A self-assembly pathway to aligned monodomain gels

A self-assembly pathway to aligned monodomain gels

(Parte 1 de 2)

A self-assembly pathway to aligned monodomain gels

Shuming Zhang1†, Megan A. Greenfield2†, Alvaro Mata3‡, Liam C. Palmer4, Ronit Bitton3, Jason R. Mantei1, Conrado Aparicio3, Monica Olvera de la Cruz1,2,3,4 and Samuel I. Stupp1,3,4,5*

Aggregates of charged amphiphilic molecules have been found to access a structure at elevated temperature that templates alignment of supramolecular fibrils over macroscopic scales. The thermal pathway leads to a lamellar plaque structure with fibrous texture that breaks on cooling into large arrays of aligned nanoscale fibres and forms a strongly birefringent liquid. By manually dragging this liquid crystal from a pipette onto salty media, it is possible to extend this alignment over centimetres in noodle-shaped viscoelastic strings. Using this approach, the solution of supramolecular filaments can be mixed with cells at physiological temperatures to form monodomain gels of aligned cells and filaments. The nature of the self-assembly process and its biocompatibility would allow formation of cellular wires in situ that have any length and customized peptide compositions for use in biological applications.

Inspired largely by biological systems, molecular self-assembly continues to be a theme of great interest in science. The targets differ broadly, from accessing ordered materials and self-assemblingdevices1–3 tounderstandinghowmisfoldedproteins self-assemble into stable fibres linked to human disease4,5. Longrange alignment of extracellular fibrils and cells in the heart6, brain and spinal cord7 must involve highly complex self-assembly mechanisms that remain largely unknown. Access to similar threedimensional (3D) artificial systems of aligned fibrils and cells is therefore of scientific and biomedical interest8–10. Spontaneous long-range alignment of molecules is known to occur in liquid crystals11 but its fixation in the solid state normally requires chemical reactions12,13 that are not likely to be compatible with living cells. Electrospinning of polymers faces similar challenges because it requires the use of high mechanical and electrical energies14 that are not highly compatible with encapsulation of living cells. We report the discovery of a thermal pathway that leads highly designable peptide-based small molecules in water to form two-dimensional (2D) plaques with filamentous texture that spontaneously template long-range alignment of bundled nanofibres on cooling. This liquid crystal of supramolecular filaments can be mixed with cells at physiological temperatures and drawn by hand from a pipette into salt solutions to form monodomain gels of aligned filaments. Cells remain viable during the process and the monodomain gels can be drawn to arbitrary lengths and geometrical contours. We hypothesize that divalent ions and slow relaxation times of the long nanofibre bundles generated by this self-assembly pathway enable formation of the macroscopic monodomains.

We prepared 0.5–1.0wt% aqueous solutions of peptide amphiphiles known to self-assemble into high-aspect-ratio nanofibres15,16. One peptide amphiphile molecule investigated contains the peptide sequence V3A3E3(CO2H) and a C16 alkyl tail

1Department of Materials Science and Engineering, Northwestern University, Evanston, Illinois 60208, USA, 2Department of Chemical and Biological Engineering, Northwestern University, Evanston, Illinois 60208, USA, 3Institute for BioNanotechnology in Medicine, Northwestern University, Chicago, Illinois 60611, USA, 4Department of Chemistry, Northwestern University, Evanston, Illinois 60208, USA, 5Department of Medicine, Northwestern University, Chicago, Illinois 60611, USA. †These authors contributed equally to this work. ‡Present address: Nanotechnology Platform, Parc Cientific, 08028 Barcelona, Spain. *e-mail:s-stupp@northwestern.edu.

at the peptide’s N-terminus, and its self-assembly into nanofibres is triggered by ions that screen the charged amino-acid residues, resulting in the formation of gels. The diameter of these nanofibres, which contain β-sheets near their hydrophobic core, is roughly equivalent to the length of two peptide amphiphile molecules and lengths in excess of micrometres. We heated the aqueous solutions unscreened by added ions to 80◦C and kept them at this temperature for 30min before cooling to 25◦C. After this heat treatment, the solution viscosity increased threefold from 5 to 15cP. When calcium chloride was added to the heated and cooled peptide amphiphile solution, we observed the formation of a gel that was at least fourfold stiffer than one formed from an unheated solution (see Supplementary Information). Using polarized optical microscopy, we also found that gels or films formed from heated solutions contained large birefringent domains (tenths of millimetres; Fig. 1), whereas those formed from unheated solutionsappearedcompletelyisotropicwithnobirefringence.

We observed that noodle-like strings of arbitrary length could be formed by manually drawing the aqueous peptide amphiphile solution into a salty medium from a pipette (Fig. 1a,b). When the solution was dragged on a surface covered by a thin layer of this medium (Fig. 1c), uniform birefringence was observed along the length of the string (Fig. 1h,i). This observation suggested that macroscopic alignment extending over centimetres was achieved. Using the same methods, the unheated solutions did not form mechanically stable string gels or show any birefringence. Scanning electron microscopy (SEM) indicated that strings formed from heated peptide amphiphile solutions contained extraordinarily long arrays of aligned nanofibre bundles (Fig. 2a,b). In great contrast, unheated peptide amphiphile solutions formed matrices of randomly entangled nanofibres (Fig. 2d,e). To verify this orientational order, we carried out small-angle X-ray scattering (SAXS) experiments and found that only strings generated from the

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Figure 1 | Strings and gels with long-range internal alignment. a,b, A peptide amphiphile solution coloured with trypan blue injected into phosphate-buffered saline after heat treatment. c, The same solution dragged through a thin layer of aqueous CaCl2 to form a noodle-like string. d, A knot made with peptide amphiphile string. e, Birefringence of a bubble gel observed between cross polars suggesting the presence of macroscopically aligned domains. f, Similar domains in a gel film. g, Peptide amphiphile noodle spirals prepared on a spin coater. h, Birefringence of a single string suggesting alignment along the string axis. i, Light extinction between cross polars at the crosspoint of two noodles demonstrating uniform alignment in each.

heated solutions revealed alignment (Fig. 2c,f). In contrast, simply dragging a peptide amphiphile solution that had not been heated does not lead to significant alignment. Similar strings can be made with other peptide amphiphile molecules, including those with bioactive epitopes, although their structural integrity depended on theamino-acidsequence(SupplementaryInformation).

To gain a mechanistic understanding of the observed transformations, we examined the effects of heating on the peptide amphiphile solution structure by quick-freeze/deep-etch (QFDE) transmission electron microscopy17 (TEM). QFDE is a sample preparation technique that allows high-resolution imaging of hydrated structures while minimizing disruption of the sample resulting from fixation or processing. The freshly dissolved peptide amphiphilesolutioncontainedavarietyofnanoscale,elongatedobjects lessthanamicrometreinlength(SupplementaryInformation).Micrographs of the peptide amphiphile solution equilibrated at 80◦C for 30min showed that the small aggregates largely disappeared; instead we observed thin ‘plaque-like’ structures up to micrometres in both length and width (Fig. 3a). Some portions of these 2D plaques had a periodic surface texture with a characteristic spacing of about 7.5nm, which corresponds to the expected diameter of a single canonical nanofibre formed by the peptide amphiphile molecules used15,18 (Fig. 3b). After cooling to room temperature, solutions were clearly composed of aligned filaments (Fig. 3d). The filaments did not have the diameter of canonical nanofibres (7–8nm) but were instead tens of nanometres in diameter, and are therefore bundles of many cylindrical nanofibres. To further visualize the 3D structure of the plaque, we fixed the structure by adding calcium chloride at 80◦C and imaged the resulting structure by SEM. Although most of the sample cooled to ambient temperature was composed of large arrays of aligned nanofibres, some plaques were captured as well (Fig. 3e). These plaques measured about 40nm in thickness and had lengths and widths comparable to those observed by QFDE-TEM. They often contained long parallel striations and, in some cases, appeared to be cracking into fibre bundles (Fig. 3f). Theseplaquestructureswerenotobservedwithoutheating.

We explored the effect of heating on structure in peptide amphiphile solutions using SAXS, circular dichroism, differential scanning calorimetry and Fourier-transform infrared spectroscopy. The X-ray scattering curve of peptide amphiphile solutions at 25◦C (Fig. 3c) shows a q−4 dependence within the low q range, indicative ofthepresenceoflargeaggregates.Onheatingto80◦C,the−4slope

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Figure 2 | SEM evidence of massive alignment versus isotropy of nanofibre bundles. a,c, Aligned nanofibre bundles in macroscopic strings formed by dragging thermally treated amphiphile solutions onto a CaCl2 solution. b,d, Isotropic network of nanofibre bundles formed by adding CaCl2 to unheated amphiphile solutions. e,f, SAXS of hydrogel strings prepared using peptide amphiphile solutions with and without heat treatment.

(log I versus log q) was retained, indicating that large aggregates are still present at high temperature19. This result is consistent with our observation of large plaque-like structures at 80◦C by QFDE-TEM. The scattering curves at 25◦C and 80◦C essentially superimpose at high q values, indicating that the local structure of supramolecular aggregates remains unchanged during heating. We alsoobserveabroadscatteringpeakintheintermediateqrangenear 0.08A−1. This q range corresponds to a spacing of approximately 8nm, which is in the range of the diameter of the fibrous structures that exist in these solutions before heating. On heating, this scattering maximum decreases in intensity, which is consistent with our observation that individual fibres are not observed at high temperature. The observed decrease in scattering at intermediate q values as solutions are heated is irreversible on cooling of the samples. In fact, the fibrous structures observed after cooling by microscopy have diameters that are several times larger than the original ones and therefore their scattering contribution would be expected in the low rather than intermediate q range. Variabletemperature circular dichroism revealed the presence of β-sheet structure throughout the temperature range studied. Effectively no change was observed in the circular dichroism spectrum during heating and cooling cycles (Supplementary Information).

Therefore, the heating and cooling of peptide amphiphile solutions does not significantly perturb the peptide secondary structure within the aggregates, which is also supported by X-ray diffraction studies (Supplementary Information). Fourier-transform infrared spectroscopy studies of the solutions before and after heating show a major peak for parallel β-sheets (1,625cm−1) and a smaller peak at about 1,660cm−1 that can be attributed to antiparallel β-sheets (Supplementary Information). Differential scanning calorimetry of a solution of the peptide amphiphile molecules did not show any peaks during heating and cooling cycles, indicating the absence of obvious phase transitions. We conclude from these experimental results that the reorganization of molecules from the 2D plaques to filaments on cooling is a subtle one, which we believe reflects changesinthehydrationoftheaggregates,asexplainedbelow.

Canonical peptide amphiphile nanofibres are known to be internally hydrated20 and their charged surfaces should also contain electrostricted water molecules around charged aminoacid residues. It is reasonable to assume that as the system is heated, bound water molecules associated with the supramolecular aggregates become bulk water molecules driven by entropy. This dehydration process could allow the fibrous aggregates to interact more closely, facilitating their fusion as depicted in Fig. 3g. We

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Figure 3 | Morphological changes resulting from thermal treatment. a, TEM obtained after a QFDE preparation of peptide amphiphile solution at 80◦C revealing a micrometre-sized, sheet-like plaque structure. b, Higher-resolution QFDE-TEM of the sheet-like structures revealing a surface pattern with a periodicity of about 7.5nm. c, SAXS of peptide amphiphile solutions treated at different thermal conditions. d, QFDE-TEM of aligned nanofibre bundles templated by the sheet-like plaque after the peptide amphiphile solution was cooled to room temperature. e, SEM of plaques that were captured by adding

CaCl2 at 80◦C. f, SEM of a plaque breaking into nanofibre bundles. g,h, Schematic representation of a plaque at high temperature (g) and its rupture into fused bundles on cooling (h). i,j, Schematic representation of the cross-section of a plaque formed by fused fibres (i) and of a fibre bundle (j).

believe the observed flat and textured plaques are the result of these fused, dehydrated fibres. During cooling, water molecules should simply rehydrate the supramolecular structures, but this process actually generates filaments with diameters that are several times greater than those of canonical nanofibres that exist before the thermal treatment (see the schematic in Fig. 3h). On the basis of microscopy, the plaque appears to break into nanofibre bundles with diameters of approximately 40nm (Fig. 2c) rather

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ARTICLES NATUREMATERIALSDOI:10.1038/NMAT2778 than individual canonical fibres with diameters of about 8nm. The diameter of the bundles corresponds to the plaque’s thickness observed by SEM (Fig. 3e).

The possibility of a transition from the plaque to a fused nanofibre bundle can be understood by computing the contributions to the difference in free energy per thermal energy kBT per amphiphile for a cylindrical fibre Ff and a lamella Fl, where Fef and Fel are the electrostatic free energies of a fibre and a lamella, respectively, and Fcf and Fcl are the cohesive free energies of molecules in these two different morphologies. The fraction of ions condensed on the surface of peptide amphiphile fibres and plaques is estimated from the modified Poisson–Boltzmann equation21, which shows that the charges are neutralized by counterions for both the plaque and the fibre (see Supplementary Information). Therefore, the transition should be dominated by the differenceincohesiveenergiesoflamellaeandfibres,1Fc =Fcf−Fcl, which is given by,

where 1HPA is the enthalpy difference per peptide amphiphile molecule between lamellar and fibre aggregates, and 1SPA and

1Swater are the entropy differences of the peptide amphiphile and water molecules, respectively. As the β-sheet signature in the

circular dichroism spectrum does not change significantly during cooling,we assumedthat theinternal energyof the β-sheet issimilar in the fibre and the plaque. Therefore, the enthalpy difference between lamellae and fibres must originate from the coupling of interactions among peptide segments and hydrophobic tails. This coupling is supported by our previous spectroscopic experiments that showed order can exist in the hydrophobic core of peptide amphiphile nanofibres and is enhanced by β-sheet orientation along the fibre axis22. The fibre architecture could also optimize interactions among peptide segments and alkyl segments. We

estimate 1HPA to be dominated by van der Waals forces, which are of the order of thermal energy. However, the increase in entropy

of the peptide amphiphile and water molecules in the plaque state can offset the enthalpy difference at elevated temperature. Specifically, the higher entropy in the plaque can originate in greater translation of water molecules (1Swater), because there is less water interface per peptide amphiphile molecule than in the

fibre structure, and therefore fewer restricted water molecules per amphiphile.Thiswouldreasonablypredictatransitiontemperature from fibrous to planar assemblies of the peptide amphiphiles (see Supplementary Information).

The observed rupture of the plaque at lower temperatures into bundles of fibres that give rise to an aqueous lyotropic liquid crystal at an unusually low concentration suggests an unusual mechanism of membrane rupture. The plaque observed by QFDE-TEM at 80 ◦C reveals a periodic surface texture with a characteristic spacing of about 7.5nm, which corresponds to the expected diameter of a single canonical nanofibre15. This strongly suggests the plaque results from the fusion of nanofibres as the dehydration occurs at elevated temperature. The microscopy also revealed the existence of ripples in the plaque of larger dimension than the fibres (Fig. 3a,b). We propose that fluctuations of the anisotropic plaque structure with 1D fibrous texture are crucial for its metamorphosis into arrays of highly aligned nanofibres. It is known that typically only membranes in curved geometries such as cylinders break by Rayleigh instabilities23; flat membranes generally rupture by creating holes24. Therefore, the possible breaking mechanism of a plaque into bundled fibres that gives rise to a lyotropic liquid crystal isuniquegivenitsunderlyinganisotropic1Dsubstructureimparted by the nanofibres composed of β-sheets. Long-range forces have been proposed to cause the rupture of surfactant membranes by means of concentration fluctuations25–27. The fibrous texture on the surface, however, generates an anisotropic surface tension, which may lead to the formation of waves of fluctuating composition on the surface similar to binary immiscible lipid membranes28. The waves resulting from the membrane tension appear as surface ripples when the β-sheets align, and this may generate the concentration of fluctuations required for rupture. However, the size of the successful composition fluctuation has to be large enough (larger than the membrane thickness D) to generate a critical size of a neck for rupture; otherwise the strain generated by the composition fluctuations in the internal structure (the interpenetrated bilayers) opposes the growth of the fluctuation29, and the lateral composition fluctuations are restored. One can assume that overall the breaking of the surface is due to a decrease in the overall surface tension γ, which is the change of free energy (F) as the interface area (A) increases, ∂F/∂A. That is, the bundled-fibre surface tension γf is lower than that of the lamellar γl because of the increase of hydration and order of peptide amphiphile molecules within the bundle. Unfortunately, the linear theory that assumes γ is constant under a deformation of the interface is not appropriate to describe the breaking of a lamella. To a first-order approximation one can assume that the interfacial energy of the fluctuation that leads to the rupture of the plaque is of the order of the γf. Consider fluctuations perpendicular to the lamellar surface plane (x,y) described by a function h(x,y), which induce a decrease in γl–γf that may result in the rupture of the plaque. To induce rupture,theresultingfree-energychange1F =Ff−F,isnegative,or 1F <0,whereFf isthefreeenergyofthefluctuatingplaquegivenby where hx = ∂h/∂x and hy = ∂h/∂y, and F the flat-plaque free energy is given by,

If we assume a 1D fluctuation along x of maximum amplitude we find in equation (2) that after the integration of both Ff and F, breaking occurs if 1F/A60, or rupture and all λ<λc would lead to stable plaques. As h0 =D/2 at rupture and there is no long-range diffusion in the rupture process, λc is the most probable size for rupture, and, given that γf/(γl−γf) is of the order of one (see Supplementary Information), λc is of the order of the membrane thickness D, in agreement with the expected sizeofthebreakingofacylinderthroughaRayleighinstability.

(Parte 1 de 2)

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