant frequency of
280-360 kHz. For imaging in solvent, a “J” type scanner was used
in tapping mode using NP-sharpened silicon nitride tapping probes
(supplied by Veeco Metrology Group) mounted on cantilevers with
spring constants of 0.032 N/m.
For CD spectra, peptide stock solution was diluted in double
distilled (dd) H2O to a final concentration of 2 mg/mL and then
immediately diluted to a final concentration of 0.04 mg/mL. The CD
spectra of peptide nanotubes that self-assembled from the dipeptides
NH2-(L)Phe-(L)Phe-COOH or NH2-(D)Phe-(D)Phe-COOH were
recorded at 25, 37, 60, 70, 80, and 90 (cid:176) C. A control dipeptide that
does not self-assemble, NH2-(L)Ala-(L)Ala-COOH, was incubated
under the same conditions, and its CD spectra at 25 (cid:176) C was recorded.
Thermal Stability of the Nanotubes in Heated Aqueous
Solution. Peptide stock solution was diluted in double distilled (dd)
H2O to a final concentration of 2 mg/mL, incubated in a water bath
at 80 (cid:176) C for 1 h, and aliquots (10 (cid:237)L) were prepared for further
analysis. For TEM, an aliquot (10 (cid:237)L) of the incubated peptide
nanotube solution was placed onto a 400 mesh copper grid (SPI
Supplies, West Chester, PA) covered by carbon-stabilized Formvar
film. Excess fluid was removed after 1 min, and the grid was
negatively stained with 2% uranyl acetate in water. After 2 min,
excess fluid was again removed from the grid and the samples were
viewed. For SEM, an aliquot (10 (cid:237)L) of the incubated peptide
nanotubes solution was dried at room temperature on a microscope
glass cover slip and viewed after coating with gold.
Thermal Stability under Dry Conditions. The peptide stock
solution was first diluted to 2 mg/mL and then immediately diluted
to a final concentration of 0.5 mg/mL. An aliquot (10 (cid:237)L) was dried
on freshly cleaved mica, under vacuum in a desiccator at room
temperature and then heated for 10 min at the required temperature.
The samples were viewed using CFEG HRSEM. For TGA, the
diluted peptide nanotubes solution (2 mg/mL) was incubated for 5
h at room temperature and then lyophilized.
Figure 1. TEM micrographs of autoclaved nanotubes (121 (cid:176) C, 1.2 atm). Untreated control (a-c) and autoclaved (d-f) samples of peptide
nanotubes viewed at magnifications of (cid:2)1000 (a and d), (cid:2)10 000 (b and e), or (cid:2)50 000 (c and f).
Figure 2. SEM micrographs of autoclaved peptide nanotubes. Untreated control (a-c) and autoclaved (d-f) samples of peptide nanotubes
viewed at magnifications of (cid:2)1000 (a and d), (cid:2)10 000 (b and e), or (cid:2)50 000 (c and f).
Conformational Stability in Organic Solvent. An aliquot (10
(cid:237)L) of the diluted nanotubes solution (2 mg/mL) was air-dried on
freshly cleaved mica, placed for 10 min in acetone or acetonitrile,
and then analyzed by SEM. In addition, 50 (cid:237)L of the peptide nanotubes
solutions were mixed with 50 (cid:237)L of each of the following solvents:
ethanol, methanol, 2-propanol, acetone, and acetonitrile and incubated
Figure 3. AFM images of autoclaved peptide nanotubes. (a)
Topography image (5 (cid:237)m (cid:2) 5 (cid:237)m) of untreated peptide nanotubes.
(b) Line plot of height across the axis of the nanotubes seen in Figure
3a. (c) Topography image (5 (cid:237)m (cid:2) 5 (cid:237)m) of autoclaved peptide
nanotubes. (d) Line plot of height across the axis of the nanotubes
seen in Figure 3c.
at room temperature for half an hour. For SEM analysis, an aliquot
(10 (cid:237)L) of the incubated peptide nanotube solution was air-dried on
a microscope glass cover slip and viewed after coating with gold.
Imaging the Peptide Nanotubes in Aqueous Environment.
For AFM analysis, the diluted peptide nanotubes solution (2 mg/
mL) was diluted in acetone to a final acetone concentration of 50%.
The samples were dropped onto freshly cleaved mica and imaged.

[Assistant] [{"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}, {"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "37.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}, {"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "60.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}, {"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "70.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}, {"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "80.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}, {"peptide_sequence": "FF", "self-assembly_phase": "tube", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "2/98 HFP/H2O", "peptide_concentration(mg/ml)": "2.0", "PH": "5.2", "temperature(celsius)": "90.0", "heating_cooling": "No", "incubation_period(min)": "5.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
Copyright © 2011 Elsevier Inc. All rights reserved.

L-phenylalanine
glucono-d-lactone, ﬂuorescein
tert-butylester hydrochloride,
isothiocyanate isomer I (FITC), isobutyl chloroformate and 4-meth-
ylmorpholine were supplied from Aldrich. Triﬂuoroacetic acid was
obtained from Fluka.
Fmoc–Phe–Phe was prepared as described previously [12].
To prepare the cryo- or hydrogels, Fmoc–Phe–Phe was sus-
pended in Milli-Q-puriﬁed water and sonicated. NaOH (0.5 M)
was added to the suspension until a solution was formed. The ﬁnal
concentration of Fmoc–Phe–Phe was between 2 and 20 mM for
this study. For samples prepared with salt 0.1 M K2SO4 were added.
To induce gelation, GdL was added as a powder to a concentration
of 4–40 mM. The sample was quickly mixed and either frozen at
(cid:2)12 (cid:3)C (cryogel) or stored at room temperature (hydrogel) for
72 h in both cases.
As an example for the preparation of the 15 mM Fmoc–Phe–Phe
sample, 4 ml of 15 mM solution of Fmoc–Phe–Phe was prepared to
which 22 mg of gluconolactone was added by mixing, and then
0.5 ml of the solution was quickly added into each glass tube
(diameter 7 mm) that was closed at the bottom with a silicon cap.
Cryogels of Fmoc–Phe–Phe for scanning electron microscopy
were freeze-dried in order to remove the water, and dried samples
were cut into thin discs and sputtered with gold/palladium (40/
60). The sputtered samples were then examined using a JEOL
JSM-5000LV scanning electron microscope.
TEM was utilised to study the formed ﬁbres of the dipeptides. A
piece of the gel was placed on a copper grid, and the gel was stained
using a 1% uranyl acetate solution for 60 s, after which the excess li-
quid was removed and the sample dried. The prepared samples were
examined using a JEOL 1230 transmission microscope.
Gels for confocal laser scanning microscopy (CLSM) were pre-
pared as described above except that 60 ll of FITC (1 mg/ml in
DMSO) was added to the sample which was then carefully mixed.
The prepared gels were carefully washed with an aqueous solution
with pH 3 to remove any non-bound dye. The gels were then cut into
thin discs and visualised using a Zeiss LSM 510 META microscope,
excitation wavelength 488 nm and emission wavelength 530 nm.
Stress sweeps were performed using a StressTech rheometer
with serrated parallel plates. Gels with a thickness of 3 mm were
used and compressed to 1.5 mm during the experiment. The stress
sweeps were performed between 0.008 and 10 Pa at 1 Hz.
Gels for FTIR were prepared in deuterated water and recorded
on a Burker-ALPHA-P using attenuated total reﬂectance. Spectra
were acquired in the 4000–400 cm(cid:2)1 range with a resolution of
4 cm(cid:2)1 over 128 scans, and a deuterated water spectrum was used
as a background and subtracted from all spectra.

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[System] polypeptide self-assembly literature information extraction

[User]
Self-assembling dipeptides conjugated to naphthalene show considerable promise
as nanomaterial structures, biomaterials, and drug delivery devices. Biomaterial
infections are responsible for high rates of patient mortality and morbidity.
The presence of biofilm bacteria, which thrive on implant surfaces, are a huge
burden on healthcare budgets, as they are highly resistant to current
therapeutic strategies. Ultrashort cationic self-assembled peptides represent a
highly innovative and cost-effective strategy to form antibacterial
nanomaterials. Lysine conjugated variants display the greatest potency with 2%
w/v NapFFKK hydrogels significantly reducing the viable Staphylococcus
epidermidis biofilm by 94%. Reducing the size of the R-group methylene chain on
cationic moieties resulted in reduction of antibiofilm activity. The primary
amine of the protruding R-group tail may not be as readily available to interact
with negatively charged bacterial membranes. Cryo-SEM, FTIR, CD spectroscopy,
and oscillatory rheology provided evidence of supramolecular hydrogel formation
at physiological pH (pH 7.4). Cytotoxicity assays against murine fibroblast
(NCTC 929) cell lines confirmed the gels possessed reduced cytotoxicity relative
to bacterial cells, with limited hemolysis upon exposure to equine erythrocytes.
The results presented in this paper highlight the significant potential of
ultrashort cationic naphthalene peptides as future biomaterials.

Materials. Fmoc and Boc protected amino acids and Wang resin
(mesh size 100−200, 0.65 mmol/g) were obtained from Novabio-
chem, Merck KGaA (Darmstadt, Germany). Cell viability reagent
alamarBlue was purchased from AbD Serotec (Oxford, U.K.). 37%
Hydrochloric acid (HCl), acetonitrile (HPLC grade, ≥99.93%), 2-
naphthaleneacetic acid (>99% purity), Whatman pH indicator paper
(pH 1−14), and hydroxyl propyl methyl cellulose (HPMC) were
obtained from Sigma-Aldrich (Dorset, U.K.). Sterile Nunc 96-well
microtiter plates and sodium hydroxide (NaOH) ≥99.0% pellets were
purchased from VWR International (Leicestershire, U.K.). NCTC
Peptide Synthesis and Characterization. All peptides (Figure
1, NapFF, NapFFKK, NapFFFKK, NapFFOO, NapFFK′K′) were
synthesized following standard Fmoc solid phase protocols on Wang
resin, using a manual nitrogen bubbler apparatus. Peptide elongation
was eﬀected using standard HBTU coupling chemistry in
dimethylformamide (DMF) solvent with 4-fold molar excess of
diisopropyl ethylamine (DIEA) in DMF and a 3-fold molar excess of
each Fmoc-protected amino acid or naphthalene acetic acid. Boc-
Lysine(Fmoc)−OH was used to synthesize the epsilon (ε) variant
NapFFK′K′. All peptides were cleaved from the resin,
carboxylic acid terminated peptides, using 95% triﬂuoroacetic acid
(TFA), 2.5% triisopropylsilane and 2.5% thioanisole (3 h, room
temperature), and precipitated using cold (−20 °C) diethyl ether. The
identity of each peptide was conﬁrmed by electrospray mass
spectroscopy (Thermo Finnigan LCQ Deca ion trap) and 1H Nuclear
Magnetic Resonance (1H NMR) analysis (Varian Unity Inova 400
spectrometer) in C2D6OS (d6-DMSO) (Supporting Information
Figures S1−S5). Peptide purity was analyzed by RP-HPLC using an
Agilent 1260 Inﬁnity system, ﬁtted with a Gemini C18, 250 mm × 4.6
mm column, a 2−60% acetonitrile gradient [30 min] in 0.05% TFA-
water at a ﬂow rate of 1.5 mL/min. All peptides were found to have
>95% purity.
Peptide Gelation. Peptides were prepared and tested for gelation
characteristics at concentrations of 2.0, 1.5, 1.0 and 0.5% w/v as
previously outlined.13 Peptides were initially suspended in 200 μL
sterile deionized water. All peptides possessed a terminal carboxylic
acid moiety. Therefore, increasing the pH of the solution to pH 9, via
titration with 1 M NaOH, resulted in deprotonation of the terminal
species and full dissolution. Protonation of the terminal carboxylic acid
via subsequent titration with 0.5 M HCl to pH 6−7 resulted in the
formation of a homogeneous gel at concentrations above the
minimum gelation concentration for the peptide. The pH was veriﬁed
using Whatman pH paper. Gels were allowed to develop for 24 h. Self-
assembly and the formation of a self-supporting gel was veriﬁed
through the use of an inversion assay, as previously described.22 Gels
and solutions were diﬀerentiated based on ﬂow characteristics with
solutions demonstrating ﬂow properties and gels remaining suspended.
Cryo-Scanning Electron Microscopy (Cryo-SEM). Samples
were freshly prepared via pH induction as outlined and analyzed for
3D ﬁbrous morphologies. Cryo-SEM images were collected by high
vacuum freeze-drying using a Hitachi Analytical Benchtop SEM
TM3030 (Hitachi High-Technologies Europe, Berkshire, U.K.) with
Deben Cool Stage (Deben, Suﬀolk, U.K.), using a method similar to
that of Pogorelov and colleagues.23 Vacuum lyophilization was applied
at 2 °C and images obtained at −10 °C, over a range of magniﬁcations
(15−30 000×) and 15 kV accelerating voltage.
Transmission Electron Microscopy (TEM). TEM was performed
using a FEI Morgagni 268 transmission electron microscope (FEI
electronics, Burlington, Massachusetts, U.S.A.) and a negative staining
technique.24 400 mesh copper grids were coated with continuous thick
carbon ﬁlm (∼35 nm) having been previously glow discharged to
increase their hydrophilicity. Samples (4 μL) were placed on the grid
and rinsed three times with doubled distilled water. The grid was
stained three times by a 2% w/v solution of uranyl acetate. Excess stain
solution from the edge of the grid was removed using ﬁlter paper.
Grids were allowed to air-dry before immediate examination.
Fourier Transform Infrared Spectroscopy (FTIR). Hydrogel
samples (2−0.5% w/v) were prepared as described previously but with
D2O replacing water.25 Prepared D2O containing hydrogels were
sandwiched between two CaF2 25 mm2 discs (0.05 mm spacer). FTIR
spectra were obtained using a Jasco 4000 series FTIR spectrometer
(Jasco Inc. Tokyo, Japan), between the wavelengths 4000−400 cm−1
with resolution 2 cm−1 and an average of 128 scans. The spectrum of
deuterated water was used as a background and subtracted from all
Circular Dichroism (CD). CD was performed using a Jasco J815
Spectropolarimeter and quartz cuvette 1 mm path length, with 1 s
integrations, 1 accumulation, and a step size of 1 nm with a bandwidth
of 1 nm over a range of wavelengths from 180 to 300 nm at 25 °C.
Peptide samples were prepared 24 h previously to allow for
hydrogelation. Measurements were repeated three times and mean
value plotted. Control samples (sterile distilled water, 1 M NaOH, 0.5
M HCl, pH 7.4) did not display any signal.
Oscillatory Rheology. The viscoelastic properties of the samples
were determined using oscillatory rheology performed on a TA
AR1500 rheometer at 25 °C. A 20 mm steel parallel plate was
employed with a set gap of 500 μm. 400 μL samples were applied to
the plate using a pipetman. An oscillatory stress sweep was performed
to determine the linear viscoelastic region (LVR) of each sample. This
region, which is determined according to a superposition principle, is
the area where stress is directly proportional to strain. There is a linear
relationship between the two variables. Following this, an appropriate
stress within the LVR was selected and a frequency sweep was
performed at the range 1−10 Hz with the selected stress remaining
constant for each sample. Three distinct replicates for each sample
were obtained and the storage modulus (G′) and loss modulus (G″)
determined.
Hinton Broth (MHB) for 24 h at 37 °C before testing. Upon visual
conﬁrmation of growth, the inoculum was taken and adjusted to
optical density 0.3 at 550 nm in Quarter Strength Ringers Solution
(QSRS), equivalent to 1 × 108 colony forming units per milliter
(CFU/mL). This suspension was further diluted in MHB (1 in 50)
and dispensed aseptically in 100 μL aliquots to each well of the
microtiter plate, corresponding to 2 × 106 CFU/mL. Bioﬁlms were
formed on the surface of the well under shear stress provided by a
Gallenkamp gyrorotary incubator at 37 °C. After an inoculation period
of 24 h, cultures were decanted and the plates irrigated thrice with 200
μL of sterile autoclaved 0.9% w/v sodium chloride (NaCl) in each
well. Washed plates were tapped gently upside down on a sterile paper
towel to remove residual wash. The prepared bioﬁlm was treated for
evaluation of viability using alamarBlue in a method similar to that
used by Pettit.26 Then, 100 μL of 2.0 1.5, 1.0 and 0.5% w/v NapFF,
NapFFKK, NapFFFKK, NapFFOO, and NapFFK′K′ was pipetted
into separate wells of the microtiter plate and bioﬁlms assessed for
reduction in viability after 24 h exposure. Gels were removed from the
microtiter plates by three rinses of each well with NaCl. A range (3−
0.5% w/v) of HPMC gels (see Supporting Information, Figure S21)
were used as a standard non-antimicrobial hydrogel comparator to
ensure removal of gels did not remove a signiﬁcant quantity of bioﬁlm
bacteria. HPMC has previously been shown to possess no signiﬁcant
antimicrobial activity.27 HPMC displayed no signiﬁcant reduction of
viable bioﬁlm compared to positive controls (no gel), thus ensuring
the validity of the assay and methods employed. The viability of
established bioﬁlms is assessed by recharging wells with 100 μL of
MHB containing 20% v/v alamarBlue, incubating the plate at 37 °C
for 2 h to allow development,
followed by spectrophotometric
measurement of absorbance at a wavelength of 570 nm (Tecan Sunrise
plate reader). AlamarBlue contains resazurin, a redox indicator that is
reduced by the metabolic activity of viable microbial cells to develop
into pink ﬂuorescent resorfurin. Positive control (100% microbicidal
eﬀect) corresponds to blank wells (no bacterial bioﬁlm) treated with
70% ethanol (added, removed and allowed to dry) immediately prior
to the addition of 20% v/v alamarBlue. Eight
replicates were
performed at each concentration. Reduction in viability of bioﬁlm
following exposure to test formulation is expressed as a percentage
based on percent reduction between untreated (blank) wells and
treated wells using the following equation:
Cell Viability Assay. Murine ﬁbroblast subcutaneous connective
tissue cell line, NCTC Clone 929 (ATCC CCL 1), was cultured in
Minimum Essential Medium (MEM) containing phenol red with
Earle’s Salts and L-glutamine, supplemented with 10% Horse Serum,
100 IU/mL penicillin, and 100 μg/mL streptomycin supplied by
Invitrogen (Paisley, U.K.). Cells were grown at 37 °C and 5% CO2 and
subcultured at 80−90% conﬂuency. Subculturing consisted of removal
of spent medium, rinsing of the adherent cell surface with sterile
autoclaved PBS before treatment with a 0.05% trypsin/0.53 mM
EDTA·4Na solution (Invitrogen, Paisley, U.K.) for cell monolayer
detachment. Cell viability was assessed by means of a quantitative
alamarBlue assay, similar to that utilized by O’Brien and colleagues.28
Cells were cultured (until at least third passage) and inoculated into
96-well tissue culture treated microtiter plates at a concentration of 1
× 104 cells/well and incubated at 37 °C and 5% CO2 for 24 ± 1 h,
until approximately 90% conﬂuency. After this time, the medium was
removed and replaced with 100 μL of 2.0 1.5, 1.0, and 0.5% w/v
NapFF, NapFFKK, NapFFFKK, NapFFOO, NapFFK′K′ in separate
wells of the microtiter plate and cell monolayers exposed for 24 h.
Absorption was measured at 570 nm in a Tecan Sunrise plate reader
after a development time of 10 h. A positive control (100% reduction
in viability) was also included in the assay using 70% ethanol. The
negative control (untreated) consisted of tissue culture cells in 100 μL
phosphate buﬀer saline (PBS). Percentage cell viability was calculated
relative to untreated control wells as per the bioﬁlm assay above. This
value was subtracted from 100 to provide a percentage viability value
displayed in Figure 6. The 3−0.5% w/v HPMC controls (Supporting
Information Figure S22) displayed no signiﬁcant reduction of viable
ﬁbroblast cells compared to positive controls (no gels),
conﬁrming the validity of the study.
Hemolysis. All peptide gels/solutions were assayed spectrophoto-
metrically for their ability to induce hemoglobin release from fresh
equine erythrocytes according to the method previously described.21
Fresh deﬁbrinated equine erythrocytes were washed three times with
equal volumes of PBS. After centrifugation for 15 min at 900 g
erythrocytes were resuspended to 4% v/v in PBS. Equal volumes (100
μL) of the erythrocyte suspension were added to each well of a 96-well
microtiter plate. Erythrocytes were subsequently exposed to peptide
concentrations of 2.0−0.5% w/v,
incubated at 37 °C for 1 h, and
centrifuged at 1000g. Aliquots of the supernatant were transferred to a
fresh 96-well microtiter plate, and hemoglobin release measured
spectrophotometrically at 405 nm (Tecan Sunrise plate reader). As a
positive control (100% hemolysis), erythrocytes were treated with
0.1% Triton X-100, while PBS (0% hemolysis) acted as a negative
control. All peptide concentrations are reported as the mean of six
replicates. Percentage hemolysis was calculated as follows:
Statistical Analysis. Statistical analyses were performed using
GraphPad Prism 6. Standard deviations were obtained at each
concentration of peptide tested based on eight
quantitative bioﬁlm viability assays and mean values obtained. For
cell cytotoxicity assays standard deviations and mean values were also
obtained from eight
replicates at each concentration. Statistical
analyses were employed using a Kruskal−Wallis test, with a Dunn’s
multiple comparisons test used to identify individual diﬀerences
between the reduction in bioﬁlm viability for each peptide hydrogel
and the positive control (0% reduction). A Kruskal−Wallis test,
followed by a Dunn’s multiple comparisons test, was also utilized for
statistical analysis of tissue culture cytotoxicity data by comparison of
percentage viability for the peptides employed to the positive control
(100% viability), with signiﬁcant
reduction in viability reported.
Hemolysis data was compared by the same statistical method with
percentage hemolysis compared to the PBS, nonhemolytic, control
(0% hemolysis). Non-parametric Kruskal−Wallis tests were employed
rather than parametric Analysis of Variance (ANOVA) as data was
shown to be non-normally distributed using the Kolmogorov and
Smirnov method. In all cases, a probability of P < 0.05 denoted
signiﬁcance.

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