[System] polypeptide self-assembly literature information extraction

[User]
Peptidic self-assembled nanostructures are said to have a wide range of
applications in nanotechnology, yet the mechanistic details of hierarchical
self-assembly are still poorly understood. The Phe-Phe recognition motif of the
Alzheimer's Abeta peptide is the smallest peptide able to assemble into
higher-order structures. Here, we show that the Ile-Phe dipeptide analog is also
able to self-associate in aqueous solution as a transparent, thermoreversible
gel formed by a network of fibrillar nanostructures that exhibit strong
birefringence upon Congo red binding. Besides, a second dipeptide Val-Phe,
differing only in a methyl group from the former, is unable to self-assemble.
The detailed analysis of the differential polymeric behavior of these closely
related molecules provides insight into the forces triggering the first steps in
self-assembly processes such as amyloid formation.

Lyophilized NH2-Ile-Phe-COOH dipeptide and NH2-Val-Phe-COOH
dipeptide with .99% purity were obtained from Bachem (Bubendorf,
Switzerland). Dipeptide samples were diluted in 1,1,1,3,3,3 hexaﬂuoro-2-
propanol to obtain stocks of 100 mg/ml and 200 mg/ml, which were further
diluted to the assay concentration in H2O, except for NMR and Fourier trans-
form infrared (FTIR) assays, where the samples were diluted in deuterium oxide
High-resolution 1H NMR spectra were recorded for several Ile-Phe and Val-
Phe dipeptide samples at different concentrations and different temperatures.
The samples used consisted of 0.05%, 0.1%, 0.5%, 1%, and 2% (w/v) of
dipeptide dissolved in D2O from stock solutions. Spectra were also collected
in the range 295–330 K to observe the temperature dependence of each in-
dividual sample.
Dipeptide samples (1.5%, w/v) were placed on carbon-coated copper grids
and left for 5 min. The grids were then stained with 2% (w/v) uranyl acetate
for another 2 min before analysis using a Hitachi (Tokyo, Japan) H-7000
transmission electron microscope operating at an accelerating voltage of
75 kV. A sample of 2% (w/v) Ile-Phe gel smeared on a 1-cm slide was
allowed to dry at room temperature, followed by gold coating, before being
imaged on a Hitachi S-570 scanning electron microscope.
Congo red (CR) was diluted in a buffer containing 5 mM sodium phosphate
and 150 mM NaCl, pH 7.0, to obtain a stock of 100 mM CR. A 2% (w/v) Ile-
Phe gel was formed in the presence of 5 mM CR ﬁnal concentration. A gel
sample was placed on a microscope slide and sealed. The CR birefringence
was detected under cross-polarized light using an optic microscope (Leica
DMRB, Heidelberg, Germany).
Diluted, gelled, and air-dried dipeptide samples were used for FTIR spectro-
scopy analysis. Exchangeable hydrogen atoms were replaced by deuterium
by dissolving the dipeptide stocks in D2O. Infrared spectra were recorded
with an FTS-6000 FT-IR spectrophotometer (BioRad, Hemel Hempstead,
UK) equipped with a liquid nitrogen-cooled mercury/cadmium telluride
detector and purged with a continuous ﬂow of nitrogen gas or with a Bruker
Tensor 27 FT-IR spectrometer. For each spectrum, 200 interferograms were
collected and averaged. In every case, the buffer spectrum was subtracted
and the baseline corrected. Second derivatives of the spectra were used to
determine the frequencies at which the different spectral components were
located. To monitor the effect of pH on self-assembly, we measured the pH
established by the dipeptides themselves upon dilution in H2O, pH 5.8. This
pH was either increased to pH 12.0 by addition of 1 N NaOH or lowered to
pH 2.0 by addition of 1 N HCl to test the effect of N- and C-terminal group
ionization in Ile-Phe self-assembly.
The ﬂuorescence emission spectra of TNS with the dipeptide samples were
recorded at 293 K in a Perkin-Elmer 650-40 ﬂuorescence spectrophotom-
eter. TNS was diluted in H2O to obtain a 1 mM stock solution. The samples
were excited at 323 nm and the ﬂuorescence emission was measured
between 350 nm and 550 nm. Both excitation and emission slits were set at
10 nm. To follow the Ile-Phe dipeptide kinetic self-assembly by TNS
binding, a 2% Ile-Phe sample in a solution containing a ﬁnal 10 mM TNS
concentration was heated to 333 K and then cooled gradually to 278 K. The
FIGURE 1 Gelation of Ile-Phe dipeptide. (A) Structures of the dipeptides
compared in this work. NH1
3 -Ile-Phe-
COO(cid:2) (lower). (B) Photograph of 2% (w/v) dipeptide samples under blue
3 -Val-Phe-COO(cid:2)
light. NH1
remains in solution (right).
(D2O). 1,1,1,3,3,3 hexaﬂuoro-2-propanol and 2-(p-toluidinylnaphthalene)-
6-sulfonate (TNS) were purchased from Sigma (St. Louis, MO). D2O en-
riched .99.97% in isotope D2O (d ¼ 1.11) was purchased from SDS
(13124, Peypin, France).
The turbidity of the different dipeptide samples at each temperature was
measured monitoring the absorbance at 360 nm on a CARY-400 Varian
spectrophotometer (Les Ulis, France). To study the dependence of peptide
self-assembly on concentration, the turbidity was measured at 293 K. To
study the dependence of peptide assembly state on the temperature, each
sample was ﬁrst heated to 333 K or cooled to 283 K before measuring the
reassembly or disassembly, respectively. These samples were subsequently
cooled or heated in 5-K stages and equilibrated for 15 min before measuring
the turbidity at each assayed temperature.

[Assistant] [{"peptide_sequence": "IF", "self-assembly_phase": "none", "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)": "0.2", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "IF", "self-assembly_phase": "none", "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)": "1.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "IF", "self-assembly_phase": "fiber", "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)": "10.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "IF", "self-assembly_phase": "fiber", "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)": "20.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "VF", "self-assembly_phase": "none", "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)": "0.2", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "VF", "self-assembly_phase": "none", "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)": "1.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "VF", "self-assembly_phase": "none", "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)": "10.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}, {"peptide_sequence": "VF", "self-assembly_phase": "none", "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)": "20.0", "PH": "5.8", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}]


[System] polypeptide self-assembly literature information extraction

[User]
Author information:
(1)Beijing National Laboratory for Molecular Sciences (BNLMS), International
Joint Laboratory, Key Laboratory of Colloid and Interface Sciences, Institute of
Chemistry, Chinese Academy of Sciences, Beijing 100080, P. R. China.

Transition of Cationic Dipeptide Nanotubes into Vesicles and
Oligonucleotide Delivery**
Xuehai Yan, Qiang He, Kewei Wang, Li Duan, Yue Cui, and Junbai Li*
The self-assembly of biocompatible molecules,
such as peptides and proteins, into regular supra-
molecular structures has important implications. In
a sense these molecules mimic biological sys-
tems.[1, 2] Additionally, such biomimetic systems
are becoming important for the delivery of drugs,
genes, and proteins. Peptide-based nanostructures
have attracted considerable attention owing to
their biocompatibility, capability of molecular
recognition, and well-defined structures.[3–10] For
example, G’rbitz first found that simple dipeptides
can self-assemble into nanotubes, and that the
resulting structures have chiral hydrophilic chan-
nels with a Van der Waals diameter of up to
10 +.[10a] Reches and Gazit showed that very short
dipeptides with the b-amyloid diphenylalanine structural
motif associated with Alzheimer-s disease can form long,
stiff nanotubes by self-assembly, and that these nanotubes can
be used as scaffolds to produce discrete silver nanowires.[6]
Subsequently, Song et al. simplified the synthesis of the
dipeptide nanotubes (DPNTs) and observed that DPNTs
turned into vesicles when the DPNT dispersion was diluted by
adding water.[8] Zhang and co-workers also reported that
linear surfactant-like oligopeptides self-assembled simulta-
neously into nanotubes and vesicles.[7, 9] Such peptide nano-
tubes (PNTs) can be used for intracellular-delivery applica-
tions. Herein, we report that cationic dipeptides (H-Phe-Phe-
NH2·HCl; see Figure 1) self-assemble into nanotubes at
physiological pH values, and that these cationic dipeptide
nanotubes (CDPNTs) can also rearrange to form vesicles
upon dilution. Moreover, the CDPNTs can traverse cell
membranes and be absorbed by the cells upon spontaneous
conversion into vesicles. We utilized this property to deliver
oligonucleotides into the interior of cells as an indication of
the potential applications of the system in gene and drug
delivery.
We selected cationic dipeptides to fabricate nanotubes so
that negatively charged nucleic acids could be bound to the
tubes and then delivered into cells. The CDPNTs were self-
assembled at physiological pH values (see experimental
section in the Supporting Information). A scanning electron
microscopy (SEM) image showed that the nanotubes are
quite homogeneous with lengths of tens of micrometers
(Figure 2 a). The inset in Figure 2 a shows a typical tubular
structure at a larger magnitude. A transmission electron
Figure 2. a) SEM image of the CDPNTs (inset: a hollow nanotube at
greater magnification); b) TEM image of the CDPNTs; c) height AFM
image (area: 5 mm 0 5 mm) of the CDPNTs; the cross-sectional profile
of a nanotube shows its height of approximately 190 nm.
microscopy (TEM) image confirmed that tubes had formed
with a diameter of approximately 200 nm (Figure 2 b). To
obtain direct topographical information, the samples were
deposited on a mica surface and investigated by atomic force
microscopy (AFM; Figure 2 c). The height of the nanotube
shown is about 190 nm, as estimated from its cross-sectional
profile. All of these results confirm that the cationic dipeptide
can self-assemble into tubes at pH 7.2.
We find that the CDPNTs transform gradually into
vesicles when the CDPNT dispersion is diluted under neutral
conditions. As observed by optical microscopy, the CDPNTs
disappear with time, and the number of vesicles increases
simultaneously (Figure 3 a). To verify the structures of the
vesicles (the black dots in Figure 3 a), the solution was diluted
Figure 3. a) Observation through an optical microscope of the conver-
sion of the CDPNTs (as a dispersion in PBS at pH 7.2) into peptide
vesicles; b) CD spectra of the CDPNTs and vesicles at different
concentrations (A: 10, B: 8, C: 7, D: 5, E: 2, F: 1 mg mL(cid:1)1); c),d) TEM
images of negatively stained CDPNTs at a concentration of 7 and
5 mg mL(cid:1)1, respectively; e) CD spectra of zwitterionic DPNTs at differ-
ent concentrations (A: 10, B: 5, C: 2 mg mL(cid:1)1).
further with pure water to a final concentration of 1 mg mL(cid:1)1.
These diluted samples were then characterized by negative-
staining TEM and AFM (see the Supporting Information).
The TEM image showed that peptide vesicles are formed
after dilution of the nanotube dispersion. Analysis by AFM
confirmed the three-dimensional, spherical shape of the
nanostructures. The cross-sectional profile of the vesicles
showed that the peptide vesicles are approximately 100 nm in
height, which is consistent with the TEM analysis. All of the
above results confirm that the CDPNTs can convert sponta-
neously into vesicles when diluted at physiological pH values.
To further clarify this transition process, circular dichro-
ism (CD) spectroscopy was used to determine changes in the
secondary structure during the conversion of the self-assem-
bled CDPNTs into vesicles. CD spectra of the CDPNTs and
vesicles at different concentrations show that the CDPNT-to-
vesicle transition causes a change in the secondary structure
of the peptide aggregations (Figure 3 b). The observed CD
curve of the assembled CDPNT (10 mg mL(cid:1)1) has some
similarities with the CD signature of a-helical polypeptides,
and the observed extrema may be indicative of transitions
found in a-helical peptides. The maximum at 192 nm and the
minimum at 206 nm correspond to the a-helical p!p*
transitions. Another absorption peak at 232 nm may corre-
spond to n!p* transitions of helical arrangements of the
PNTs.[11, 12] The CD signal reflects the interactions between
the three-dimensional aromatic stacking arrangement and the
hydrogen-bonded cylinders of peptide main chains.[13] When
the CDPNT dispersion was diluted to a concentration of
7 mg mL(cid:1)1, the minimum peaks for the b sheets of the peptide
nanostructures were observed at 206 nm and 212 nm.[14, 15]
When the concentration was decreased further, these peaks
disappeared gradually. At 1 mg mL(cid:1)1, the secondary structure
of CDPNT vanished.
Although the CD spectra of the self-assembled structures
have not yet been assigned completely, they demonstrate that
dilution indeed results in a change in the peptide nano-
structures. Furthermore, TEM images show that at a concen-
tration of 8 mg mL(cid:1)1, the system maintains a tubular structure
(see the Supporting Information). However, when the
solution is diluted to 7 mg mL(cid:1)1, vesicles start to form and
coexist with the tubular structures (Figure 3 c). The ratio of
peptide vesicles to nanotubes becomes larger upon a further
decrease in the concentration to 5 mg mL(cid:1)1. The intermedi-
ate state of the conversion of CDPNTs into vesicles can be
observed directly (Figure 3 d). When the concentration of the
system was decreased to 1 mg mL(cid:1)1, the transition is com-
plete, and the system is composed entirely of vesicles. (see the
Supporting Information). The transition from tube to vesicle
occurs at a concentration of approximately 7 mg mL(cid:1)1, which
is consistent with the CD results.
For comparison, we chose a zwitterionic dipeptide,
diphenylalanine (l-Phe-l-Phe),
similar experiments
under the same conditions. We found that this dipeptide
could self-assemble into nanotubes at a concentration of
10 mg mL(cid:1)1 (see the Supporting Information). This self-
assembly is similar to results reported previously.[6] A clear
Cotton effect is observed at 232 nm in the CD spectra of these
zwitterionic DPNTs at a concentration of 10 mg mL(cid:1)1 (Fig-
ure 3 e). Upon a decrease in the concentration of these
zwitterionic DPNTs, the intensity of the CD signal at 232 nm
decreases dramatically. When the concentration of the system
is decreased to 2 mg mL(cid:1)1, no new peak appears. This result
indicates that within this concentration range the zwitterionic
dipeptide forms mainly tubular structures rather than vesicles,
unlike the cationic dipeptide, for which the transition from
nanotubes to vesicles can be observed at a concentration of
7 mg mL(cid:1)1.
 15213773, 2007, 14, Downloaded from https://onlinelibrary.wiley.com/doi/10.1002/anie.200603387 by Massachusetts Institute of Technolo, Wiley Online Library on [21/03/2024]. See the Terms and Conditions (https://onlinelibrary.wiley.com/terms-and-conditions) on Wiley Online Library for rules of use; OA articles are governed by the applicable Creative Commons Licenselabeled with 5-((5-aminopentyl)thioureidyl)fluorescein was
bound to the CDPNTs through electrostatic interactions.
Figure 4 a shows the green fluorescent CDPNTs detected by
confocal laser scanning microscopy (CLSM). The CDPNTs
are stable after binding with ssDNA, and the nanotube
Figure 4. CLSM images of a) the CDPNTs with bound fluorescently
labeled ssDNA; b) a selected single nanotube, which was used for the
intensity measurements (inset: intensity profile of a fluorescent
CDPNT); c) peptide vesicles from the reassembly of the peptides
derived from the CDPNTs with bound fluorescently labeled ssDNA;
d) a solution of fluorescently labeled ssDNA.
structure can be still observed after four weeks (see the
Supporting Information). Analysis of the fluorescence-inten-
sity distribution on an individual nanotube (Figure 4 b) shows
that the fluorescence remains mainly on the sidewalls of the
nanotubes (see inset in Figure 4 b). Furthermore, peptide
vesicles formed from CDPNTs retain the green fluorescence
(the green dots in Figure 4 c, in contrast with ssDNA alone in
Figure 4 d), which indicates that the ssDNA is localized on the
peptide vesicles after the conversion of the CDPNTs. We also
investigated the ability of CDPNTs to inhibit the migration in
gel electrophoresis of ssDNA. When the volume ratio of
CDPNTs to ssDNA was changed to 3:1 (see the Supporting
Information), the migration of ssDNA in gel electrophoresis
was inhibited. This observation indicates that ssDNA can be
immobilized securely on the cationic dipeptide nanostruc-
tures through electrostatic interactions. In contrast, zwitter-
ionic DPNTs and cationic dipeptide molecules in solution
without aggregation have no effect on the migration of
ssDNA in gel electrophoresis at different volume ratios (see
the Supporting Information), which indicates that ssDNA
does not bind to zwitterionic DPNTs and free cationic
dipeptides. These results confirm that ssDNA can be bound
to the cationic dipeptide nanostructures through charge
interactions, which are very important for the immobilization
on the nanostructures and intracellular release of DNA.[16, 17]
The dispersion of the CDPNT/ssDNA complexes was
dialyzed for 2 h with constant stirring in phosphate buffer
saline (PBS) to eliminate cytotoxic hexafluoro-2-propanol.
We then studied the interaction of the resulting complexes
with HeLa cells. CDPNTs labeled with Congo red (CR-
CDPNTs) were used for the control experiment of cellular
uptake. It was found that Congo red does not influence the
tubular structure of the cationic dipeptide (see the Supporting
Information). Both TEM and confocal-microscopy images
confirmed the stability of CDPNTs in the presence of Congo
red. The HeLa cells were incubated with CR-CDPNTs and
CDPNTs/ssDNA (c(CDPNTs) (cid:3) 0.5 mg mL(cid:1)1)
in a cell-
growth medium with 5 % CO2 at 37 8C for 24 h. The
medium containing CR-CDPNTs or the complexes was then
removed, the cells were rinsed gently twice, and a fresh
culture medium was added. Subsequently, the cells were
imaged directly in glass-bottomed Petri dishes by CLSM.
CLSM scans normal to the dish plane revealed that fluores-
cence originated mainly from the interior of
the cells
(Figure 5; see also the Supporting Information). This result
indicates that the CR-CDPNTs and CDPNT/ssDNA enter
the cells by converting into vesicles and accumulate in the
cytoplasma of the cells. No significant cell death was observed
when HeLa cells were incubated under the conditions
described above. As negative-control experiments, cells
were incubated with a solution that contained zwitterionic
Figure 5. Uptake of the CR-CDPNTs and CDPNTs/ssDNA by HeLa
cells: a) bright-field image of cells; b) fluorescence image of positive-
control cells after incubation for 24 h with the CDPNTs labeled with
Congo red; c),d) fluorescence images of cells after incubation for 24 h
with the complexes of CDPNTs and fluorescently labeled ssDNA; the
cell nuclei (dark circular regions) appear free of fluorescence; the
complexes of peptide vesicles/ssDNA are accumulated within the
cytoplasm of the cell.
dipeptide or only fluorescently labeled ssDNA. No fluores-
cence of the cells was detected by CLSM (see the Supporting
Information), which means that the CDPNTs can traverse cell
membranes by converting spontaneously into vesicles, and
that they can transport ssDNA into the cells.
The self-assembly of cationic dipeptides is driven by
hydrogen bonding and by p–p stacking interactions, as
described by Reches and Gazit for the self-assembly of
aromatic dipeptides into nanotubes.[6] Zhang and co-workers
pointed out previously that the self-assembly and disassembly
processes of linear surfactant-like peptide nanostructures are
dynamic over time.[7a] On the basis of molecular simulations
of the d-Phe-d-Phe dipeptide, Song et al. suggested that the
concentration of the peptide is key to the self-assembly
process of PNTs.[8] The conversion of CDPNTs into vesicles
also appears to be a dynamic process. When the concentration
their disassembly occurs
probably as a result of electrostatic repulsion rather than
hydrogen bonding or p–p stacking interactions between
peptide molecules. As observed under the optical microscope,
vesicles of relatively incompact stacking are formed rapidly
by peptide reassembly. Modeling and simulation studies of
this process are still required for further understanding. As for
the mechanism of cellular uptake of the CDPNTs, we propose
that their intracellular internalization occurs by endocytosis
of the reassembled vesicles (Figure 1), which is a well-known
mechanism for many species.[18–22]
In summary, self-assembled positively charged DPNTs
underwent spontaneous conversion into vesicles upon dis-
persion dilution. CDPNTs with fluorescently labeled ssDNA
bound electrostatically could enter cells readily, most likely
after conversion into vesicles with attached ssDNA. The
fluorescently labeled ssDNA accumulates in the cytoplasm of
the cells after internalization. Thus, because of their biocom-
patibility, bioabsorption, and recyclability, it may be possible
to exploit the peptide nanostructures as a new class of
molecular transporter for the delivery of a wide range of
foreign substances, such as drugs, genes, and proteins.

[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": "8/92 HFP/H2O", "peptide_concentration(mg/ml)": "10.0", "PH": "7.2", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "nan"}


[System] polypeptide self-assembly literature information extraction

[User]
the sol-
vent  was  removed  on  a  rotary  evaporator  under  reduced
pressure.  The  residue  was  redissolved  in  ethyl  acetate  and
washed sequentially with solutions of 10% citric acid (3x20
mL), water (3x20 mL), saturated solution of NaHCO3 (3x20
mL) and water (3x20 mL). The organic phase was collected
and dried over anhydrous sodium sulphate. The solvent was
removed under reduced pressure to yield Boc-Phe-Gly-OMe
(2) in ̴ 75% yield (2.51 g). 1H-NMR (CDCl3) δ: 7.4-7.2 (m,
5H of Phe), 4.35 (br, 1H, α-CH of Phe), 4.1 (m, 2H, CH2 of
Gly), 3.7 (s, 3H of OCH3), 3.1 (d, 2H, β-CH2 of Phe) and 1.4
(s, 9H, 3xCH3 of Boc).
The  Boc-  group  removed  from  (2)  (1.69  g,  5  mmol)  by
treatment  with  50%  TFA  in  DCM  (10  mL).  The  reaction
mixture  was  stirred  for  2h  and  completion  of  the  reaction
was  monitored  on  TLC.  The  solvent  was  removed  com-
pletely  and  excess  of  TFA  was  removed  by  trituration  with
dry ether  to obtain  TFA.H-Phe-Gly-OMe (3) in quantitative
To  a  solution  of  TFA  salt  (3),  DIPEA  (0.98  mL,  5.5
mmol) and Boc-L-Proline (1.07 g, 5 mmol) in dry DMF (20
mL),  DIPCI  (0.85  mL,  5.5  mmol)  was  added  at  5oC  with
continuous stirring. The reaction was monitored on TLC and
after completion of the reaction, solvent was removed under
reduced pressure. The work-up was carried out as discussed
above  and  the  crude  compound  purified  on  silica-column
chromatography  using  methanol-ethyl  acetate  (1:9  ratio)  as
an  eluent  to  obtain  pure  Boc-Pro-Phe-Gly-OMe  (4)  in  80%
yield  (1.73  g).  The  compound  (4)  was  further  purified  on
HPLC  using  a  reverse  phase  C18  column  with  a  gradient  of
water-acetonitrile  (0-100%  acetonitrile  containing  0.01%
TFA);  1H-NMR  (CDCl3)  δ:  7.4-7.2  (m,  5H  of  Phe),  4.2-3.9
(m, 4H, α-CH of Pro, α- CH of Phe and CH2 of Gly), 3.7 (s,
3H of OCH3), 3.1 (br, 4H, δ-CH2 of Pro and β-CH2 of Phe),
2.3-1.7 (m, 4H, β-CH2 and γ-CH2 of Pro), 1.4 (s, 9H, 3xCH3
of  Boc);  MS:  m/z,  434.38  (calculated)  and  434.26  (ob-
served).
For the formation of self-assembled nanostructures, a so-
lution  of  peptide  (4)  in  100μL  methanol  or  trifluoroethanol
was  diluted  with  900  µL  of  water  and  vortexed  for  2  min.
The  dispersed  sample  was  left  at  room  temperature  for  2h
and DLS studies of the resulting solution was carried out.
For  transmission  electron  microscopic  (TEM)  imaging,
grids were prepared by depositing 10 μL solution of samples
on carbon-coated copper grids with 1% uranyl acetate nega-
tive  staining  and  images  were  observed  at  an  accelerating
voltage of 200 kV on HR-TEM (Tecnai G2 20 twin, Tecnai
200 kV twin microscope).
Aspirin-loaded nanostructures  were prepared by dissolv-
ing peptide (4) (5mg) and aspirin (1mg) in 200 μL of metha-
nol  with  continuous  vortexing  for  1  min.  Then  800  μL  of
water was added and again vortexed for 1min. The solutions
were left overnight for stabilization of self-assembled nanos-
tructures.  The  resulting  mixture  was  lyophilized  to  obtain
desired drug loaded nanostructure. Similarly, eosin and cur-
cumin were also loaded into the nanostructures.
The  stabilization  of  drug-encapsulated  nanostructures
was  carried  out  with  vitamin  E–TPGS  by  using  standard
method [23]. Briefly, a solution of vitamin E–TPGS in min-
eral  oil  (100  mL,  0.4%  w/v)  was  prepared  and  heated  at
35oC, subsequently drop wise addition of aspirin-loaded pep-
tide  nanostructures  (5  mg  in  1  mL  water)  to  the  vitamin  E-
TPGS  solution  was  done  with  continuous  stirring  at  1400
rpm  to  form  water  in  oil  emulsion  (w/o  emulsion).  After
emulsification, solution was stirred for 2 h at 4oC to stabilize
the  Vitamin  E-TPGS  coated  nanoparticle  through  self-
assembly. The resulting suspension was diluted with hexane
to make the final concentration of 20% (v/v). The suspension
was  centrifuged  and  the  organic  layer  was  removed.  The
aqueous layer was washed three times with hexane (1mL) to
remove  the  remaining  mineral  oil  and  followed  by  vacuum
drying. The same procedure was repeated for eosin and cur-
cumin loaded nanostructures.
To study the interaction of drugs with peptide nanostruc-
tures,  fluorescence  assay  was  performed  following  reported
procedure [21]. Briefly, 20 µL of 4, (10 mg/mL in methanol)
was added to aqueous solution of eosin (10 µM, 190 µL) and
vortexed  for  1  min.  Emission  fluorescence  spectra  of  the
resulting  solution  (excitation  wavelength  518  nm)  was  re-
corded  in  the  range  from  540  to  600  nm  using  spectro-
fluorimeter (Fluoromax-4, Horiba) at different time intervals.
The release studies of entrapped drugs were done by tak-
ing the above vitamin E-TPGS stabilized samples (̴ 5mg) in
a dialysis membrane (MWCO 500-1000 Da). The membrane
was  placed  in  a  beaker  containing  50  mL  of  distilled  water
(ELIX) stirred at a speed of 100 rpm. Aliquots of 1mL were
taken out  at fixed  time  intervals from the beaker and absor-
bance  of  the  samples  was  read  at  wavelength  295  nm  for
aspirin and 517 nm in case of eosin using UV-Visible Spec-
trophotometer.
In vitro cytotoxicity assay of void peptide nanostructures,
curcumin  loaded  peptide  nanostructures,  TPGS  stabilized
void  peptide  and  TPGS  stabilized  curcumin  loaded  peptide
nanostructures  was  performed  in  cancerous  MCF-7  (human
breast  cancer) and normal HaCaT (human keratinocyte) cell
lines  by  MTT  colorimetric  assay.  Briefly  5000  cells/well
were plated in a 96-well plate. These cells were treated with
void  peptide  nanostructures  and  curcumin  loaded  peptide
nanostructures in range of 3.2 - 64 μg/mL and a range of 10 -
100  μg/mL  was  used  for  TPGS  stabilized  void  peptide
nanostructures and TPGS stabilized curcumin loaded peptide
nanostructures.  These  cells  were  exposed  to  above  nanos-
tructures  for  24  h  and  subsequently  100  μL  of  MTT
(1mg/mL)  in  DMEM  was  added  to  wells.  The  cells  were
incubated  for  2h  at  37°C.  The  formazan  crystals  so  formed
were  solubilized  in  DMSO.  The  intensity  of  the  color  was
measured  spectrophotometrically  at  540  nm  on  an  ELISA
plate  reader.  The  untreated  cells  were  taken  as  control.  The
cell viability was calculated using the formula: Cell Viability
(%) = (Abs)treated / (Abs)control x 100
Molecular  self-assembly  offers  unique  directions  for  the
fabrication of novel supramolecular structures and advanced
materials.  Peptide-based  nanostructures  indicate  an  impor-
tant route toward the production of ordered nanostructures as
several  studies  have  demonstrated  their  ability  to  form  well
organized  assemblies.  Small  and  modified  peptides  have
been  reported  to  self-assemble  through  various  interactions
into shape-specific nanostructures such as micelles, vesicles,
nanospheres,  nanotubes,  and  other  morphologies.  Most  of
the  small  peptides  have  aromatic  residues,  which  facilitate
self-assembly  via  hydrophobic  and π -π  interactions  making
less soluble  in  aqueous systems  and thus hampering several
potential  applications.  To  address  these  concerns,  sincere
efforts have been  made by introducing surface modification
or stabilization by surfactants [23-25]. In the present investi-
gation,  we  synthesized  a  protected  tripeptide  (4),  Boc-Pro-
Phe-Gly-OMe,  by  the  standard  solution  phase  peptide  syn-
thesis  method  using  L-amino  acids  (Scheme  1)  and  charac-
terized  by  mass  and  1H-NMR.  The  peptide  self-assembly
was carried out in aqueous solutions.
The peptide self-assembly was triggered by adding water
to a methanolic solution of peptide (4) and vortexing resulted
in the formation of nanostructures, which were analyzed by
DLS (Table 1  and Fig. 1c).  The average size of the peptide
nanostructure  was  found  to  be  119.6  nm.  Similar  results
were  obtained  when
trifluoroethanol (TFE) followed by addition of water to form
nanostructure  of  average  size  117.5  nm.  The  morphology
and size of the nanostructures were also confirmed by TEM.
The self-assembled peptide nanostructures displayed spheri-
cal  sized  particles  of  77.37  nm  and  72.08  nm  in  methanol-
water and TFE-water systems respectively (Fig. 1 a, b). The
smaller  size  of  self-assembled  nanostructures  in  TEM  com-
pared  to  DLS  might  be  due  to  measurement  of  hydrody-
namic diameter in DLS, which measured the size of swollen
nanostructures [26,27].
In  order  to  evaluate  the  encapsulation  of  hydrophobic
molecules  in  the  self-assembled  nanostructures,  eosin,  aspi-
rin and curcumin were subjected to encapsulation. DLS stud-
ies  of  the  nanostructures  after  encapsulation  were  also  car-
ried out (Table 1). In comparison to the unloaded nanostruc-
tures the average size of the nanostructures was found to be
higher  after  loading  the  compounds.  The  data  showed  that
after  loading,  the  size  of  the  drug-loaded  nanostructures  in-
creased from 119.6 nm to 135.5 nm, 217.1 nm and 143.9 in
case of eosin, aspirin and curcumin, respectively. The encap-
sulation occurred due to the hydrophobic-hydrophobic inter-
actions which confirmed the presence of hydrophobic matrix
in self-assembled nanostructures. The loading of eosin, aspi-
rin  and  curcumin  was  performed  at  different  w/w  ratios  of
peptide:drug and it was found to be 13.8% (w/w 4:1), 33.3%
(w/w  2:1)  and  3.84%  (w/w  25:1)  respectively.  The  %  drug
entrapment efficiency of nanostructures was found to be 66.6
(eosin),  100  (aspirin)  and  0.4  (curcumin)  at  above  w/w  ra-
tios.  The  morphology  and  size  of  the  nanostructures  were
observed  by  TEM  which  showed  spherical  shaped  self-
assemblies  with  average  size  of  the  nanostructures  ̴  72.08
nm, 94.97 nm and 48.29 nm in eosin, aspirin and curcumin-
loaded  nanostructures  respectively  (Fig.  2a,  3a,  4a).  As  ex-
pected,  the  average  size  of  nanostructures  in  the  TEM  was
found to be smaller as compared to DLS.
The  loaded nanostructures were further stabilized with  a
biocompatible  and  biodegradable  surfactant,  Vitamin  E-
TPGS,  which  improved  the water dispersibility of the drug-
loaded  nanostructures.  The  DLS  measurements  showed  that
after vitamin E-TPGS stabilization, the size of the nanostruc-
tures  increased  from  135.5  nm  to  309.2  nm,  217.1  nm  to
313.5 nm and 143.9 to 249.9 nm in case of eosin, aspirin and
curcumin,  respectively  (Table  1).  These  structures  were  of
spherical  shaped  with  average  size  was  around  80.68  nm,
116.6  nm  and  125.83  nm  in  eosin,  asprin  and  curcumin
loaded  nanostructures,  respectively  (Fig.  2b,  3b,  4b).  The
increase in average  size  confirmed  the stabilization of  these
nanostructures with Vitamin E-TPGS, which may be due  to
the hydrophobic- hydrophobic interaction.
Peptide  based  nanostructures  are  an  important  class  of
biomaterials having potential use in tissue engineering, food
industry, micro fluidics and drug delivery. Loading of mole-
cules in nanostructures  is of the main  issues  in the develop-
ment  of  drug  delivery  systems.  Different  types  of  weak  in-
teractions  such  as  electrostatic,  hydrophobic  and  H-bonding
have been found to be responsible for the formation of self-
assembled nanostructures. Here,  the  loading of hydrophobic
molecules  was  confirmed  by  eosin-fluorescence  quenching
study.  Interaction  of  eosin  with  peptide  was  monitored  by
emission  fluorescence  quenching  (Fig.  5).  It  was  observed
that  eosin  fluorescence  at  ~540  nm  got  quenched  in  time
dependent manner. For this purpose, a solution of peptide (4)
in methanol was mixed with an aqueous solution of eosin (10
µM in water) and the emission of eosin (λex - 515 nm, λem -
540 nm) was measured at different intervals of time (Fig. 5).
intensity  gradually
that  fluorescence
It  was  observed
dropped at 540 nm with increasing time  intervals. These re-
sults confirmed that loading of eosin occurred via hydropho-
bic  interactions  between  hydrophobic  eosin  molecules  and
the hydrophobic moiety in peptide nanostructures [21,28,29].
Fig. (1). TEM images of  self-assembled nanostructures of (a) peptide (4)  in 10% methanol,  scale bar : 0.

[Assistant] [{"peptide_sequence": "PFG", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Boc", "C-terminal_modification": "Methoxy", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "10/90 Methanol/H2O", "peptide_concentration(mg/ml)": "1.0", "PH": "7.5", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "120.0"}, {"peptide_sequence": "PFG", "self-assembly_phase": "nanosphere", "N-terminal_modification": "Boc", "C-terminal_modification": "Methoxy", "Non-terminal_modification": "nan", "peptide_conjugate_mixture": "Peptide", "conjugate_partner": "nan", "solution": "10/90 TFE/H2O", "peptide_concentration(mg/ml)": "1.0", "PH": "6.0", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "120.0"}]


[System] polypeptide self-assembly literature information extraction

[User]
 that are based on the proteinogenic
amino acid phenylalanine in combination with serine, cysteine, glutamate,
histidine and lysine are described as simple and remarkable low molecular weight
hydrogelators. Blends of selected DKPs show remarkable pH-dependent properties
and can be applied as easy to tune materials in drug delivery.

Phenylalanine-containing cyclic dipeptides – the lowest
molecular weight hydrogelators based on unmodified
proteinogenic amino acids†
Physical molecular hydrogels based on low molecular weight
gelators (LMWGs) have become an important class of materials
due to their versatile applications as innovative soft materials
in injection based drug delivery and tissue engineering.1 The
gel-resulting self-assembly process of LMWGs is initiated by
non-covalent forces such as hydrogen bonding, p-stacking or
van der Waals interactions. Above all, amino acids and small
peptides are popular LMWGs since they are easy to vary and
often show promising biocompatibility.2,3 However, intermolec-
ular hydrogen bonding networks between amide bonds alone, in
particular in water, are in general not suﬃcient to make a small
peptide or amino acid an eﬃcient hydrogelator. Thus, function-
alization of the peptide N- or C-terminus with protecting groups
capable of excessive p-stacking interactions is necessary. Low
molecular weight peptide based hydrogelators which only contain
unmodified proteinogenic amino acids are rare.4 In our search for
the smallest amino acid based units that can form higher ordered
structures in an aqueous environment,5 we herein want to present
cyclic dipeptides (diketopiperazines – DKPs) as representative
examples for remarkable small LMWGs that self-assemble into
nanofibers and form stable, self-healing hydrogels.6,7
University of Tuebingen, Institute of Organic Chemistry, Auf der Morgenstelle 18,
D-72076 Tuebingen, Germany. E-mail: boris.nachtsheim@uni-tuebingen.de;
Fax: +49 7071 295897; Tel: +49 7071 29 72035
† Electronic supplementary information (ESI) available: Synthetic procedures,
spectroscopic data as well as 1H and 13C spectra of DKPs 1–6, rheological
measurements and concentration dependent sol–gel transition temperatures of
1–6 and blends of 4 and 6. See DOI: 10.1039/c3cc44110e
open-chained di- or oligopeptides. Therefore, we synthesized a small
set of DKPs containing the lipophilic amino acid phenylalanine and
a variety of hydrophilic amino acids, in particular glycine (1), serine
(2), cysteine (3), glutamate (4), histidine (5) and lysine (6) (Fig. 1).
In an initial experiment we were intended to verify the principle
hydrogelation ability of cyclic dipeptides 1–6. Thus, we heated an
aqueous suspension containing 4 wt% of each DKP until the solid
completely dissolved. Subsequently, the saturated peptide-containing
solution was cooled to rt or even lower. To our great delight, each
DKP 1–6 formed stable hydrogels under these conditions (Fig. 2).
DKPs 1–3 and 5 form opaque hydrogels. Glutamate-containing
DKP 4 yields a transparent hydrogel (at pH 6.0 in phosphate buffer).
The lysine-containing DKP 6 also yields a transparent hydrogel. To
further elucidate their thermal stability, we examined the sol–gel
transition temperature of these hydrogelators systematically as a
function of DKP-concentration (ESI,† Fig. S1A). Cyclo(L-Phe-L-Cys) 3
was the most effective hydrogelator since it formed a stable hydrogel
down to a remarkably low concentration of only 0.25 wt%. It shows a
notable high sol–gel transition temperature of up to 100 1C at 1 wt%.
In contrast, the corresponding serine-containing DKP 2 turned out to
be the least efficient hydrogelator since high peptide loadings (at least
4 wt%) are needed for a hydrogelation with a low sol–gel transition
Fig. 2 Macroscopic appearance of DKP-based hydrogels; (A) cyclo(L-Phe-Gly) 1.
(B) Cyclo(L-Phe-L-Ser) 2. (C) Cyclo(L-Phe-L-Cys) 3. (D) Cyclo(L-Phe-L-Glu) in phosphate
buﬀer at pH 6.0. (E) Cyclo(L-Phe-L-His). (F) Cyclo(L-Phe-L-Lys).
temperature of 12 1C. In general, all DKPs, except 2, form stable
hydrogels at rt with a low concentration of 2 wt%. Cyclo(L-Phe-Gly) 1
which bears no hydrophilic functional group forms stable hydrogels
down to a concentration of 1 wt%. To the best of our knowledge, 1 is
the lowest molecular weight peptide-based hydrogelator containing
only proteinogenic amino acids that has been described so far.8,9
Next, we were interested in the shape and morphology of the
hydrogels on a nanoscale. Therefore, we prepared xerogels of the
corresponding hydrogels through freeze drying and performed
scanning electron microscopy (SEM) (Fig. 3). Xerogel morphology
of 1 showed porous, but highly ordered planes (Fig. 3A and B). Each
plane is woven by knotted nanofibers. DKP 2 also consists of porous
planes (Fig. 3C and D) which contain entangled nanofibers and
partly microcrystalline domains. DKP 3 is constructed by undefined
nanosheets (Fig. 3E and F) while the glutamate-containing DKP 4
(Fig. 3E and F) forms a defined porous network at pH 6.0 (in
phosphate buﬀer).
DKPs 5 and 6 form dense lamellar sheets with bundled nano-
fiber connections between the sheets (Fig. 3I–L). The viscoelastic
properties of the hydrogels were investigated by rheological strain
sweep experiments (ESI,† Fig. S2–S7A). The values of the storage
modulus G0 are in the remarkable range of 103–106 Pa. A strong age-
dependent storage modulus was observed for the lysine containing
hydrogel 6 which increases from 697 Pa for a freshly prepared gel
sample to 146 kPa for a gel sample aged for 20 h prior to the
rheological measurement. On the other hand, the freshly prepared
sample showed the highest mechanical stability with a high critical
strain level of 10% deformation. Further time sweep experiments
showed a remarkable increase of G0 upon aging for all DKPs (ESI,†
Fig. S2–S7D). In agreement with comparable hydrogels, the loss
moduli (G00) were found to be at least one order of magnitude lower
than G0 indicating that the hydrogels described herein are true
physical hydrogels and not viscoelastic fluids.10 Frequency sweep
experiments of all hydrogels showed a very weak dependency on the
angular frequency suggesting that the gel matrices have excellent
tolerance towards external forces (ESI,† Fig. S2–S7B). Furthermore,
most of the hydrogels (except of 2) showed a significant thixotropic
behaviour and thus have self-healing properties (ESI,† Fig. S2–S7C).
We further asked ourselves whether we can tune the mechanical
and morphological properties of the hydrogel by mixing two or more
DKPs. Due to recent reports regarding tuneable two-component
hydrogels based on the interaction between diamines and carboxylic
acids,11,12 we studied the gelation behaviour of a 1 : 1 mixture
between the glutamate containing DKP 4 and the lysine containing
We initially investigated the sol–gel transition temperature of the
blended hydrogel and found remarkable pH-dependency. While pure
4 only formed stable hydrogels at pH 6.0 and pure 6 only formed
stable hydrogels in basic solution (pH 8–11), the blend formed a clear
hydrogel within a remarkably widespread pH-range between pH 2 and
pH 11. The sol–gel temperature reaches a maximum at pH 2 (79 1C)
and a minimum at neutral pH-values (27 1C, Fig. 4B). The viscoelastic
properties showed significant pH-dependency as well, which
was inverse to the sol–gel temperatures with a maximum G0 value
Fig. 3 SEM images of xerogels. Unless otherwise stated, gelation was performed
in an unbuﬀered aqueous solution (A and B) cyclo(L-Phe-Gly) 1 at pH = 7. (C and D)
Cyclo(L-Phe-L-Ser) 2 at pH = 7. (E and F) Cyclo(L-Phe-L-Cys) 3 at pH = 7. (G and H)
Cyclo(L-Phe-L-Glu) 4 at pH = 6.0 (phosphate buﬀer). (I and J) Cyclo(L-Phe-L-His) 5 at
pH = 8. (K and L) Cyclo(L-Phe-L-Lys) 6 at pH = 10–10.5.
Fig. 4 (A) SEM image of a freeze-dried xerogel (2 wt% 4 and 2 wt% 6 at pH 7).
(B) pH-dependent sol–gel temperature of a 1 : 1 blend of 4 and 6 (2 wt% each)
compared to pure DKPs (2 wt%).
3 For selected recent examples for peptide-based LMW hydrogelators see:
(a) V. J. Nebot, J. Armengol, J. Smets, S. F. Prieto, B. Escuder and
J. F. Miravet, Chem.–Eur. J., 2012, 18, 4063–4072; (b) S. Marchesan,
C. D. Easton, F. Kushkaki, L. Waddington and P. G. Hartley, Chem.
Commun., 2012, 48, 2195–2197; (c) R. Orbach, I. Mironi-Harpaz, L. Adler-
Abramovich, E. Mossou, E. P. Mitchell, V. T. Forsyth, E. Gazit and
D. Seliktar, Langmuir, 2012, 28, 2015–2022;
(d) K. A. Houton,
K. L. Morris, L. Chen, M. Schmidtmann, J. T. A. Jones, L. C. Serpell,
G. O. Lloyd and D. J. Adams, Langmuir, 2012, 28, 9797–9806;
(e) S. Grigoriou, E. K. Johnson, L. Chen, D. J. Adams, T. D. James
and P. J. Cameron, Soft Matter, 2012, 8, 6788–6791; ( f ) M. Hughes, P.
W. J. M. Frederix, J. Raeburn, L. S. Birchall, J. Sadownik, F. C. Coomer,
I. H. Lin, E. J. Cussen, N. T. Hunt, T. Tuttle, S. J. Webb, D. J. Adams
and R. V. Ulijn, Soft Matter, 2012, 8, 5595–5602; (g) G. Pont, L. Chen,
D. G. Spiller and D. J. Adams, Soft Matter, 2012, 8, 7797–7802;
(h) X. Zhang, X. Chu, L. Wang, H. Wang, G. Liang, J. Zhang, J. Long
and Z. Yang, Angew. Chem., Int. Ed., 2012, 51, 4388–4392; (i) C. Ou,
J. Zhang, X. Zhang, Z. Yang and M. Chen, Chem. Commun., 2013, 49,
1853–1855; ( j) J. Nanda, A. Biswas and A. Banerjee, Soft Matter, 2013,
9, 4198–4208;
(k) R. Wieduwild, M. Tsurkan, K. Chwalek,
P. Murawala, M. Nowak, U. Freudenberg, C. Neinhuis, C. Werner
and Y. Zhang, J. Am. Chem. Soc., 2013, 135, 2919–2922.
4 For examples of low molecular weight peptide self-assemblies/hydro-
gels see: (a) M. Reches and E. Gazit, Science, 2003, 300, 625–627;
(b) N. S. de Groot, T. Parella, F. X. Aviles, J. Vendrell and S. Ventura,
Biophys. J., 2007, 92, 1732–1741; (c) M. J. Krysmann, V. Castelletto,
A. Kelarakis, I. W. Hamley, R. A. Hule and D. J. Pochan, Biochemistry,
2008, 47, 4597–4605; (d) J. Naskar, G. Palui and A. Banerjee, J. Phys.
Chem. B, 2009, 113, 11787–11792; (e) A. Saiani, A. Mohammed,
H. Frielinghaus, R. Collins, N. Hodson, C. M. Kielty, M. J. Sherratt
and A. F. Miller, Soft Matter, 2009, 5, 193–202; ( f ) S. Marchesan,
C. D. Easton, F. Kushkaki, L. Waddington and P. G. Hartley, Chem.
Commun., 2012, 48, 2195–2197.
5 S. Zhang, Acc. Chem. Res., 2012, 45, 2142–2150.
6 A. D. Borthwick, Chem. Rev., 2012, 112, 3641–3716.
7 For DKP-derived organo- and hydrogelators see: (a) Z. Xie, A. Zhang,
L. Ye, X. Wang and Z.-G. Feng, J. Mater. Chem., 2009, 19, 6100–6102;
(b) Z. Xie, A. Zhang, L. Ye and Z.-G. Feng, Soft Matter, 2009, 5,
1474–1482; (c) S. Manchineella and T. Govindaraju, RSC Adv., 2012,
2, 5539–5542; (d) K. Hanabusa, Y. Matsumoto, T. Miki, T. Koyama
and H. Shirai, J. Chem. Soc., Chem. Commun., 1994, 1401–1402.
8 An exceptional low molecular weight squalen-based hydrogelator
was recently described by Ohsedo and co-workers: Y. Ohsedo,
M. Miyamoto, M. Oono, K. Shikii and A. Tanaka, RSC Adv., 2013,
3, 3844–3847.
Fig. 5 Drug release experiment with BSA (dotted lines) and tetracycline
(continuous lines) in diﬀerent blend hydrogels of DKPs 4 and 6. (DKP 4 : DKP 6).
at pH 7 (ESI,† Fig. S8A). However, the fastest regeneration is
observed at pH 10 (ESI,† Fig. S8B). A SEM image of a freeze dried
sample at pH 7 showed a so far unseen morphological shape
with a highly ordered lamellar structure (Fig. 4A). Xerogel
morphologies for other pH values are shown in the ESI† (Fig. S9).
Finally, we were interested in an application of our newly found
hydrogel blends in a drug release experiment. We particular asked
ourselves whether we can modulate the drug releasing properties of
the hydrogel by simply varying the DKP ratio. Thus 1 : 1, 1 : 2 and 2 : 1
mixtures of 4 and 6 were investigated in the release of (1) tetracycline
(TC) as a lipophilic small molecule and (2) bovine serum albumin
(BSA) as a highly charged polypeptide as model substrates (Fig. 5).13
For both substrates a significant diﬀerence in drug release was
observed. While nearly all BSA was released after 24 h for a 2 : 1
mixture of 4 and 6, only 55% was released with the 1 : 2 mixture. The
hydrogel containing a 1 : 1 mixture of 4 and 6 lies in-between (75%).
For TC the fastest release was observed for the 1 : 1 and the 2 : 1
mixtures (82% and 80%) while the 1 : 2 mixture showed a signifi-
cantly delayed release (62%).
In summary, we herein present the remarkable hydrogelation
properties of phenylalanine-containing cyclic dipeptides (diketo-
piperazines – DKPs) which only contain proteinogenic amino acids
as building blocks. To the best of our knowledge these structures are
the smallest peptide-based hydrogelators that have been described
so far. Despite their simplicity and the low molecular weight of their
monomeric building blocks, the so formed hydrogels are remarkably
stable and self-healing. We could further demonstrate that a blend
of two well-chosen DKPs shows novel pH-dependent mechanical
properties and can be used as soft materials for delayed drug release.
We gratefully

[Assistant] [{"peptide_sequence": "FG", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FS", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FC", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "7.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FE", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "6.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FH", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "8.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "8.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "8.6", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "9.2", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "9.8", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "10.4", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}, {"peptide_sequence": "FK", "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 PB", "peptide_concentration(mg/ml)": "20.0", "PH": "11.0", "temperature(celsius)": "25.0", "heating_cooling": "Yes", "incubation_period(min)": "nan"}]


[System] polypeptide self-assembly literature information extraction

[User]
Self-assembly of peptide into nanostructures is believed to be stabilized
primarily by aromatic interactions. Using a minimalistic approach, we probed the
importance of aromatic interactions in the self-assembly of simple model
dipeptides. Our results suggest that aromaticity may not be absolutely essential
for self-assembly, even though it tends to provide directionality to the
assembly. We found that peptides containing cyclic/linear side chain hydrophobic
residues were also capable of forming stable self-assemblies that are stabilized
by hydrophobic interactions. Our observations will find relevance in the design
of small peptide based nanoparticles.

The relevant N-terminal protected amino acid (Boc-Xaa-COOH)
(5 mM) was dissolved in dry tetrahydrofuran (Sigma-Aldrich) and
the resulting solution stirred in an ice-salt bath at (cid:2)15 (cid:4)C. N-
Methylmorpholine (Sigma) (0.65 ml, 5 mM) was added to the solu-
tion followed by isobutyl chloroformate (Sigma) (0.7 ml, 5 mM).
After 30 min, a pre-cooled aqueous solution of the incoming amino
acid (H-Yaa-COOH) (5.5 mM) and sodium hydroxide (0.22 g,
5.5 mM) was added and mixture stirred overnight at room temper-
ature. The reaction mixture was concentrated in vacuo, acidiﬁed with
citric acid to pH 3.0 and extracted with ethyl acetate (Spectrochem)
(3 (cid:5) 20 ml). The ethyl acetate layer was washed with water (2 (cid:5) 15
ml), with saturated sodium chloride (1 (cid:5) 20 ml), dried over anhy-
drous sodium sulfate and evaporated to yield Boc-Xaa-Yaa-COOH
dipeptide. Deprotection at the a-amino group and side chain
protection were achieved by treatment with 98% formic acid (30 ml)
for 3 h or 50% TFA : DCM for 30 min at room temperature. The
reaction mixture was evaporated to dryness and the residue was
precipitated with anhydrous diethyl ether in cold (Cha–Cha dipeptide
was precipitated with ice-cold double distilled water). The resulting
precipitate was ﬁltered, washed several times with dry ether and
subsequently lyophilized from 10% acetic acid–water (20 ml) to yield
the ﬁnal compound. Synthetic yields were typically between 90 and
The peptides were puriﬁed on a preparative reverse phase C18
column (Deltapak, C18, 15m, I.D. 300 (cid:5) 19 mm) using acetonitrile–
water linear gradient 5–45% acetonitrile (0.1%TFA)/water (0.1%
TFA) at a ﬂow rate of 4 ml min(cid:2)1 over 25 min. The puriﬁed peptides
were reinjected into an analytical reverse phase C18 column (Phe-
nomenex, C18, 5m, I.D. 250 (cid:5) 4.6 mm) using a acetonitrile–water
linear gradient 5–95% acetonitrile (0.1%TFA)/water (0.1% TFA) at
a ﬂow rate of 1 ml min(cid:2)1 over 25 min. The puriﬁed peptides were
veriﬁed by mass spectroscopy (Applied Biosystems QStar (Q-TOF)).
Self-assembly was initiated by dissolving 1 mg of a puriﬁed peptide
in 50 ml of hexaﬂuoroisopropanol (HFIP) (Sigma) and by subsequent
addition of 1 ml of double distilled water. Samples were aged for 2–
24 h prior to their characterization by TEM, CD and DLS.
Spectra were collected on a Perkin-Elmer Spectrum BX FTIR
spectrometer. The assembled peptide samples were spotted on
a CaF2 window and air-dried at 99% relative humidity. The samples
were then rehydrated with D2O, and hydrogen–deuterium exchange
was monitored every minute for 10 min in the spectral range of 1600–
1700 cm(cid:2)1. The FTIR spectra were smoothed with a smoothing
length of 20 units. Two-dimensional correlation spectroscopy (2D
COS) maps were generated from the stack of spectra using the freely
available software package 2D-Shige. All spectral assignments were
done according to published reports.
Published on 11 January 2011. Downloaded on 3/21/2024 2:58:10 PM. View Article OnlineNetherlands). Photomicrographs were digitally recorded using
a Megaview III (SIS, Germany) digital camera.
Light scattering studies were performed in Photocor complex.
Dynamic light scattering experiment was carried out at 37 (cid:4)C and an
angle of 90(cid:4) using 632 nm laser and Photocor-FC digital correlator
operating in multiple-tau mode. The correlation curves were ﬁtted
using DYNAL-S (intensity distribution analysis) for the estimation of
hydrodynamic radius. For measuring the thermal stability of the
assemblies, the samples were prepared at the peptide concentration of
1 mg ml(cid:2)1, vigorously sonicated and were subsequently diluted to an
effective peptide concentration of 0.18 mg ml(cid:2)1 at which the
suspension was stable for the duration of data collection. Tempera-
ture dependent scattering intensity and hydrodynamic radius were
measured and plotted.
Cha–Cha was dissolved in d6-DMSO to a concentration of 6 mg
ml(cid:2)1. The monomeric nature of the preparation was ascertained using
DLS. To assemble the peptide, D2O was added to a ﬁnal concen-
tration of 20%. Self-assembly of Cha–Cha under these conditions was
again ascertained using DLS. All NMR experiments were carried out
at 298 K on a Bruker Avance III spectrometer equipped with 5 mm
cryogenic triple-resonance probes, operating at a ﬁeld strength of
500 MHz. One-dimensional 1H NMR spectra were measured with 32
scans with a relaxation delay of 2 s. Free induction decays (FIDs)
were collected with a spectral width of 8012.82 Hz and an acquisition
time of 2.045 s (t1max). All NMR spectra were processed using
Topspin 2.1 (Bruker AG).

[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": "98/2 HFP/H2O", "peptide_concentration(mg/ml)": "1.0", "PH": "3.5", "temperature(celsius)": "25.0", "heating_cooling": "No", "incubation_period(min)": "75.0"}


[System] polypeptide self-assembly literature information extraction

[User]
BACKGROUND: Gene therapy is most effective when delivery is both efficient and
safe. However, it has often proven difficult to find a balance between
efficiency and safety in case of viral or polymeric vectors for gene therapy.
Peptide based delivery systems may be attractive alternatives but their relative
instability to proteolysis is a major concern in realizing their potential
application in biomedical sciences. In this work we report gene delivery
potential of nanoparticles (Nps) synthesized from cationic dipeptides containing
a non-protein amino acid α, β-dehydrophenylalanine (∆Phe) residue.
METHODS: Dipeptides were synthesized using solution phase peptide synthesis
method. Nps were formed using self-assembly. Nps were characterized using light
scattering, electron microscopy. Transfection efficiency was tested in
hepatocellular carcinoma (HuH 7) cells.
RESULTS: The cationic dipeptides condensed plasmid DNA into discrete vesicular
nanostructures. Dipeptide Nps are non-cytotoxic, protected the condensed DNAs
from enzymatic degradation and ferried them successfully inside different types
of cells. GFP encoding plasmid DNA loaded dipeptide Nps showed positive
transfection and gene expression in HuH 7 cells.
CONCLUSIONS: The cationic dipeptide Nps can successfully deliver DNA without
exerting any cytotoxic effect. Owing to their simple dipeptide origin, ease of
synthesis, enhanced enzymatic stability as well unmatched biocompatibility,
these could be successfully developed as vehicles for effective gene therapy.

Materials
Materials used in this study are: THF, NMM, DMF, pipe-
ridine, DIPCDI, IBCF, TIPS, TFA, HFIP, phenol, DL-threo-
phenylserine, MTT (Sigma-Aldrich, Munich, Germany)
Boc-Arg(Mtr)-OH,
Boc-Lys(Boc)-OH (Novabiochem,
Merck, Darmstadt, Germany) sodium acetate, ethyl ace-
tate, acetonitrile (Spectrochem Pvt Ltd, Mumbai, India),
anhydrous sodium sulfate, citric acid (Merck, Munich,
Germany), human ovarian cancer (HeLa), hepatocellular
carcinoma (HuH-7) and human dermal fibroblast (L929)
cells were obtained from American Type Culture
Peptide synthesis
Dipeptides were synthesized at a scale ranging from
10–30 mM by procedures described earlier [18,19] and as
provided in the Additional file 1: Information S1.
Assembly of dipeptides
Assembly of dipeptides into Nps in water was done from
their HFIP stock solutions at a concentration of 2 mg/ml.
Briefly, 2 mg of cationic peptides were first dissolved in
20 μl of HFIP and then diluted with 1 ml of filtered MilliQ
water to form self-assembled Nps.
Dynamic Light Scattering (DLS) studies
Light scattering studies were performed in Photocor com-
plex (Photocor, Moscow, Russia) using multiple tau digital
correlator. For DLS measurement samples were prepared
in dust free environment. Experiments were carried out at
an angle of 90° using 632 nm laser at room temperature.
Data represent mean of three different sets.
Loading DNA in preformed peptide Nps
Arg-ΔPhe and Lys-ΔPhe, Nps (2 mg/ml) were prepared
as described above and were titrated with increasing
concentrations of plasmid DNA (25, 37.5, 50, 62.5, 75,
100 μg) and analyzed using DLS to determine the
optimum DNA concentration needed to form compact
nanostructures.
Determination of entrapment efficiency of DNA in Arg-
ΔPhe and Lys-ΔPhe Nps
Plasmid DNA was first labeled with Pt-conjugated Alexa-
488 dye using Ulysis DNA-labeling kit (Sigma-Aldrich,
Munich, Germany) and then added to Nps (100 μg of
plasmid DNA : 1 mg/ml of peptide Nps). Samples were in-
cubated for 30–60 min, filtered through a filter plate with
10 kDa membrane cut-off (Millipore, Corp., Billerica, MA,
USA) by centrifuging at 2800 rpm for 20 min at 25°C. Fil-
trates were analysed spectrophotometerically to measure
the amount of free DNA and quantified for percentage en-
trapment of the nucleotides in Nps as described above by
using the following formula:
Effect of plasmid DNA size on their interaction with
Arg-ΔPhe and Lys-ΔPhe
Peptides (2 mg/ml) were incubated with 100 μg of plas-
mids with varying sizes for 30 min. The DNA-peptide
Nps so obtained were analyzed using DLS and electron
microscopy.
Transmission Electron Microscopy (TEM)
DNA-peptide Nps were adsorbed on a 300 mesh copper
grid with carbon coated formvar support and stained
with 1% uranyl acetate and viewed under transmission
electron microscope operating at an accelerated voltage
of 120 kV (Tecnai 12 BioTWIN, FEI Netherlands). Pho-
tomicrographs were digitally recorded using a Megaview
II (SIS, Germany) digital camera. Image analysis was car-
ried out using Analysis II (Megaview, SIS, Germany) and
ImageJ (http://rsb.info.nih.gov/ij/) software packages.
Cellular uptake of DNA-peptide Nps
For estimating the cellular uptake of DNA-peptide Nps,
plasmid DNA and peptide Nps were labeled with Alexa-
488 using ULYSIS DNA labeling kit
(Sigma-Aldrich,
Munich, Germany) and NHS-modified Alexa-588 dyes
(Invitrogen, Life Technologies Corp. NY, USA) respect-
ively. DNA-peptide Nps were prepared as described above.
HeLa cells were first cultured in tissue culture dishes in
RPMI-1640 media, trypsinized and seeded in 6 well plates
(1 x 105 cells/well). After 12 hrs of culture, cells were incu-
bated with plasmid DNA, peptide Nps, DNA-peptide Nps
for 18 hrs, washed with serum and phenol red free media
and imaged under fluorescence microscope.
Nuclear localization of DNA-peptide Nps
HeLa cells were seeded on cover slips placed in 6-well
plates, allowed to grow for 12 hrs for cell adherence and
spreading and subsequently incubated with DNA pep-
tide Nps carrying Alexa-488 labeled DNA for 12 hrs.
After this, media was removed, cells were treated with
DAPI for nucleus staining, washed and visualized under
fluorescent and bright-field microscopes.
Agarose gel electrophoresis of DNA peptide Nps
Complex formation between Nps and plasmid DNA was
analyzed by 1% agarose gel electrophoresis. Free plasmid
DNA (4 μg) and DNA peptide Nps complexes with pep-
tide/DNA ratio of 20/1 (4 μg DNA/80 μg of peptide in
40 μl nuclease free water) were run on a 1% (w/v) aga-
rose gel with TBE buffer (89 mmol/l Tris, 89 mmol/
l boric acid, and 2 mmol/l EDTA pH 8.0) at 75 V for
90 minutes. DNA was visualized by staining gels with
ethidium bromide (0.5 g/l) and images were acquired
using an UV trans-illuminator (Vilber, Lourmat, France).
Stability of DNA peptide Nps towards enzymatic degra-
dation was determined by incubating plasmid DNA
(4 μg) and DNA peptide Nps complexes with peptide/
DNA ratio of 20/1 (4 μg DNA/80 μg of peptide in 40 μl
nuclease free water) with 5 μl of DNase I solution (10 μg/
ml in DNase/Mg2+ digestion buffer, which consisted of
50 mM Tris–HCl, pH 7.6, and 10 mM MgCl2) each at
37°C for 30 min, and degradation of plasmid DNA was
analyzed by 1% agarose gel electrophoresis.
Cytotoxicity of DNA-peptide Nps
HeLa and L929 cells were cultured in RPMI-1640 and
DMEM supplemented with 10% FBS, and maintained on
TCTP plates at 37°C in a humified atmosphere containing
5% CO2, till 70-80% confluency. Cells were then harvested
by trypsinization, washed and resuspended in RPMI-1640
/DMEM medium (supplemented with 10% FBS). Live cells
were counted by trypan blue dye exclusion test using a
hemocytometer, diluted in completed media, seeded in 96
well plates (2 x 104 cells/well). After 12 hrs, cells were in-
cubated with 20 μl each of dipeptide and DNA-dipeptide
Nps (2 mg/ml) for a period of 24 hrs. As a control, cells
were incubated with 20 μl of PBS. MTT assay was used to
determine the cell viability of treated cells. The percentage
viability was calculated using the equation given below:
Percentage viability of cells = [(Absorbance of MTT
formazan produced by cells grown in presence of nps /Ab-
sorbance of MTT formazan produced by cells grown on
TCTP control plate)] × 100.
In vitro transfection efficiency of DNA peptide Nps
HuH-7 cells were cultured in DMEM supplemented with
10% FBS and 1% pencillin-streptomycin and maintained
on TCTP plates at 37°C in a humified atmosphere
containing 5% CO2, till 70-80% confluency. 24 h prior to
transfection cells were seeded in 6 well plates (2 x 105).
DNA-Nps were prepared by incubating 4 μg of EGFP
encoding plasmid DNA with 80 μg each of dipeptides
Lys-ΔPhe and Arg-ΔPhe in 40 μl of nuclease free water.
Nps so formed were visualized under TEM to determine
particle size and morphology as described above. On the
day of transfection, the culture medium in each well was
replaced with 1 ml of complete medium containing 1 μg
of DNA loaded in DNA-peptide Nps at (1: 200 DNA to
peptide ratio) and 1 μg of DNA in lipofectamine. The cells
were then incubated for 48 hrs, washed to remove excess
Nps and DNA and analysed using fluorescence micros-
copy for GFP expression.

[Assistant] [{"peptide_sequence": "KF", "self-assembly_phase": "particle", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "Delta-F", "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)": "nan"}, {"peptide_sequence": "RF", "self-assembly_phase": "particle", "N-terminal_modification": "Free", "C-terminal_modification": "Free", "Non-terminal_modification": "Delta-F", "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)": "nan"}]


[System] polypeptide self-assembly literature information extraction

[User]
DI) and
zeta-potential of DNPs before and after ligand
conjugation was analyzed using a dynamic light
scattering (DLS) technique (Zetasizer Nano ZS90;
Malvern Instruments Ltd, Malvern, UK) at an
angle of 90° and at RT, using a 632-nm laser.
The  mean  of  readings  from  three  different
batches was considered as actual particle size and
zeta-potential.
n Stability of DNPs
For assessing stability, RDF NPs (1 mg/ml) were
diluted with filtered Milli-Q® water (Millipore
n Drug entrapment
DOX HCl (Sigma-Aldrich, Munich, Germany)
was mixed with preformed NPs in an equal weight
ratio, incubated for 48 h at RT with constant
shaking  and  then  ultracentrifuged  at  120,000
rpm  for  4  h  at  10°C  (OptiMax™  MLA-130
rotor; Beckman Coulter, IN, USA) to separate
the unentrapped drug molecules. Concentration
of unbound drug in the supernatant was quan-
tified using UV-visible spectroscopy at 480 nm
(Lambda  35;  PerkinElmer,  MA,  USA)  and
results were expressed as percentage entrapment,
determined using equation 1:
n Derivatization of DNPs with FA
NPs were prepared at desired concentrations, and
conjugated with the carboxylic acid group of FA
by  the  standard  1-ethyl-3-(3-dimethylamino-
propyl)carbodiimide/N-hydroxysuccinimide
(EDC/NHS)  activation  method  [18,19],  with
minor modification (Supplementary method S3).
n Cytotoxicity of DOX-loaded NPs
HeLa  and  MDA-MB-231  cells  were  plated
(2 × 104 cells/well) in triplicates in 96-well, ster-
ile microtiter plates (Corning, MA, USA) and
allowed to grow for 12 h, allowing proper cell
adhesion. Cells were then treated with DOX,
DOX-loaded  RDF  (DOX-RDF)  and  DOX-
loaded FA-RDF (DOX-FA-RDF) NPs contain-
ing various concentrations of DOX (1, 5, 10 and
future science groupReseaRch aRticlePanda, Kaul, Kumar et al.20 µg/ml) and incubated for 24 h, before cell
viability assays. Cells were also treated with 20
µl of phosphate-buffered saline as control. At the
end of the incubation period, cells were washed
with complete media and cell viability was deter-
mined using the 3-(4,5-dimethylthiazol-2-yl)-
2,5-diphenyltetrazolium bromide assay.
n Wound migration assay
The assay was performed as previously described
[20]. Briefly, MDA-MB-231 cells were seeded and
grown  to  confluency.  Wounds  with  constant
diameter were made using sterile tips and then
treated with DOX, DOX-RDF and DOX-FA-
RDF NPs having various DOX concentrations (1,
5 and 10 µg/ml) and incubated for 24 h. Wound
photographs of the same scratch at 0 and 18 h were
taken under a microscope (Nikon, NY, USA).
n Anti-tumor activity of DOX-FA-RDF
NPs in EAT-bearing mice
For tumor regression studies, mice bearing EATs
were randomly divided into five groups (n = 10 in
each group) and intravenously injected (through
the  tail  vein)  with  100  µl  of  DOX  solution
equivalent to 5 mg/kg bodyweight of the animal
[12], 100 µl of DOX-RDF (stock: 4 mg/ml) and
DOX-FA-RDF NPs (stock: 2.5 mg/ml) contain-
ing an amount of DOX equivalent to 5 mg/kg
bodyweight of the animal, every fourth day for
a  period  of  3  weeks.  Void  RDF  and  FA-RDF
NPs (1 mg/ml) were used as the vehicle control.
Tumor sizes were measured using calipers and
tumor  volume  was  calculated  using  equation  2,
where  L  is  the  length  and  W  is  the  width  of
the tumor:
n Toxicity studies (in vivo)
Animals were intravenously administered eight
times with 100 µl of DNPs (1 mg/ml; equivalent
to therapeutic dose) and monitored continuously
for any behavioral change and bodyweight loss.
After these animals were sacrificed, organs were
excised, sectioned and studied for morphological
changes by hematoxylin and eosin staining.
Results & discussion
n Selection of the most suitable DNPs
for drug delivery
Amphipathic dipeptides such as EDF, KDF and
RDF formed discrete nanostructures at differ-
ent  concentrations  [21,22].  Since  self-assembly
of  short  peptides  is  generally  a  concentration-
dependent phenomenon, the authors reinvesti-
gated self-assembly of the four dipeptides, EDF,
KDF, RDF and DDF, at different concentrations.
Dipeptides were synthesized in high yields, puri-
fied  using  HPLC  and  characterized  by  mass
spectrometry (Supplementary table S1). Self-assembly
was initiated by dissolving different amounts of
dipeptides (1–10 mg) in HFIP and subsequently
adding 1 ml of water. The formation and stabil-
ity in population was investigated by DLS and
TEM.  RDF  showed  remarkable  self-assembly,
characterized  by  high  counts  per  second  and
low PDI at a concentration of ≥1 mg/ml, and
formed NPs with a mean diameter of approxi-
mately 240 ± 10 nm (table 1 & Figure 1a) and a zeta-
potential of 40 ± 2mV (Figure 1a). KDF and EDF
also self-assembled, but at a much higher con-
centration than RDF; at 5 mg/ml, KDF formed
NPs with a mean diameter of 400 ± 10 nm, while
EDF formed NPs of approximately 370 ± 5 nm,
but only at a concentration of 10 mg/ml (table 1).
DDF formed gel in HFIP. Minor changes in the
charge and molecular mass, contributed here only
by  the  N-terminal  residues,  had  a  remarkable
www.futuremedicine.comfuture science groupModified dipeptide-based nanoparticles: vehicles for targeted tumor drug deliveryReseaRch aRticleTable 1. Self-assembly (hydrodynamic diameter, polydispersity index and particle count rate) of dipeptides at
different concentrations determined using dynamic light scattering.
Count rate
Values represent the mean of readings from three different batches ± standard deviation.
EDF: Glu–a,b-dehydrophenylalanine; KDF: Lys–a,b-dehydrophenylalanine; Kcps: Kilo counts per second; ND: Not detectable; PDI: Polydispersity index;
RDF: Arg–a,b-dehydrophenylalanine.
effect on the self-assembly of these dehydrodipep-
tides. Results of TEM corroborated with those
of  DLS  studies.  At  1  mg/ml,  RDF  assembled
into well-defined vesicular nanostructures with a
Figure 1. Hydrodynamic diameter and zeta-potential measurements of
Arg–a,b-dehydrophenylalanine nanoparticles. Hydrodynamic diameters of
Arg–a,b-dehydrophenylalanine nanoparticles at concentrations of (A) 1 and
(B) 0.25 mg/ml. (C) Zeta-potential measurements of Arg–a,b-dehydrophenylala nine
nanoparticles at a concentration of 1 mg/ml.
mean diameter of 30 nm (Figure 2a), whereas KDF
and EDF did not show significant assembly at this
concentration (Figure 2a), although at a higher con-
centration of 5 mg/ml and 10 mg/ml, they formed
pleomorphic  vesicular  structures  with  mean
diameters of 370 and 110 nm, respectively [22].
Concentration-dependent  self-assembly  of
peptides has been reported earlier  [8,23], but the
dipeptides described here represent a remarkable
example of minor structural changes leading to
significant differences in self-assembly behavior.
Gorbitz and others have also shown that small
changes at the N-termini of dipeptides resulted in
different packing arrangements, probably leading
to differential assembly [24,25]. Since RDF assem-
bled into stable NPs at a relatively low concen-
tration (0.25 mg/ml), further investigations were
mostly focused on this dipeptide. RDF showed
a  concentration-dependent  assembly  (Figure  1b)
with a critical aggregation concentration below
1 mM. Such a low assembling concentration of
RDF  compares  well  with  other  peptide-based
NPs shown to have drug-delivery potential  [12].
NPs intended for drug delivery would inevitably
be diluted significantly upon in vivo delivery; the
stability of RDF NPs under different dilutions was
investigated using DLS. It was observed that the
NPs, once formed, were stable up to 100-fold dilu-
tions and retained their particulate nature with a
size of 203 nm and a PDI of 0.3 (table 2). RDF NPs
were also stable in size, as well as polydispersity, at
pH 7.4 over a 24-h period (Figure 2b).
Stability of NPs in serum is essential for their
application as a drug-delivery vehicle. Moreover,
since RDF is cationic in nature, its interaction
with serum proteins may lead to charge shielding
and aggregation. In most studies, serum stabil-
ity of NPs has been carried out in the presence
of  10%  (v/v)  fetal  bovine  serum  [26,27],  mainly
because whole serum contains many proteins and
protein aggregates that interfere with the DLS
Figure 2. Electron microscopy and in vitro stability of dipeptide nanoparticles.
(A) Transmission electron microscope images of amphipathic dipeptide nanoparticles.
(Ai) Arg–a,b-dehydrophenylalanine (Arg–DPhe) nanoparticles, (Aii) Lys–DPhe nanoparticles and
(Aiii) Glu–DPhe nanoparticles. Only Arg–DPhe formed discrete nanovesicles with a mean diameter of
30 nm. (B) Stability of Arg–DPhe nanoparticles over 30 days. (C) Stability of Arg–DPhe nanoparticles
in presence of 1 and 10% FBS determined for 24 h. Error bars represent mean ± standard
deviation (n = 3).
FBS: Fetal bovine serum.
measurement. Hence, the serum stability of RDF
NPs was assessed in the presence of 10% (v/v) fetal
bovine serum using DLS. For a period of 6 h, no
significant changes were seen in the particle size
of RDF NPs (except for the formation of 2–5%
large aggregates) (Figure 2C), as obtained for other
nanoparticulate systems  [26,27]. However, after a
period of 24 h, the NPs showed a twofold increase
in their overall particle size, possibly due to interac-
tion with serum proteins. However, the fact that
they still retain their nanodimension demonstrates
the suitability of these systems for cellular delivery.
0.2 ± 0.1
0.3 ± 0.09
0.3 ± 0.03
0.2 ± 0.07
0.3 ± 0.05
0.8 ± 0.1
Nanoparticles were diluted with water. Size and PDI was
measured using dynamic light scattering. Values represent
the mean of readings from three different
batches ± standard deviation.
PDI: Polydispersity index.
www.futuremedicine.comfuture science groupModified dipeptide-based nanoparticles: vehicles for targeted tumor drug deliveryReseaRch aRticle
(Figure 3) [18], and the conjugation (with efficiency
of 50 ± 5% w/w) was characterized by HPLC and
mass  spectrometry  (Supplementary  FigureS  S1  &  S2).
TEM showed that RDF NPs were stable to con-
jugation and retained their vesicular structure
with a mean diameter of 50 nm (Figure 4a). DLS
studies showed that the functionalized NPs had
a mean hydrodynamic diameter of 600 ± 120 nm
(86%) (Supplementary Figure S3) and a zeta-potential
of 20 ± 2 mV (Figure 4b). Although FA conjugation
led to an increase in the hydrodynamic diameter
of the NPs, they were still expected to have bet-
ter tumor accumulation through enhanced per-
meability and retention effect, as a recent study
reported that bacteria larger than 1000 nm could
even  show  enhanced  permeability  and  reten-
tion effect and high tumor accumulation [33,34].
Moreover, as conjugation of specific ligands to
Figure 3. Conjugation of folic acid to dipeptide nanoparticles for tumor-targeted drug delivery.
DNP: Dipeptide nanoparticle; EDC: 1-ethyl-3-(3-dimethylaminopropyl)carbodiimide; FA: Folic acid; NHS: N-hydroxysuccinimide.
Figure 4. Electron microscopy, surface charge, cytotoxicity and cellular uptake of folic acid-conjugated Arg– a,b-
dehydrophenylalanine nanoparticles. (A) Transmission electron microscopy image of folic acid-conjugated Arg–RDF
nanoparticles (NPs). NPs retained their vesicular structure even after surface modification with the ligand. (B) Zeta-potential of
FA-RDF NPs. (C) Fluorescent microscopic image showing uptake of RDF and FA-RDF NPs by HeLa cells. Magnification: 20×.
(Ci & Cii) RDF, (Ciii & Civ) FA-RDF and (D & E) FACS analysis showing uptake of RDF and FA-RDF NPs by HeLa cells and
MDA-MB-231 cells. (F) Cytotoxicity of RDF and FA-RDF NPs. Error bars represent mean ± standard deviation (n = 3).
*p < 0.005 versus R DF NPs; **p <  0.05 versus R DF NPs; ***p > 0.05 versus R DF NPs.
BF: Brightfield image; Con.: Untreated control; FA: Folic acid; FA-RDF: Folic acid-conjugated Arg–a,b-dehydrophenylalanine;
FI: Fluorescence image; FL1-H: Fluorescence channel 1 height; HEK: Human embryonic kidney cell;
RDF: Arg–a,b-dehydrophenylalanine.
www.futuremedicine.comfuture science groupModified dipeptide-based nanoparticles: vehicles for targeted tumor drug deliveryReseaRch aRticle
NPs for targeted delivery is often complex and
involves multiple steps, the ease of conjugation
of ligands such as FA to free DNPs makes them
an attractive platform for delivery purpose [35,36].
The serum stability study of the derivatized NPs
demonstrated that the particles were almost stable
up to the 6-h time point, with a minor increase
in their mean hydrodynamic diameter. However,
similar to RDF NPs, a longer incubation time led
to particle aggregation with a noticeable increase
in overall particle size (Supplementary Figure S4).
HeLa cells incubated with Alexa Fluor®-labeled
(Life Technologies Corp., CA, USA) FA-RDF
NPs  showed  intense  fluorescence  compared
with  those  treated  with  RDF  NPs,  suggest-
ing higher uptake of FA-RDF NPs  (Figure  4C).
FACS analysis also showed that in cancer cells
overexpressing FA receptors (HeLa and MDA-
MB-231), uptake of FA-RDF NPs was signifi-
cantly  higher  (p  <  0.005)  than  underivatized
RDF NPs (77 vs 44% in MDA-MB-231; 49 vs
11% in HeLa) (Figure 4d & e). No significant dif-
ference was observed in uptake of the two NPs in
MCF-7 cells that do not overexpress FA receptors
(Supplementary Figure S5). However, in the presence
of 5 mM FA in the incubation medium, uptake
of FA-RDF NPs was reduced by approximately
50% in MDA-MB-231 and almost 35% in HeLa
Figure 5. Efficacy of doxorubicin-loaded nanoparticles in tumor cells. Cytotoxicity of DOX,
DOX-R∆F and DOX-FA-R∆F against (A) HeLa and (B) MDA-MB-231 cells cultured in serum-
containing media, as a function of DOX-equivalent concentrations (0, 1, 5, 10 and 20 µg/ml). Folic
acid-conjugated nanoparticles significantly increased the efficacy of DOX. Error bars represent
mean ± standard deviation (n = 3). (C) Wound migration assay of cells treated with drug-loaded
nanoparticles (wound photographs taken at 0 and 18 h). FA-R∆F nanoparticles increased the potency
of the drug towards inhibiting wound migration of MDA-MB-231 cells.
*p > 0.05 versus DOX; **p < 0.05 ver

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