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Supramolecular Hydrogel Formation in a Series of SelfAssembling Lipopeptides with Varying Lipid Chain Length. Valeria Castelletto, Amanpreet Kaur, Radoslaw M. Kowalczyk, Ian W. Hamley, Mehedi Reza, and Janne Ruokolainen Biomacromolecules, Just Accepted Manuscript • Publication Date (Web): 23 May 2017 Downloaded from http://pubs.acs.org on May 27, 2017

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Supramolecular Hydrogel Formation in a Series of Self-Assembling Lipopeptides with Varying Lipid Chain Length.

V. Castellettoa*, A. Kaura, R. M. Kowalczyk a, I.W. Hamleya, M. Rezab, J. Ruokolainenb. a

School of Chemistry, Pharmacy and Food Biosciences. University of Reading,

Whiteknights, Reading RG6 6AD, United Kingdom. b

Department of Applied Physics, Aalto University School of Science, Aalto FI-00076,

Finland.

* Author for correspondence.

ACS Paragon Plus Environment

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Abstract

The self-assembly in aqueous solution of three lipopeptides comprising a bioactive motif conjugated at the N terminus to dodecyl, tetradecyl or hexadecyl lipid chains has been examined. The bioactive motif is the peptide block YEALRVANEVTLN; a C-terminal fragment of the lumican proteoglycan. This study was motivated by our previous studies on the hexadecyl homologue C16-YEALRVANEVTLN which showed aggregation into β-sheet structures above a critical aggregation concentration (cac), but most remarkably we found that these aggregates were stable to dilution below the cac1. Here we find that the C12- and C14- homologues also self-assemble above a cac into β-sheet nanotapes based on bilayer packing. The cac decreases with increasing lipopeptide hydrophobicity. Unexpectedly, the β-sheet secondary structure is present upon dilution and the aggregates are thermally stable. These results indicate that the dilution trapping of β-sheet secondary structure is not associated to lipid chain melting behaviour. Instead we associate it with pH-dependent favourable inter-molecular electrostatic interactions. Investigation of the pH-dependence of aggregation led to the discovery of conditions for formation of lipopeptide hydrogels (initial sample preparation at pH 10 in NaOH solution, followed by reduction to pH ~1 by addition of HCl). The lipopeptide hydrogels comprise networks of bilayerbased peptide nanotape bundles and to our knowledge this type of hydrogel is unprecedented. These hydrogels may have future applications based on processes such as encapsulation and release that involve fast switches between solution and hydrogel nanostructures.


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Introduction Lipopeptides combine peptide and lipid motifs to create surfactant-like molecules with tunable aggregation properties and nanostructures in aqueous solution.2 The peripherally attached peptide units enable the incorporation or stimulation/modulation of bioactivity and this has led to a diversity of demonstrated applications with potential biomedical relevance.3 Lumican is an extracellular matrix proteoglycan with an important role in the structure of the cornea. In previous papers, our group in collaboration with that of C. J. Connon has investigated the self-assembly4 and bioactivity1 of peptide amphiphiles incorporating a C-terminal fragment YEALRVANEVTLN from the lumican proteoglycan attached to a hexadecyl (palmitoyl) chain. This lipopeptide selfassembles, above a critical aggregation concentration (cac) into β-sheet based nanotape structures comprising interdigitated lipopeptide bilayers. The aggregated form was found to be stable on dilution and in this form, a two-fold increase in collagen production per cell was observed at low concentration (below the cac) compared to a directly dissolved sample at the same concentration.1 This work motivated the present study in which we investigate the influence of alkyl chain length on the self-assembly of lipopeptides comprising the YEALRVANEVTLN lumican-derived peptide and C12, C14 and C16 lipid chains. There have been few studies on the effect of alkyl chain length on the selfassembly of peptide amphiphiles. Meijer et al. investigated the effect of lipid chain length on the secondary structure and thermal stability of peptide amphiphiles incorporating the KTVIIE peptide.5 Samples with C12, C14 and C16 tails showed thermally stable β-sheet structures (as probed by CD spectroscopy) whereas C8 and C10 functionalized lipopeptides showed a loss of secondary structure on heating. Apart


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from the C6 PA, all other peptide assemblies were found to be stable on dilution down to low micromolar concentrations. The peptide itself showed a loss of β-sheet structure below 0.06 mM.5 This group also investigated the conformation of CnGANPNAAG lipopeptides with n = 0 to 16 (the peptide is a bioactive sequence derived from a malaria parasite protein).6 C16-GANPNAAG lipopeptide showed stable β-sheet structure up to 90 oC. In contrast, the C6, C10 and C12 samples exhibited a random coil conformation. The C14 lipopeptide underwent a transition from β-sheet to random coil structure on heating.5 Xu and co-workers have studied the self-assembly behaviour of a series of lipopeptides with C9, C11, C13 and C15 chains and a VRGDV peptide motif.7 All lipopeptides

formed β-sheet nanofibers in aqueous solution at pH 7, although

adjustment of pH was used as a means to influence nanostructure formation and the two lipopeptides with shorter alkyl chains formed spherical micelles at pH 11 (coexisting with fibrils in the case of the C11 PA). The influence of hydrophobic chain length on DNA transfection was studied for non-linear gemini surfactants comprising two lipid chains attached to a Lys-spacer-Lys type headgroup (the spacer length was also varied).8 Optimal properties were observed for lipopeptides with oleoyl chains and an n= 6 spacer. The self-assembly properties of the molecules were not systematically examined although the molecule with optimal properties was found to form liposomes. An extensive study of the alkyl tail modification influence on the selfassembly of peptide-amphiphile molecules was undertaken by Hartgerink, Stupp and co-workers.9 In that work, pH control, divalent ion addition and concentration were defined as the three tools for the self-assembly of peptide fibres. In particular, the family of molecules corresponding to the CCCCGGGS(PO4)RGD motif with C6 C10,


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C16 or C22 chains was used to illustrate the reversibility of self-assembly and covalent capture in those molecules, based on pH and oxidation state. Here, we study the secondary structure and aggregation properties of C12YEALRVANEVTLN

(1),

C14-YEALRVANEVTLN

(2)

and

C16-

YEALRVANEVTLN (3). The original aim was to investigate the influence of shorter alkyl chains on the self-assembly and kinetic stability of aggregates, motivated by our previous studies on 3,

1, 4

especially in regard to the interesting observation that pre-

aggregated self-assemblies (nanotapes) could be diluted below the cac and remained kinetically stable against the presumed equilibrium monomeric state at low concentration. Here, we explore the nature of the stability to dilution of aggregates of three lipopeptides with three different lipid chain lengths as this was expected to have a significant influence on molecular packing and mobility. We have also unexpectedly discovered conditions under which it is possible to form gels of these lipopeptides. Gel formation by lipopeptides comprising alkylated peptides has been reported for nanofiber-forming lipopeptides when the fibrils form a sample-spanning network,9-10 although it has not been reported previously for bilayer-based lipopeptide assemblies to our knowledge. In this work, a combination of spectroscopy, microscopy, scattering techniques, together with rheology are used to provide new insight into the self-assembly of lipopeptides 1-3 and their hydrogel formation.

Methods Peptides. Lipopeptides 1-3 were custom synthesized by CS Bio (USA). Lipopeptides 1 and 2 were received as ammmonium salts, while lipopeptide 3 was received as an acetate salt. The counterions present are based on the purification process. While lipopetides 1 and 2 were purified once using ammonium hydroxide buffer to obtain


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target purity, 3 was purified with ammonium hydroxide as a first step, then repurified with acetic acid buffer in a second step and finally purified with ammonium hydroxide buffer as a third step. As a result, 3 contains 1.5 % acetate while 1 and 2 contain only 0.08 and 0.11% ammonium ions respectively. The purity of samples 1, 2 and 3 was determined by the supplier to be 95.15%, 95.47% and 95.01% respectively, as determined from HPLC in ammonium hydroxide (0.1 % NH4OH in water/ acetonitrile gradient). Our own electro-spray ionization mass spectroscopy (ESI-MS) analysis (spectra in Figure S1a-c) show that the measured molecular weight was 1674.96, 1702.99 and 1731.02 Da (expected: 1673.96, 1702.01 and 1730.05 Da) for 1, 2 and 3 respectively. PA solutions were prepared by mixing weighed amount of peptide powder with water and concentrations were calculated as wt%. Solutions were then heated to 55 ̊C for 20 minutes with intermittent vortex mixing at 1800 rpm for 30 seconds. PA solutions prepared in this way were allowed to rest for at least 3 hours before the experiments. Since the molar mass values are within 3% of each other (1.5% deviation from the mean), our data comparing characteristics of the lipopeptides at fixed wt% concentration are essentially also comparisons at fixed molarity. For example the molarities of 1 wt% solutions of 1, 2 and 3 are 5.97 mM, 5.87 mM and 5.78 mM which are the same within uncertainty. Mass Spectroscopy. Electrospray-ionization mass spectra were recorded using a Thermofisher Obitrap XL instrument. Samples, which were presented as 1 mg/ml, were diluted 33 fold (30 µL sample + 970 µL MeOH). The two LCMS mobile phase buffers (water and acetonitrile) included 0.1% formic acid to assist the reverse phase separation and to aid protonation.


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Zeta Potential Measurements. The zeta potential was measured using a Zetasizer Nano zs from Malvern Instruments. 1 ml of sample was placed inside a disposable folded capillary cell. The sample was left to equilibrate for 120 sec before measuring the zeta potential, using an applied voltage of 50.0 V. The results presented are the average over three measurements. Fluorescence Assays. ThT fluorescence depends on the formation of amyloid-like structures11 (β-sheet fibrils) and is used to detect amyloid formation by peptides. For the ThT assay, the spectra were recorded from 460 to 600 nm using an excitation wavelength λex= 440 nm, and the peptide was dissolved in a 5.0×10-3 wt % ThT solution. Circular Dichroism (CD) Spectroscopy. CD spectra were recorded using a Chirascan spectropolarimeter (Applied Photophysics, UK). Peptide solutions were placed in a quartz cover slip cuvette (0.1 mm thick), or in a quartz bottle (1 mm path length). Spectra are presented for absorbance A < 2 at any measured point with a 0.5 nm step, 1 nm bandwidth, and 1 s collection time per step. The CD signal from the water background was subtracted from the CD data of the sample solutions. Where data has been smoothed, the smoothed trace was calculated in a way that no distortions were observed in the residual trace. Fourier Transform Infra-Red (FTIR) Spectroscopy. Spectra were recorded using a Nexus-FTIR spectrometer equipped with a DTGS detector. Liquid samples were measured using a transmission configuration with a PEARL liquid cell, or with the sample placed between two CaF2 plate windows (spacer 1.2×10-2 mm thick). Spectra were scanned 128 times over the range of 900-4000 cm-1. Rheology. Rheological properties were determined using a controlled stress TA Instruments AR-2000 rheometer (TA Instruments). Experiments were performed


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using a plate-and-plate geometry (plate radius= 20 cm; gap= 1000 µm). The linear regime of the sample was first identified from the dependence of the storage (G’) and loss (G’’) moduli on the oscillatory stress at a fixed frequency of 6.283 rad s-1. The dependence of G’ and G’’ on the oscillatory frequency was then determined at a fixed oscillatory stress within the linear regime. X-ray Diffraction (XRD). Measurements were performed on stalks prepared by drying a drop of solution or a thread of hydrogel suspended between the ends of waxcoated capillaries. The stalks were mounted onto the four axis goniometer of an Oxford Diffraction Gemini Ultra instrument. The sample-detector distance was 44 mm. The X-ray wavelength was λ = 1.54 Å. The wavenumber scale (q = 4πsinθ/λ, where 2θ is the scattering angle) was geometrically calculated. The detector was a Sapphire CCD. Cryogenic-Transmission Electron Microscopy (Cryo-TEM). Imaging was carried out using a field emission cryo-electron microscope (JEOL JEM-3200FSC), operating at 200 kV. Images were taken in bright field mode and using zero loss energy filtering (omega type) with a slit width of 20 eV. Micrographs were recorded using a Gatan Ultrascan 4000 CCD camera. The specimen temperature was maintained at -187 oC during the imaging. Vitrified specimens were prepared using an automated FEI Vitrobot device using Quantifoil 3.5/1 holey carbon copper grids with a hole size of 3.5 µm. Just prior to use, grids were plasma cleaned using a Gatan Solarus 9500 plasma cleaner and then transferred into the environmental chamber of a FEI Vitrobot at room temperature and 100 % humidity. Thereafter 3 ml of sample solution was applied on the grid and it was blotted twice for 5 seconds and then vitrified in a 1/1 mixture of liquid ethane and propane at temperature of -180 °C. The grids with


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vitrified sample solution were maintained at liquid nitrogen temperature and then cryo-transferred to the microscope. Nuclear Magnetic Resonance (NMR). 1H NMR experiments were performed on sample 3 dissolved in DMSO-d6. 1D 1H and

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C as well as 2D COSY, TOCSY,

NOESY, HSQC and HMBC spectra were recorded at room temperature using a Bruker Avance III 700 MHz spectrometer (16.44T; equipped with a four channel cryoprobe) and a Bruker Avance III 500 MHz spectrometer (11.74T; standard BBO probe). Each 1H spectrum corresponds to an average of 128 to 1024 transient, while the

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C spectrum required 8192 transients. The indirect (F1) dimension of the 2D

experiments was varied between 256 (COSY) and 1024 (TOCSY, HSQC and HMBC) points, in an attempt to further improve resolution of the overlapped signals in the spectra using the second dimension. Each indirect point in the 2D COSY, TOCSY and HSQC correspond to an average of 32 transients, while an average of 100 transients were measured from HMBC. All spectra were referenced using DMSO-d6 impurity signal as secondary reference at 2.50 ppm (1H) and 39.52 ppm (13C). Small-Angle and X-Ray Scattering (SAXS). Synchrotron SAXS experiments were performed on solutions at beamline BM29 at the ESRF (France), using a BioSAXS robot, and on gels at beamline I22 (Diamond, ESRF). On BM29 solutions were loaded into the 96 well plate of an EMBL BioSAXS robot, and then injected via an automated sample exchanger into a quartz capillary (1.8 mm internal diameter) in the X-ray beam. The quartz capillary was enclosed in a vacuum chamber, in order to avoid parasitic scattering. The sample was injected in the capillary and flowed in front of the X-ray beam during the SAXS data acquisition. BM29 operated with an X-ray wavelength λ = 1.03 Å (12 keV). The images were


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captured using a PILATUS 1M detector, while data processing was performed using dedicated beamline software ISPYB (BM29). On I22, hydrogels were placed in DSC pans modified with glass or Teflon windows to enable transmission of the X-ray beam. The sample to SAXS detector distance was 7.483 m. A Pilatus P3-2M detector was used to acquire the 2D SAXS scattering patterns. SAXS data collected for samples with Teflon windows was corrected for the Teflon background scattering. Diffraction from silver behenate was used to calibrate the wavevector scale of the scattering curve. Data processing was performed using software DAWN (Data Analysis Software group, Diamond Light Source Ltd.). Polarized Optical Microscopy (POM). Images were obtained with an Olympus BX41 polarized microscope by placing the sample between crossed polarizers. Samples were stained with a 0.3 wt% Congo red solution before being placed between a glass slide and a coverslip. Images were captured with a Canon G2 digital camera. Hydrogel formation. The measured pH of a 1 wt% peptide solution in water (pH 6.64, 4.3 and 6.81 for 1, 2 and 3 respectively), was increased to pH 10 by titration of 0.98 M NaOH. The hydrogel was formed by injecting 1 M HCl solution into the peptide solution at pH 10, which immediately gelled on contact with the injected 1 M HCl solution. The amount of 1M HCl injected corresponded to 7.1%, 7.7% and 9.5% of the total final weight for 1, 2 and 3 hydrogels respectively. The resulting pH of the hydrogels was 1.48, 1.33 and 1.1 for 1, 2 and 3 respectively. Several attempts to make a hydrogel omitting the intermediate titration of the pH to 10 proved to be unsuccessful. Scanning Electron Microscopy (SEM). The hydrogel was immersed in a fixative solution containing 2 % paraformaldehyde and 2.5% glutaraldehyde for 30 min. This was followed by gradual dehydration from 10 to 100% ethanol (10, 30, 50, 70, 90,


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100% ethanol), waiting 30 seconds between dehydration steps. The sample was then extracted from the 100% ethanol solution and subjected to critical point drying. The dried sample was placed on a stub covered with a carbon tab (Agar Scientific, UK), and then coated with gold. A FEI Quanta FEG 600 environmental scanning electron microscope (SEM) in high vacuum mode (20 kV high tension) was used to study and record SEM images. Dynamic Light Scattering (DLS). Experiments were performed using an ALV CGS3 system with 5003 multi-digital correlator. The light source was a 20 mW He-Ne laser, linearly polarized, with λ= 633 nm. Scattering angle of 90o was used for all the experiments. Samples were placed standard 0.5 cm diameter cylindrical glass cells.

Results Solutions nanostructure. The isoelectric point of the peptide block in lipopeptides 13 is at pH 0.85 as calculated using web-based software.12 For pH values higher than 0.85, the net charge Z becomes increasingly negative until reaching Z= -2 at pH 7 and Z~ -2.5 at pH 10 (the charge changes in this range due to crossing pKa,2 associated with the α-amino group in the asparagine residue). The software TANGO13 was used to estimate the aggregation propensity into different secondary structures of the residues in the peptide sequence (0.15 wt % peptide, 20 oC)

at pH 7 and pH 10. Figure S2 shows the predictions for the

aggregation state for each residue. There is a predicted tendency for β-sheet formation in the N-terminal residues of the peptide (i.e. EVTLN region) at both pH 7 or pH 10 (Figure S2). Results obtained from this calculation highlight the importance of the amphiphilic nature of the lipopeptides as well as the secondary structure propensity in the self-assembly processes discussed below.


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The pH of solutions in water is ~ 6.7 for lipopeptides 1 and 3 and ~4.3 for lipopeptide 2, at the concentrations studied in this work (Figure S3). We believe that the lipopeptides purification process and consequent counterion concentration has affected the final pH of the solutions introducing the variation measured in Figure S3. A negative charge at the surface of the self-assembled nanostructures is expected for these pH values. Accordingly, the zeta potential measured for 0.1 wt% samples was 56.2±4.7 and -42.2±2.6 mV for 1 and 3 respectively. It was not possible to measure the zeta potential for a 0.1 wt% solution of 2 due to the cloudiness of the sample, caused by the formation of large aggregates in the solution. Increasing the pH of the solutions to 10 through titration of NaOH, a process discussed below in relation to hydrogel formation, preserves the negative charge at the surface of the self-assembled nanostructures. Zeta potential values at pH 10 are -42.9±3.3, -48.9±1.7 and -50.5±1.3 mV for 1, 2 and 3 respectively. NaOH titration reduces the cloudiness of the solution of 2 and allows for zeta potential measurements. The high values of zeta potential measured for the solutions in pure water or with NaOH, indicates the formation of highly charged species in solution. All following experimental results are at native pH, unless pH 10 is indicated. Since zeta potential experiments suggested the formation of self-assembled structures at native pH and pH 10, fluorescence assays were undertaken to determine the cac for the formation of such nanostructures. The cac was determined through a Thioflavin (ThT) assay, which is sensitive to the formation of amyloid fibrils.11,

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Results of the ThT assay for 1, 2 and 3 are shown in Figure 1. Increasing the lipid chain length increases the hydrophobicity of the PA. Consequently, the cac values in Figure 1, obtained from the discontinuity in fluorescence intensity using ThT as a probe of aggregation, decrease with increasing lipid chain length. Lipopeptide 2 is the


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weakest fibrillizer, since the intensity of the ThT fluorescence emission is on average 4x lower for 2 than it is for 1 and 3. In a previous work we reported cac = 0.03 wt% for lipopeptide 3, from pyrene (Pyr) fluorescence assays.4 Here, we show that the ThT assay is better suited to determine cac for lipopeptide 3. In our previous work the Pyr fluorescence was found to unexpectedly decrease with increasing 3 concentration,4 possibly introducing anomalies in the determination of the cac. In addition, the concentration range of the Pyr assay did not extend low enough to cover the cac evident in Figure 1c. We performed CD and DLS experiments to understand the difference between ThT and Pyr assays for lipopeptide 3. We measured the CD signal for 3x10-3 wt% and 3x10-2 wt% lipopeptide 3 at 5 and 25 hrs after mixing the lipopeptide in water (Figure S4a). We also used DLS to calculate the hydrodynamic radius, RH, for samples in Figure S5a at 25 hrs after mixing the lipopeptide in water (Figure S4b). The CD spectra in Figure S4a has a minimum at ~216 nm ([Θ]min) and a maximum at ~197 nm ([Θ]max), characteristic of a β-sheet secondary structure,15 such that the population of β-sheets increases with concentration and time. RH is 603 and 1086 nm at 3x10-2 and 3x10-3 wt% lipopeptide 3 respectively, showing that the fibrils are more folded at 3x10-2 wt% than at 3x10-3 wt% lipoptide. We think that fibril folding might favour the encapsulation of Pyr previously observed by us at concentrations ≥ 0.03 wt% lipopeptide 3.4 Therefore cac measured by Pyr4 might correspond to the onset of encapsulation of Pyr by fibril folding, instead of the encapsulation of Pyr within the fibril core. In that case, cac= 0.03 wt% lipopeptide 3 measured by the Pyr assay can be dismissed against cac measured by the ThT assay (7.4x10-4 wt% lipopeptide 3).


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Results reported by Stupp and co-workers prove that self-assembly and βsheet formation do not need to be taking place at the same concentration.16

For

example, we have previously found that NH2-K(Boc)LVFF-CONH2 peptide undergoes hydrophobic collapse at a low concentration, followed by amyloid formation at a higher cac concentration.17

However, results discussed in the

paragraph above support the idea that lipopeptide 3 undergoes simultaneous selfassembly and β-sheet formation, at cac concentration measured in Figure 1c. CD was used to determine the secondary structure at native pH, above the cac. Figure 2a shows that lipopeptides 1 and 2, similarly to 3 discussed above, adopt a βsheet secondary structure.15 At 6.5x10-3 wt% lipopeptide, the β-sheet content is similar for 1 and 3 but simultaneously higher than for 2. The same spectra as in Figure 2a were observed upon increasing the temperature from 10 up to 75 oC. This shows that no change in peptide conformation occurs across 21 and 41.5 o

C, which are the critical melting temperatures cmTs for C14 and C16 respectively18

(cmT= -1oC for C1218 could not be reached). The inset in Figure 2a shows that [Θ]max/[Θ]min is relatively independent of T, only presenting a small deviation for 1 at 75 oC. In a separate experiment, CD was used to examine the behaviour of the secondary structure upon dilution. Figure 2b shows the CD data measured for a sample obtained by diluting a solution above the cac (0.1 wt% lipopeptide) down to a concentration below the cac (4.4x10-4 wt% lipopeptide). The results show that the βsheet secondary structure was preserved upon sample dilution below the cac for all the samples (Figure 2b), and the ratio [Θ]max/[Θ]min was also similar comparing initial and diluted solutions for a particular lipopeptide (inset Figure 2b). These results confirm that our previous observation of the stability of self-assembled structure to


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dilution below the cac for 3,4 can now be extended to 1 and 2, i.e. lipid chain length in the range C12-C16 does not influence the fact that aggregates can be stable below the measured cac. The FTIR spectra in the amide I region (Figure 3) support the CD assignment, with a strong peak at 1616-1620 cm-1.19 The shoulder peak around 1650-1660 cm-1 indicates a small contribution from unordered and/or α-helical secondary structure.19 The self-assembled nanostructure was imaged using cryo-TEM for samples containing 1 wt% lipopeptide. The morphologies obtained for 1, 2 and 3 are displayed in Figure 4. A coexistence of long twisted and short straight homogeneous fibres, ~7 nm thick, are observed for 1. Short crystalline-like striated tapes ~ 14 nm thick, with ~ 4.5 nm thick stripes, are observed for 2. In contrast, the morphology of 3 is dominated by more flexible twisted long tapes ~ 9 nm thick. Amphiphilicity drives the formation of tape-like nanostructures while fibril self-assembly is controlled by β-sheet formation.20 Lipopeptides 1 and 3 are possibly on the borderline between nanotape and fibril structure. In contrast, 2 forms thicker nanotapes with no evidence of fibril formation showing that amphiphilicity dominates the self-assembly of this sample. XRD was measured for stalks prepared by drying 1 wt % solutions. 2D XRD patterns show that the fibres and tapes in 1 and 3 are slightly oriented upon drying (Figure S5a-c). Reduced 1D intensity profiles correspond to a cross-β sheet pattern (Figure 5), where d-spacings at ~9 and ~10 Å indicate the lateral distance between β-sheets while peaks at 4.69 Å describe the strand separation in the β-sheets.21 Distances at 3.78 Å correspond to the Cα-Cα spacing in a β-sheet structure.21 We attempted to investigate the conformation of the lipopeptides via NMR spectroscopy. NMR spectra were measured for a 0.3 wt% solution of 3 in DMSO-d6


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(Scheme S1, Figure S6, Table S1), for which TEM experiments show that the peptide is self-assembled into fibres (results not shown). The partial assignment of the peptide side chains was successful for residues such as tyrosine (Y), glutamic acid (E), alanine (A), arginine (R) or leucine (L), and for the terminal group –CH3 of the C16 chain. It was not possible to assign a proton signal to the asparagine (N) at the peptide block, possibly because these residues are fully exposed to the solvent and exchange protons at the very high rate with the solution.22 The fast exchange is also expected for the hydroxyl groups of Y, threonine (T) and E, as denoted by the analysis of the linewidth recorded at 700 and 500 MHz which shows increase by at least 20% with decrease of the spectrometer frequency. The opposite trend or no change is observed for amine groups of the backbone suggesting slower exchange and less exposure to the solvent. The self-assembly in solution was also studied using SAXS. Figure 6 shows the SAXS data measured for 0.5, 2 and 4 wt% lipopeptide solutions. The SAXS data has been fitted using SASfit software23 (Figure 6); parameters extracted from the fitting are listed in Table S2. SAXS data for 0.5 wt% solutions in Figure 6a was fitted using a form factor for a lipid bilayer structure, comprising three Gaussian functions to represent the electron density variation across the two head groups and the dense lipid core.24 The model and its application to lipopeptide bilayer structures have been described in detail in our previous papers.25 SAXS data for concentrated samples (Figure 6b) considered the interactions between lipid bilayers using a modified Caillé structure factor26 in addition to the lipid bilayer structure. The SAXS curves for 2 and 3 in Figure 6 could be fitted to single bilayers. However, SAXS for 1 show a slightly inhomogeneous structure. The SAXS data for


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0.5 wt% lipopeptide 1 in Figure 6a was fitted as a co-existence of bilayers with long cylinders, consistent with the existence of tapes and fibres in the solution. The SAXS data for 2 wt% lipopeptide 1 in Figure 6b was fitted according to a co-existence of bilayers with two different thicknesses. Our theoretical models provide excellent fits to the SAXS data. The extended length of a lipopeptide containing a 13-residue peptide in a parallel β-sheet is l = length of the lipid chain+13 × 3.2 Å27 from the 13-residue peptide (lipid chain length= 18, 16, and 14 Å for C16, C14 and C12 respectively). Tables S2a-c show that l is on average higher than the total bilayer thickness lT. This result indicates that the bilayers comprise highly interdigitated molecules, with possibly also some folding of peptide residues. The optical texture of the samples could be studied at high concentrations by POM. Samples containing ~ 2 wt% lipopeptide were stained by Congo red, since they showed an apple green birefringent pattern (Figure 7), which is a diagnostic of amyloid formation.28 Flow alignment was observed in the POM when the samples were sheared between the glass slide and the coverslip (Figure 7 b and Figure 7f) for 1 and 3. A sample containing 2 did not orient under shear ( Figure 7d). Again, these results are in agreement with Figures S5a-c that show partial alignment for the 2D XRD for 1 and 3 but not for 2. The stability of the peptide block within the lipopeptide fibril was tested by carrying out an enzymatic assay of asparaginase (Asp). The enzyme is expected to act on the –H2N groups of the asparagine (E) changing them into -OH groups and turning that residue into aspartic acid (D). A solution containing 0.1 wt% sample 3 and 0.3 wt% Asp was incubated at pH 8.6 adjusted with 3.3 wt% NaOH. TEM images show that the lipopeptide is self-assembled into fibres after the incubation (results not


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shown). However, retention times and ES-MS spectra (Figure S7) show that the enzyme does not reduce the E residue during the Asp assay. It is likely that within the self-assembled fibre, the E-side is not exposed to the water or is in close contact to the side chains of other neighbouring residues in the lipopeptide block in a way that it is not accessible to the Asp.

Tuning the pH for Peptide Hydrogel formation. As mentioned above, peptide hydrogels were made by changing the pH of the solution. In this process, the pH of a 1 wt% lipopeptide solution in water (Figure S3) was increased to pH 10 through NaOH titration in the solution. Cryo-TEM images of the solutions at pH 10 are displayed in Figure 8. For all three samples, increasing the pH to 10 resulted in longer and straighter fibres than those displayed in Figure 4 for the lipopeptides in water. In addition, there is a co-existence of micelles with fibres, not observed for the peptides mixed in pure water. The thickness of the fibres is about 10, 6.5 and 8 nm for 1, 2 and 3 respectively. The micelle diameter is approximately 15, 13 and 15 nm for 1, 2 and 3. Hydrogels were obtained by injecting 1 M HCl to the solutions corresponding to those used to produce the cryo-TEM images in Figure 8. Figure 9a-c displays the self-standing hydrogels obtained for each sample. Hydrogels are made by increasing the pH of a 1 wt% lipopeptide solution in water (native pH) to pH 10, and then decreasing it to pH 1.48, 1.33 and 1.21 for 1, 2 and 3 respectively. Hydrogel formation is triggered by changes in electrostatic interactions between the peptide block of neighbouring lipopeptides, induced by changes in the pH. Nanotapes in 1 wt% solutions in water are at pH ~ 4-7 (Figure S3). The arginine (R), a very polar basic and hydrophilic residue, is positively charged at pH ~ 4-7. In contrast, at pH 10, the R-side chain is uncharged, while all the other


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amino acids in the peptide block are negatively charged. A negative charge for the nanotape surfaces is confirmed by zeta potential values (-42.9±3.3), (-48.9±1.7) and (50.5±1.3) mV measured at pH 10 for 1, 2 and 3 respectively. The R-side chain becomes positively charged upon decreasing the pH from 10 to ~1, while all the other residues in the peptide block become neutralized. The intermediate step at pH 10 is essential for hydrogel formation, as illustrated by the following experiment. The discussion above can be illustrated by CD results Figure S8, measured for a 6.5x10-3 wt% lipopeptide 1 sample, initially dissolved in water (pH 6.15), which pH was first raised to 10 (by NaOH sol titration) and then decreased to 1.48 (HCl sol titration). The CD for the lipopeptide in water above the cac (pH 6.14) corresponds to a β-sheet order. When the pH is increased to 10 the peptide block exhibits a disordered conformation. A further reduction of the pH to 1.48 returns the peptide block to the β-sheet conformation. Results in Figure S8 can be used to understand the process for hydrogel formation observed at 1 wt% lipopeptide. At pH 6.14 the lipopeptide fibres are stabilized by the competition between β-sheets bonds and electrostatic interactions, balanced by the amphiphilic nature of the molecule. At pH 10, repulsive interactions between Y, E, A, L, V, N, V and T negatively charged residues overcome the tendency of the peptide to self-assemble into β-sheets and the peptide structure becomes disordered. Negatively charged residues exposed to the water at pH 10, are immediately neutralized upon reduction of the pH to 1.48. Peptide fibres formed at pH 1.48 benefit from the initial cancellation of repulsive interactions from Y, E, A, L, V, N, V and T residues and are stabilized by a weaker repulsive interaction between R residues competing with β-sheet bonding. The initial cancellation of the repulsion between Y, E, A, L, V, N, V and T residues when the pH is reduced from 10 to 1.48


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allows the peptide fibres to make a network and consequently form an hydrogel at pH 1.48. This is consistent with previous reports on conditions that favour peptide gelation when the net charge on the peptide is in a favourable range to balance interpeptide interactions.29 The high relevance of the intermediate step at pH 10 for hydrogel formation is proved by the fact that several tests shown that hydrogels are not formed when the pH is reduced directly from 6.14 to 1.48. Dried hydrogels, their nanostructure fixed with a binary solution of paraformaldehyde and glutaraldehyde, were studied by high vacuum SEM. The resulting images are displayed in Figure 10. According to SEM, the hydrogels comprised an entangled network of aggregates of nanotapes (57.7±9.3) nm, (76.2±10.1) nm and (58.6±6.1) nm thick for 1, 2 and 3 respectively. The hydrogels shown in Figure 9 instantly adsorb methylene blue when dipped in a 2.9x10-3 wt% methylene blue solution, acquiring a bright blue colour, as it is exemplified in Figure 9d for an hydrogel made from 2. The uptake of the cationic dye methylene blue by the lipopeptide hydrogel is not favoured by the positive charge of the lipopeptide in the hydrogel. Instead, uptake of methylene blue is probably due to a diffusion phenomenon favoured by the network structure of the peptide hydrogel. Rheology experiments were performed to evaluate the viscoelastic properties of the hydrogels. Stress sweeps show approximately linear viscoelastic behaviour up to a stress σ = 90 Pa (Figure S9). Frequency sweeps at a fixed shear stress within the linear regime (σ = 80 Pa), reveal frequency-dependent moduli G ′′ > G ′ consistent with a hard gel-like response (Figure 11). These self-standing hard gels, easily release water by being compressed by the rheometer tool, resulting in the relatively high shear modulus reported in Figure 11.


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Isotropic 2D XRD spectra measured on stalks of the hydrogels are displayed in Figures S5d-f. The 1D radially averaged profile and d-spacings of data in Figures S5d-f is shown in Figure S10. 1D XRD data for hydrogel from 3 shows a higher number of peaks, probably associated to partial crystallization of the peptide during stalk preparation. However, there are no major structural differences between the stalks measured for the solutions or the hydrogels for a particular peptide, suggesting a similarity in the internal structure of the peptide fibres. This hypothesis was confirmed by SAXS experiments on the hydrogels. Figure 12 shows the SAXS data measured for hydrogels, fitted according to the form factor of bilayers (represented by a sum of Gaussian functions), which is basis for the tape-like self-assembled motif

24

The parameters extracted from these fittings (Tables S2) show that the lipid chains are overlapped inside the core of the bilayers, since the estimated bilayer length, lT, is smaller than two times the length of the extended molecule, l. In fact, SAXS confirms that the internal structure of the bilayers in the hydrogel is very similar to that observed for the lipopeptide nanotapes in water (Figures 4, 6).

Conclusions We studied the self-assembly of three lipopeptides, with the same peptide block (YEALRVANEVTLN) but different lipid chain lengths (C12, C14 and C16 for 1, 2 and 3 respectively). The lipopeptides self-assemble into nanotapes above a cac which decreases with increasing lipid chain length. CD spectroscopy indicates that the β-sheet secondary structure is stable upon dilution to concentrations below the cac. The fact that this is observed for all three lipid chain lengths suggests that this phenomenon is not a kinetic process resulting from partial vitrification or crystallization of the lipid chains, which was a possible scenario for the C16


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homologue but cannot be for the C12 since its melting points is too low. Of note in relationship to these observations is the recent report by the Stupp group of kinetically trapped monodisperse long fibres with β-sheet structure. This behaviour was dependent on ionic strength of the solution of the charged lipopeptide molecules studied.30 The thermodynamically stable state (accessed via annealing) at low ionic strength was found to be short monodisperse fibrils with a random coil secondary structure. A kinetically trapped state of long β-sheet fibres was accessible via dilution from high ionic strength to below a critical ionic strength. Re-annealing drove the system back

to the thermodynamically stable state. Here we show via CD

measurements that the β-sheet structures for our three lipopeptides are thermally stable in a range 10 – 70 oC, which crosses the expected lipid chain melting temperature in the case of the -C16 and –C14 homologues. This suggests that lipid chain melting is either suppressed in the lipopeptides, or if it occurs it does not influence the peptide conformation probed by CD. It thus seems most likely that the kinetic trapping process may be related to favourable electrostatic interactions dependent on ionic strength. Further examination of this, along with kinetic aspects, will be an interesting topic for future research. The nanotapes have a negatively charge surface in water and an internal bilayer structure composed of overlapped lipopeptide molecules. Mass spectroscopy indicates that the E-residues are buried inside the nanotape core in a way that is not accessible for enzyme degradation. Nanotapes of 1 and 3 are easily oriented under shear in solution. Lipopeptide hydrogels can be prepared by first changing the pH of the solution to 10 and then decreasing it to ~1. Hydrogel formation is driven by changes in the electrostatic charge of the peptide residues at different pHs; the intermediate step at


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pH 10 being essential for hydrogel formation.

Lipopeptide hydrogels are self-

standing hard gels that release water upon compression. The hydrogels are built from a network of bundles of lipopeptide

nanotapes. The internal structure of the

nanotapes in the hydrogel is very similar to the internal structure of the nanotapes in water. Hydrogels presented in this work are instantly formed through pH switching, which represents an advantage in applications such as encapsulation and release formulations.

Acknowledgements This work was supported by EPSRC grant EP/L020599/01 to IWH. We thank the ESRF for the award of bioSAXS beamtime on BM29 (ref. MX1869) and Susana Gonçalves Pires and Martha Brennich for assistance with the measurements. Diamond Beamtime at I22 (ref. SM15750-1) and help from Olga Shebanova. We acknowledge Nick Michael for assistance with MS and Nick Spencer for assistance with XRD experiments, and the access to the Chemical Analysis Facility Laboratory (University of Reading).

Additional Information Supplementary information. NMR data and peak assignments, SAXS fitting parameters, ES-MS spectra, concentration dependence pH data, rheology and XRD figures can be found in the Supporting Information. This material is available free of charge via the Internet at http://pubs.acs.org.


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References 1. Walter, M. N. M.; Dehsorkhi, A.; Hamley, I. W.; Connon, C. J. Biomat. Sci. 2016, 4, 346-354. 2. (a) Lowik, D.; Leunissen, E. H. P.; van den Heuvel, M.; Hansen, M. B.; van Hest, J. C. M. Chem. Soc. Rev. 2010, 39 (9), 3394-3412; (b) Löwik, D. W. P. M.; van Hest, J. C. M. Chem. Soc. Rev. 2004, 33, 234-245 ; (c) Cavalli, S.; Kros, A. Adv. Mat. 2008, 20 (3), 627-631; (d) Cui, H. G.; Webber, M. J.; Stupp, S. I. Biopolymers 2010, 94 (1), 1-18; (e) Zhao, X. B.; Pan, F.; Xu, H.; Yaseen, M.; Shan, H. H.; Hauser, C. A. E.; Zhang, S. G.; Lu, J. R.. Chem. Soc. Rev. 2010, 39, 3480-3498; (f) Hamley , I. W. Soft Matter 2011, 7, 4122-4138; (g) Dehsorkhi, A.; Castelletto, V.; Hamley , I. W. J. Pept. Sci. 2014, 20, 453-467; (h) Hamley, I. W. Chem. Comm. 2015, 51, 8574-8583. 3. (a) Matson, J. B.; Stupp, S. I. Chem. Comm. 2012, 48, 26-33; (b) Arslan, E.; Garip, I. C.; Gulseren, G.; Tekinay, A. B.; Guler, M. O. Adv. Healthc. Mat. 2014, 3, 1357-1376. 4. Hamley, I. W.; Dehsorkhi, A.; Castelletto, V.; Walter, M. N. M.; Connon, C. J.; Reza, M.; Ruokolainen, J. Langmuir 2015, 31, 4490-4495. 5. Meijer, J. T.; Roeters, M.; Viola, V.; Lowik, D.; Vriend, G.; van Hest, J. C. M. Langmuir 2007, 23, 2058-2063. 6. Lowik, D.; Garcia-Hartjes, J.; Meijer, J. T.; van Hest, J. C. M. Langmuir 2005, 21, 524-526. 7. Xu, X.-D.; Jin, Y.; Liu, Y.; Zhang, X.-Z.; Zhuo, R.-X. Colloids Surf. B 2010, 81, 329-335. 8. Damen, M.; Cristobal-Lecina, E.; Sanmarti, G. C.; van Dongen, S. F. M.; Rodriguez, C. L. G.; Dolbnya, I. P.; Nolte, R. J. M.; Feiters, M. C. Soft Matter 2014, 10, 5702-5714. 9. Hartgerink, J. D.; Beniash, E.; Stupp, S. I. PNAS 2002, 99, 5133-5138. 10. (a) Paramonov, S. E.; Jun, H. W.; Hartgerink, J. D. JACS 2006, 128, 72917298; (b) Pashuck, E. T.; Cui, H. G.; Stupp, S. I. JACS 2010, 132, 6041-6046; (c) Stendahl, J. C.; Rao, M. S.; Guler, M. O.; Stupp, S. I. Adv. Funct. Mat. 2006, 16, 499508; (d) Kim, J. K.; Anderson, J.; Jun, H. W.; Repka, M. A.; Jo, S. Mol. Pharm. 2009, 6, 978-985; (e) Rexeisen, E. L.; Fan, W.; Pangburn, T. O.; Taribagil, R. R.; Bates, F. S.; Lodge, T. P.; Tsapatsis, M.; Kokkoli, E. Langmuir 2010, 26, 1953-1959; (f) Anderson, J. M.; Andukuri, A.; Lim, D. J.; Jun, H. W. Acs Nano 2009, 3 (11), 34473454; (g) Hosseinkhani, H.; Hosseinkhani, M.; Khademhosseini, A.; Kobayashi, H.; Tabata, Y. Biomaterials 2006, 27, 5836-5844; (h) Zhou, J.; Li, J.; Du, X.; Xu, B. Biomaterials 2017, 129, 1-27; (i) De Leon Rodriguez, L. M.; Hemar, Y.; Cornish, J.; Brimble, M. A. Chem. Soc. Rev. 2016, 45, 4797-824. 11. (a) LeVine, H. Prot. Sci. 1993, 2, 404-410; (b) LeVine, H. In Methods in Enzymology, Wetzel, R., Ed. Academic Press: San Diego, 1999; Vol. 309, pp 274284. 12. Innovagen's Peptide Property Calculator. Copyright © 2012 Innovagen AB. 13. (a) Rousseau, F.; Schymkowitz, J.; Serrano, L., Protein aggregation and amyloidosis: confusion of the kinds? Curr. Op. Struct. Biol. 2006, 16, 118-126; (b) Fernandez-Escamilla, A. M.; Rousseau, F.; Schymkowitz, J.; Serrano, L.. Nat. Biotech. 2004, 22., 1302-1306; (c) Linding, R.; Schymkowitz, J.; Rousseau, F.; Diella, F.; Serrano, L. J. Mol. Biol. 2004, 342, 345-353. 14. (a) Lindgren, M.; Sorgjerd, K.; Hammarstrom, P. Biophys. J. 2005, 88, 42004212; (b) Khurana, R.; Coleman, C.; Ionescu-Zanetti, C.; Carter, S. A.; Krishna, V.;


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Grover, R. K.; Roy, R.; Singh, S., J. Struct. Biol. 2005, 151, 229-238; (c) Krebs, M. R. H.; Bromley, E. H. C.; Donald, A. M. J. Struct. Biol. 2005, 149, 30-37. 15. (a) Hamley , I. W. Angew. Chem. Int. Ed. 2007, 46, 8128-8147; (b) Nordén, B.; Rodger, A.; Dafforn, T. R., Linear Dichroism and Circular Dichroism: A Textbook on Polarized-light Spectroscopy. Cambridge, 2010; (c) Woody, R. W., Circular Dichroism of Peptides and Proteins. In Circular Dichroism: Principles and Applications, Nakanishi, K.; Berova, N.; Woody, R. W., Eds. VCH: New York, 1994; pp 473-496. 16. Korevaar, P. A.; Newcomb, C. J.; Meijer, E. W.; Stupp, S. I. JACS 2014, 136, 8540-8543. 17. Castelletto, V.; Ryumin, P.; Cramer, R.; Hamley, I. W.; Taylor, M.; Allsop, D.; Reza, M.; Ruokolainen, J.; Arnold, T.; Hermida-Merino, D.; Garcia, C. I.; Leal, M. C.; Castano, E. Sci, Reports 2017, 7, article number 43637. 18. Cevc, G. Biochemistry 1991, 30, 7186-7193. 19. (a) Jackson, M.; Mantsch, H. H. Crit. Rev. Biochem. Mol. Biol. 1995, 30, 95120; (b) Stuart, B., Biological Applications of Infrared Spectroscopy. Wiley: Chichester, 1997. 20. Castelletto, V.; Hamley, I. W.; Perez, J.; Abezgauz, L.; Danino, D. Chem. Commun. 2010, 46, 9185-9187. 21. Serpell, L. C., BBA - Mol. Basis Dis. 2000, 1502, 16-30. 22. Yamazaki, T.; Pascal, S. M.; Singer, A. U.; Forman-Kay, J. D.; Kay, L. E.,. JACS 1995, 117, 3556-3564. 23. Breßler, I.; Kohlbrecher, J.; Thünemann, A. F. J. Appl. Cryst. 2015, 48, 15871598. 24. Pabst, G.; Rappolt, M.; Amenitsch, H.; Laggner, P.. Phy. Rev. E 2000, 62, 4000-4008. 25. (a) Castelletto, V.; Cheng, G.; Stain, C.; Connon, C.; Hamley, I.. Langmuir 2012, 28 11599-11608 ; (b) Castelletto, V.; Gouveia, R.; Connon, C. J.; Hamley, I. W. Far. Disc. 2013, 166, 381-397. 26. (a) Caillé, A. CRendus Acad. S. B: S. Phys. 1972, 274, 891-893; (b) Caillé, M. C. R. Acad. Sci. Paris 1972, 274, 891-893. 27. Creighton, T. E., Protein Folding W.H. Freeman: New York, 1992. 28. Hamley, I. W.; Castelletto, V.; Moulton, C.; Myatt, D.; Siligardi, G.; Oliveira, C. L. P.; Pedersen, J. S.; Abutbul, I.; Danino, D. Macromol. Biosc. 2010, 10, 40-48. 29. Rajagopal, K.; Lamm, M. S.; Haines-Butterick, L. A.; Pochan, D. J.; Schneider, J. P. Biomacromolecules 2009, 10, 2619-2625. 30. Tantakitti, F.; Boekhoven, J.; Wang, X.; Kazantsev, R. V.; Yu, T.; Li, J.; ZHuang, E.; Zandi, R.; Ortony, J. H.; Newcomb, C. J.; Palmer, L. C.; Shekhawat, G. S.; Olvera de la Cruz, M.; Schatz, C.; Stupp, S. I. Nat. Mat. 2016, 15, 469-477.


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Figures and Schemes

Scheme 1. Molecular structure of lipopeptides (a) 1, (b) 2 and (c) 3.


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90

(a)

1

60

cac ~1.8x10-3 wt%

30 0

(b)

2

20

I / Io

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

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-3

cac ~1.5x10 wt%

10 0

(c)

3

100 cac ~7.4x10-4 wt%

50 0

-5

-4

log(C/ wt%)

-3

-2

Figure 1. Critical aggregation concentration (cac) assay for lipopeptides (a) 1, (b) 2 and (c) 3 using ThT fluorescence assay. The fluorescence emission of the peptide/ThT solution, I, has been normalized by the fluorescence emission of the solution without peptide, Io.


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(a)

[Θ] max / [Θ] min

2

-1

3

3

-2

1

-3

0

25

50

75

T / oC

2

20 oC 6.5x10-3 wt% lipopeptide

1 3

-3 [Θ] max / [Θ] min

10-5x[Θ] / degrees cm2dmol

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3

original diluted

(b)

-1

1

2

0

2 lipopeptide #

3

1 3

-3 190

200

210

4.4x10-4 wt% lipopeptide

220 230 λ / nm

240

250

260

Figure 2. CD data for solutions of the three lipopeptides at the concentration indicated. The inset in (a) shows the temperature dependence of [Θ]max/[Θ]min and the inset in (b) shows this quantity for the three samples at 20 oC comparing original or diluted samples. [Θ]max and [Θ]min are measured at 197 and 216 nm respectively.


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0.8 0.4

(a) 1, (b) 0.5, (c) 0.1 wt% lipopeptide 1

(a)

2

(b)

1 b c

0.0 0.4

2

1: 1660 cm-1 2: 1616 cm-1

a

Intensity / a. u.

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(a) 1, (b) 0.5, (c) 0.1 wt% lipopeptide 2 1: 1660 cm-1 2: 1620 cm-1

0.2

1 a b

0.0

c

(a) 1, (b) 0.5, (c) 0.1 wt% lipopeptide 3

(c)

2

0.4 1: 1660 cm-1 2: 1620 cm-1

0.2

1 a

0.0 1800

c

b

1650 wavenumber / cm-1

1500

Figure 3. FTIR spectra measured for lipopeptides (a) 1, (b) 2 and (c) 3, at 0.1, 0.5 and 1 wt% lipopeptide.


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Figure 4. Cryo-TEM images collected for 0.5 wt% solutions of lipopeptides (a) 1, (b) 2 and (c) 3. Arrows indicate twists in the fibres.


30

3.81 Å

1

2.38 Å

4.69 Å

3.79 Å

4.12Å 3.78 Å

4.69 Å

10.22 Å 9.6 Å 9.37 Å

Intensity / a. u.

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4.69 Å

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2 1 wt % lipopetide 3

1

2 q / Å-1

3

4

Figure 5. XRD data for stalks made from 1 wt% lipopeptide (a) 1, (b) 2 and (c) 3.


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(a)

1 2 3

Intensity / a. u.

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0.5 wt% lipopeptide

(b)

2 wt% 1 4 wt% 2

2 wt% 3

0.01

q / Å-1

0.1

Figure 6. SAXS data measured at different concentrations of lipopeptide. Only samples in (b) show birefringence under optical polarized microscopy. The full line is a fitting to the data, as described in the text. Parameters extracted from the fitting are listed in Tables S2a-c.


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Biomacromolecules

Figure 7. Polarized optical microscope images obtained for samples stained with Congo red. 2 wt% lipopeptide 1 (a) before and (b) after shear; 2 wt% lipopeptide 2 (c) before and (d) after shear; and 2.4 wt% lipopeptide 3 (e) before and (f) after shear.


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Biomacromolecules

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Figure 8. Cryo-TEM images for 1 wt% lipopeptide (a) 1, (b) 2 and (c) 3 at pH 10.


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Biomacromolecules

Figure 9. Hydrogels made from 1 wt% lipopeptide (a) 1, (b) 2 and (c) 3. (d) corresponds to an hydrogel of 2 dyed with 2.9x10-3 wt% methylene blue.


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Biomacromolecules

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Figure 10. SEM images for hydrogels made from 1 wt% lipopeptide (a) 1, (b) 2 or (c) 3.


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Hydrogels, G' lipopeptide: G" lipopeptide:

G', G'' / Pa

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Biomacromolecules

1, 1,

2 and 2 and

3 3

1E+04

1E+03

0.1

1 10 ang. freq. / rad s-1

100

Figure 11. Dependence of G’ and G’’ on the oscillatory frequency measured for hydrogels made from 1 wt% lipopeptide. The oscillatory stress fixed to 80 Pa (1 and 2) or 20 Pa (3), was in the linear regime according to data in Figure S6.


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Biomacromolecules

Hydrogel, 1

Intensity / a. u.

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35 36 37 38 39 40 41 42 43 44 45 46 47 48 49 50 51 52 53 54 55 56 57 58 59 60

Hydrogel, 2

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(a)

(b)

(c) Hydrogel, 3

0.01

0.1 -1

q/Å

Figure 12. (circles) SAXS data for hydrogels made from 1 wt% of lipopeptides (a) 1, (b) 2 and (c) 3. The full line is a fitting to the data, as described in the text. Parameters extracted from the fitting are listed in Tables S2a-c.


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Biomacromolecules

Table of Contents Figure

Lipopeptide hydrogels are formed by changing the pH of a 1 wt% lipopeptide solution


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Supramolecular Hydrogel Formation in a Series of ... - ACS Publications

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