Controlling the multiscale structure of nanofibrous fibrinogen scaffolds

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Controlling the multiscale structure of nanofibrous fibrinogen scaffolds for wound healing Karsten Stapelfeldt, Stephani Stamboroski, Irina Walter, Naiana Suter, Thomas Kowalik, Monika Michaelis, and Dorothea Brueggemann Nano Lett., Just Accepted Manuscript • DOI: 10.1021/acs.nanolett.9b02798 • Publication Date (Web): 16 Aug 2019 Downloaded from pubs.acs.org on August 16, 2019

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Controlling the multiscale structure of nanofibrous fibrinogen scaffolds for wound healing Karsten Stapelfeldt+ ,† Stephani Stamboroski+ ,‡ Irina Walter,† Naiana Suter,† Thomas Kowalik,‡ Monika Michaelis,∗,¶,§ and Dorothea Brüggemann∗,†,k †Institute for Biophysics, University of Bremen, Otto-Hahn-Allee 1, 28359 Bremen, Germany ‡Fraunhofer Institute for Manufacturing Technology and Advanced Materials, Wiener Strasse 12, 28359 Bremen, Germany ¶Interdisciplinary Biomedical Research Centre, Nottingham Trent University, Clifton Lane, Nottingham NG11 8NS, U.K. §Hybrid Materials Interfaces Group, University of Bremen, Am Fallturm 1, 28359 Bremen, Germany kMAPEX Center for Materials and Processes, University of Bremen, 28359 Bremen, Germany E-mail: [email protected]; [email protected] Phone: +49 (0)421 218-64578; +49 (0)421 218-62286

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Abstract As a key player in blood coagulation and tissue repair fibrinogen has gained increasing attention to develop nanofibrous biomaterial scaffolds for wound healing. Current techniques to prepare protein nanofibers, like electrospinning or extrusion, are known to induce lasting changes in the protein conformation. Often, such secondary changes are associated with amyloid transitions, which can evoke unwanted disease mechanisms. Starting from our recently introduced technique to self-assemble fibrinogen scaffolds in physiological salt buffers, we here investigated the morphology and secondary structure of our novel fibrinogen nanofibers. Aiming at optimum self-assembly conditions for wound healing scaffolds, we studied the influence of fibrinogen concentration and pH on the protein conformation. Using circular dichroism and Fourier-transform infrared spectroscopy we observed partial transitions from α-helical structures to β-strands upon fiber formation. Interestingly, a staining with thioflavin T revealed that this conformational transition was not associated with any amyloid formation. Towards novel scaffolds for wound healing, which are stable in aqueous environment, we also introduced cross-linking of fibrinogen scaffolds in formaldehyde vapor. This treatment allowed us to maintain the nanofibrous morphology while the conformation of fibrinogen nanofibers was re-developed towards a more native state after rehydration. Altogether, self-assembled fibrinogen scaffolds are excellent candidates for novel wound healing systems since their multiscale structure can be well controlled without inducing any pathogenic amyloid transitions.

Keywords self-assembly, fibers, fibrinogen, secondary structure, solid-state CD, FTIR

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Current approaches for the therapy of acute and chronic wounds have benefited immensely from recent advances in nanotechnology. 1,2 Over the past years a variety of synthetic and natural nanomaterials was found to promote wound healing and tissue repair. 3,4 In particular, nanofiber mats and scaffolds offer promising strategies for regenerative medicine as they mimic the nanoarchitecture of native blood clots and many tissues. 5,6 During wound healing in vivo, blood coagulation induces the formation of nanofibrous fibrin clots upon enzymatic cleavage of the plasma protein fibrinogen. 7 Together with thrombocyte aggregation hemostasis takes place, which initiates wound closure. 8 During this process the nanofibrous blood clot functions as a provisional extracellular matrix (ECM), into which fibroblasts and endothelial cells migrate to facilitate tissue regeneration. 8,9

Being a trigger factor in blood coagulation and wound healing, fibrinogen has become increasingly attractive to develop nanoscaffolds for wound treatment. 10,11 Previously, fibrinogen nanofibers were fabricated with extrusion through ceramic nanopores 12 or using the well-established method of electrospinning. 13–15 We could previously show that extrusion induces lasting changes in the protein conformation. 16 With electrospinning nanofibrous fibrinogen mats in the size of several centimeters have already been fabricated. 13 Nevertheless, despite its simple use, electrospinning also often introduces conformational changes in the native protein structure due to the use of organic solvents. 17 In combination with the high electric fields used during electrospinning, organic solvents can lead to significant changes in the biofunctionality of nanofibrous scaffolds and can even induce protein degradation. 18 Beyond electrospinning it was previously reported that acidic buffer conditions, 19 organic solvents 20 and various hydrophobic surfaces 21–23 can induce the assembly of fibrinogen into fibrils and fiber scaffolds. Nevertheless, these versatile approaches mainly yielded fibrinogen scaffolds with low fiber density.

Recently, self-assembly of proteins into amyloid fibrils has been discussed to achieve

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tailored functions of nanofibrous biomaterials. 24 However, similar to electrospinning, the self-assembly of amyloid fibers is based on considerable changes in the protein structure. Amyloids are characterized by long, insoluble, unbranched fibers, which are rich in β-sheets, wherein each β-strand is arranged perpendicular to the fiber axis. 25,26 Many diseases are associated with amyloid formation, which are therefore known as protein misfolding disorders. 27 Such disorders often share common molecular mechanisms with neurodegenerative prion diseases where misfolded protein aggregates are known to be infectious. 28,29 Various protein-based biomaterials are also discussed to potentially exhibit amyloidogenic properties due to structural changes. 30 Current research on the occurrence of Alzheimer’s disease, for instance, suggests that fibrinogen is infected by β-amyloids, which results in the oligomerization of fibrinogen. 31,32 Hence, to tailor nanofibrous fibrinogen scaffolds for regenerative medicine without evoking any disease mechanisms, it is crucial to understand the underlying mechanisms of amyloid-induced conformational transitions .

When fibrinogen is enzymatically converted to fibrin in vitro no significant changes in the secondary structure occur during fibrillogenesis. 33 Yet, changes in the quaternary structure take place when fibrinopeptides A and B are cleaved off during in vivo fibrillogenesis. 7 However, controlled deformations upon fibrin clot formation in vivo are accompanied by a conformational conversion from α-helices to β-strands. 34,35 Interestingly, it was recently established that the adhesion of thrombocytes to fibrinogen crucially depends on the protein conformation. 36 These results indicate that future fibrinogen scaffolds for wound healing applications do not only require a biomimetic nanoarchitecture but also a precise control of the secondary structure. Hence, innovative scaffold design will involve a multiscale approach, which reaches from the molecular structure via the nano- and microscale to the macroscopic scaffold dimensions.

We recently introduced a novel in vitro method to fabricate nanofibrous fibrinogen scaf-

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folds with overall dimensions in the centimeter range using salt-induced self-assembly. 37 In contrast to other self-assembly routines 19–23 this process leads to dense fibrinogen nanofibers with well-controlled macroscopic scaffold dimensions. After we previously introduced saltinduced self-assembly as proof-of-concept to control nanofiber assembly on the nano- and microscale we now need to investigate whether these morphological effects are accompanied by changes in the secondary structure. These findings will enable us to tailor our novel biomaterial class to promote wound healing on different length scales, i.e. from the secondary structure via the nanoarchitecture towards the macroscale. Hence, we combined the two well-established techniques of circular dichroism (CD) and Fourier-transform infrared (FTIR) spectroscopy 38 to analyze the secondary structure of dried fibrinogen scaffolds. Although CD spectroscopy is commonly applied to proteins in solution, 39–41 solid-state CD investigations of dried protein films have also been performed 42,43 including studies on post-adsorptive transitions in the fibrinogen conformation. 44,45 FTIR spectroscopy has previously been used for the structural analysis of fibrinogen, 37,46,47 fibrin 48,49 and fibrinogen fragments 50 and is therefore also well-suited to study the secondary structure of our novel self-assembled fibrinogen scaffolds.

Substrate preparation CD experiments were carried out in Suprasil quartz cuvettes (Hellma UK Ltd.), which were cleaned in a UV-ozone cleaner (BioForce Nanosciences Inc., Salt Lake City, United States). For FTIR experiments 15 mm diameter glass slides (VWR, Darmstadt, Germany) were sputter-coated in an EM ACE600 high vacuum sputter coater (Leica Microsystems, Wetzlar, Germany) with a 5 nm adhesion layer of chromium and 50 nm of gold. For morphological analysis of fibrinogen scaffolds with scanning electron microscopy (SEM) we either used gold substrates as they were prepared for FTIR experiments or we used 15 mm diameter glass slides, which were cleaned by immersion into a 3:1 mixture of 95% sulfuric acid (VWR) and 30% hydrogen peroxide solution (VWR) for 5 min. For rehydration studies the cleaned glass slides and CD cuvettes were further modified with (3-

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Aminopropyl)triethoxysilane (APTES, Sigma, Steinheim, Germany) by overnight incubation in an ethanol solution containing 5% APTES. Unbound APTES was subsequently removed by washing the glass slides three times with pure ethanol for 5 min.

Preparation of fibrinogen scaffolds Nanofibrous fibrinogen scaffolds were fabricated by our recently introduced salt-induced self-assembly process 37 and planar fibrinogen layers were prepared by incubation in an aqueous fibrinogen solution. In brief, fibrinogen (100% clottable, Merck, Darmstadt, Germany) was dissolved in NH4 HCO3 (Roth, Karlsruhe, Germany) and dialyzed overnight against NH4 HCO3 using 14 kDa cut-off cellulose membrane dialysis tubing (Sigma). All solutions were prepared with deionized water from a TKA water purification system (Thermo Fisher Scientific, Schwerte, Germany). Planar fibrinogen layers were prepared by drying a final concentration of 5 mg ml−1 fibrinogen in 5 mM NH4 HCO3 for 12 h using a self-built climate chamber at a relative humidity of 30 %. Fibrinogen nanofiber scaffolds were prepared in the presence of 2.5x PBS (Thermo Fisher, pH 7.4) during the 12 h drying process. Fibrinogen samples for CD experiments were directly prepared in Suprasil quartz cuvettes (Hellma UK Ltd.) by drying a total volume of 40 µl. For morphological analysis with SEM and for FTIR spectroscopy a final volume of 200 µl fibrinogen solution was dried on gold-coated glass slides. In further experiments the fibrinogen concentration as well as the pH value of the PBS buffer were varied to study the influence of these parameters on the protein conformation. Samples, which were rehydrated, were cross-linked in formaldehyde vapor beforhand. For this procedure the samples were placed in a sealed beaker for 2 h. 1 µl of a 37% formaldehyde solution per cm3 was added into the beaker and allowed to evaporate. After crosslinking the samples were washed with deionized water. Cross-linked samples for CD analysis were prepared and rehydrated in APTES-modified cuvettes, to which 40 µl of deionized water were added before the cuvette was closed. Fibrin samples were directly prepared in a 0.01 mm path length CD cuvette by incubating 5 mg ml−1 fibrinogen solution in 5 mM NH4 HCO3 in the presence of 25 U ml−1 thrombin (Sigma) for 15 min.

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Circular dichroism spectroscopy CD spectra were recorded with an Chirascan spectrometer (Applied Photophysics, UK) running the Pro-Data Chirascan software (v4.2.22). For solid-state CD measurements we followed the procedure summarized in the SI (see S2: Methods). In brief, at least three repeat scans for each sample were measured at 25◦ C over the wavelength range of 190 to 250 nm using intervals of 1 nm and Suprasil quartz cells with a path length of 0.01 mm. The scans were averaged, baseline-corrected and the resulting net spectra were smoothed with a Savitsky-Golay filter using smoothing windows of 5 to 10 data points. The mean residue ellipticity (ΘM RE ) was defined as: 39

ΘM RE =

Θ c·l·n

(1)

where Θ is the raw CD ellipticity and n is the number of amino acids in the protein. In this case we describe l as the thickness of the protein films and c was estimated based on the UV absorbance of the films at 214 nm. 51 To estimate the contribution of specific secondary conformational components in the samples the measured CD spectra were analyzed using the BeStSel webserver. 52,53

Fourier-transform infrared reflection spectroscopy FTIR spectra were measured with a Bruker Vertex 70 system with an IR Scope II microscope (Bruker, Ettlingen, Germany) to analyze the energy absorption of fibrinogen scaffolds, which were prepared under varying self-assembly conditions. Spectra were recorded at 4 cm−1 resolution and 64 scans per measurement. At least three samples of each parameter setting were measured in 15 different positions following a zig-zag pattern. The reference spectrum was measured against air. After subtraction of water vapor absorbencies with OPUS included atmospheric compensation function, spectra were smoothed by a 7 to 17 point Savitsky-Golay function,

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baseline-corrected by rubber band baseline correction and averaged. Amide I and amide II peak positions were determined by peak integration using the Origin 9.0 software (OriginLab Northampton, USA). The evaluation of the secondary structure of amide I peaks was performed in the averaged spectra using curve fitting and second derivative spectral analysis with Origin software. Amide I bands were normalized and baseline-corrected. Peak positions were determined by second-derivative analysis. Curve fitting was performed using a non-linear least-squares fitting applying Gaussian band shape with peaks allowed to assume any height and area, but half height band width restricted to 11 to 15 cm−1 . The peaks obtained from deconvolution were then assigned to specific secondary structures based on previous FTIR studies (e.g. Roach et al., 46 Litvinov et al. 48 or Belton et al. 54 ). The quantification of each secondary structure was calculated by dividing the area of the peak of each component by the total area of the amide I peak.

Morphological analysis of fibrinogen scaffolds The morphology of planar fibrinogen layers and nanofibrous fibrinogen scaffolds was analyzed using scanning electron microscopy (SEM). A Bal-Tec SCD 005 sputter system (Leica Microsystems, Wetzlar, Germany) was used to sputter-coat the samples with a 7 nm gold layer before they were analyzed with acceleration voltages of 3 kV in a Zeiss Auriga field emission microscope (Carl Zeiss, Oberkochen, Germany). An MFP3D AFM (Asylum Research, Santa Barbara, CA, USA) was used to measure the topography of nanofibrous fibrinogen scaffolds in dry and wet conditions. An optical light microscope was combined to the AFM to control the scanning process. Glass slides were silanized with APTES prior to the assembly of nanofibrous fibrinogen scaffolds. Subsequently, the surface topography of dried fibrinogen scaffolds was analyzed in ambient air. Topographical analysis in wet environment was carried out after rehydration in water. We used silicon nitride cantilevers (MLCT Bio, Bruker, Wissembourg, France) with average nominal resonant frequencies of 38 kHz and a nominal spring constant of 0.1 Nm−1 . Height

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profiles of all samples were obtained in contact mode with a scan rate of 1 Hz and 256 scanning lines.

To achieve our aim of controlling the nanoarchitecture and secondary structure of selfassembled fibrinogen scaffolds, we studied conformational transitions from planar to nanofibrous fibrinogen. First, we focused on the influence of protein concentration and pH during the self-assembly process to identify optimum conditions for fabricating wound healing scaffolds. To ensure scaffold longevity with regard to future in vivo applications in regenerative medicine we also analyzed the influence of cross-linking and rehydration on the secondary structure.

Secondary structure of planar and nanofibrous fibrinogen scaffolds Fibrinogen nanofibers were prepared by drying fibrinogen (c.f. Figure S-2 for reference structure) in the presence of concentrated phosphate buffered saline (PBS) solution, while planar fibrinogen was prepared by a drying step in the absence of PBS. 37 The results of the secondary structure analysis of planar and fibrous fibrinogen with CD and FTIR spectroscopy are shown in Figure 1. The insets in Figure 1 show the results of scaffold analysis with scanning electron microscopy (SEM) and confirm the morphological differences between planar and fibrous fibrinogen 37 for both substrates, i.e. quartz glass (Figure 1A) and gold (Figure 1B). The CD spectra of planar fibrinogen layers show higher intensities than for fibrous fibrinogen scaffolds (Figure 1A). However, the characteristic spectral features remain unchanged also in comparison to the CD spectra in solution (c.f. Figure S-3), namely two minima around 222 and 210 nm and a maximum below 195 nm. 36

The effect of salt addition on the fibrinogen morphology can be clearly seen from FTIR spectroscopy. The FTIR spectra in the region between 1440 cm−1 and 1700 cm−1 , which contains the amide I band at ∼ 1650 cm−1 and the amide II peak at ∼ 1550 cm−1 , showed higher

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Figure 1: Comparison of conformational changes between planar fibrinogen and fibrinogen nanofibers (both 5 mg ml−1 no PBS for planar and 2.5x PBS pH 7.4 for nanofibers in accordance with Stapelfeldt et al.). 37 A: Results from CD spectroscopy, insets show the morphological differences using SEM analysis for fibrinogen on quartz glass; the bar chart summarizes the secondary structure obtained from BeStSel analysis. 52,53 B: Results from FTIR spectroscopy, insets show the morphological differences using SEM for fibrinogen on gold; the bar chart summarizes the secondary structure analysis using the deconvolution procedure described in the SI, S1: Methods and Figure S-1. intensities for planar than for nanofibrous fibrinogen in the amide I band (Figure 1B). The ratio between amide I/amide II intensities was 2.02 for planar fibrinogen and 1.19 for fibrous fibrinogen, which indicates an influence of the salt addition on the protein conformation. Upon transition from planar to nanofibrous fibrinogen the peak positions were found to be shifted towards lower wavenumbers. The amide I band was shifted from 1656 cm−1 to 1650 cm−1 while the amide II peak shifted from 1547 cm−1 to 1543 cm−1 , which is in good agreement with our previous study. 37 Furthermore, broadening of the amide I peak was observed upon fibrillogenesis. This change in shape could originate from the loss of homogeneity upon fiber formation, as proposed by Dutta et al.. 33 However, with 20 cm−1 the peak broadening in our study is stronger pronounced than the previously reported 5 cm−1 . The peak shifts and the change in amide peak shape are indicatives of conformational changes. 55,56 Moreover, as the dipole moments of the amide I and II bands are approxi-

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mately perpendicular to each other, the ratio of these two bands changes, if the adsorbed proteins change their conformation. 46 Hence, our findings from FTIR spectroscopy suggest that the morphological differences between planar and nanofibrous fibrinogen, which we found with SEM analysis, are accompanied by a change in protein secondary structure.

To ensure comparability between both spectroscopic methods, we summarized the secondary structure compounds obtained from deconvolution analysis to: α-helical content, content based on β-strands and ”other” contents. When the spectra obtained from CD and FTIR spectroscopy were deconvoluted both methods revealed similar structural features for our planar fibrinogen samples. Figure 1 clearly shows that the transition from planar to nanofibrous fibrinogen scaffolds resulted in a decrease of α-helical structures (from 23±1 % to 19±1 % for CD and from 28±3 % to 18±1 % for FTIR), which was accompanied by an increase in β-strands (from 28±1 % to 32±1 % for CD and from 25±1 % to 41±3 % for FTIR). Previously, a transition of α-helical structures to β-strands has been reported for the adsorption of fibrinogen to various surfaces, which would indicate an influence of the underlying substrate material in secondary structure changes. 44,46 Nevertheless, our novel method for the in vitro fabrication of fibrinogen nanofibers was found to be independent of the underlying substrate material as it mainly relies on the presence of salt ions during the drying process. 37 Hence, we conclude that the changes in secondary structure content, which we observed with CD and FTIR spectroscopy, are solely induced by the presence of salt during the drying process without any involvement of the underlying substrate.

The observed change in secondary structure upon salt-induced fiber formation clearly distinguishes our in vitro method from the in vivo formation of fibrin fibers. When fibrinogen polymerizes into fibrin after enzymatic cleavage by thrombin no conformational changes occur. 33 Despite this lack of conformational changes fibrin nanofibers are insoluble 57 whereas our self-assembled fibrinogen fibers dissolve again in aqueous environments. 37 This solubility

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of our scaffolds is the first indication for the absence of a potential amyloid transitions. 25,26 Additionally, the CD spectrum of amyloid fibers would be characterized by a single minimum at 218 nm with intensities below -20,000 deg cm2 dmol−1 , 26 which was not observed for our self-assembled nanofibers. To investigate further whether the structural changes, which occur during our salt-induced self-assembly process, might be related to amyloid transitions we carried out a standard fluorescence test using thioflavin T (ThT). 58 Despite the previously measured increase in β-strand content the ThT-staining did not reveal any characteristic shift of fluorescence emission to 480 nm, which would occur in the presence of amyloid structures (see Figure S-4). Based on this finding we conclude that the observed increase in β-strands during salt-induced fiber assembly does not lead to the formation of amyloid fibers. Hence, despite being rich in β-strand structures, our novel fibrinogen scaffolds are not composed of amyloid-like fibers and will most likely not trigger any disease mechanisms.

Concentration dependence of fibrinogen secondary structure Varying protein concentrations during the preparation of planar and nanofibrous fibrinogen scaffolds yielded different trends for the secondary structure (see Figure 2). Concentration-dependent CD measurements of planar fibrinogen are summarized in Figure 2A. The changes in the mean residue ellipticity for increasing protein concentrations are subtle, and the obtained CD spectra mainly overlap. Again, the characteristic wavelengths associated with minimum and maximum positions remain unchanged with respect to the CD spectra in solution (c.f. Figure S-3). The determination of secondary structures via the BeStSel server revealed maximum changes of about 2 % between the secondary structures of different planar scaffolds. In contrast, the CD analysis of nanofibrous fibrinogen scaffolds in Figure 2C showed a decrease of α-helices when the protein concentration was increased from 1 to 5 mg ml−1 . Concurrently, the β-strand content increased from low to high concentrations. Interestingly, with CD spectroscopy we identified the most pronounced conformational changes between 2 mg ml−1 and 3 mg ml−1 . These observations are also asso-

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Figure 2: Comparison of concentration-dependent conformational changes between planar fibrinogen (no PBS) and fibrinogen fibers (2.5x PBS). A: Results from CD spectroscopy for planar samples with a bar chart summarizing the secondary structure analysis obtained from BeStSel. 52,53 B: Results from FTIR spectroscopy for planar scaffolds with a bar chart summarizing the secondary structure analysis using the deconvolution procedure described in the SI, S1: Methods and Figure S-1. C: CD spectroscopy results for nanofibrous fibrinogen and a bar chart summarizing the secondary structure. D: Results from FTIR spectroscopy for nanofibrous samples with a bar chart to summarize the secondary structure.

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ciated with a slight reduction in the contributions of ”other” secondary structure compounds. Furthermore, we could observe a change in the extinction coefficients obtained from the absorbance at 214 nm (c.f. Figure S-5) between 2 mg ml−1 and 3 mg ml−1 . These findings are in very good agreement with our previous morphological studies, 37 which identified that a minimum concentration of 2 mg ml−1 on glass substrates was necessary to obtain fibrinogen fibers. In the present study we could also confirm this concentration-dependent morphological transition for fibrinogen scaffolds on gold substrates (c.f. Figure S-6) as they were used for subsequent FTIR analysis.

During FTIR spectroscopy we observed that the intensities of the amide I and II bands increased with increasing protein concentration for both, planar and nanofibrous, fibrinogen scaffolds (Figure 2B and D). For all concentrations the amide I and II peaks of planar fibrinogen were found to be more narrow than for fibrinogen nanofibers, which exhibited a broader shape. No evident change in the amide I and II peak positions was identified in both, planar and nanofibrous, fibrinogen when the concentration was varied. The deconvolution of the amide I peak revealed an α-helix content between 25 and 29% for planar fibrinogen and between 18 and 21% for fibrinogen nanofibers. The β-strand content of planar fibrinogen was in the range of 25 to 32% while fibrinogen nanofibers contained between 40 and 43% of β-strands.

In contrast to the structure dependence of the protein concentration, which we found with CD spectroscopy, we did not observe any conformational transition regime with FTIR spectroscopy - neither for planar nor for nanofibrous fibrinogen. However, both methods agree in terms of a general decrease in α-helicity and an associated increase of β-strands in the presence of PBS, i.e. upon transition from planar to nanofibrous fibrinogen. We attribute the observed differences in concentration dependence between CD and FTIR to the intrinsically different regions of interest, which were investigated with both methods.

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Within solid-state CD measurements spectra were obtained and averaged from the whole cuvette, while FTIR analysis was carried out in 15 localized scaffold positions, which were then averaged.

Secondary structure of fibrinogen nanofibers under varying pH conditions For many different proteins and peptides self-assembly of nanofibers has been found to be sensitive for pH changes. 26,48,59,60 Therefore, we studied the influence of pH on the secondary structure during fibrinogen fiber assembly using a physiological range from pH 5 to 9 (see Figure 3).

CD analysis revealed a general decrease of the intensity from pH 5 to pH 9. Similar to the concentration dependence, the characteristic wavelengths associated with minimum and maximum positions remained unchanged in comparison to the CD spectra in solution (c.f. Figure S-3). Subsequent analysis of the CD spectra via the BeStSel server 52,53 revealed a clear dependence of the structural features from the pH value. A decrease in α-helicity from low to high pH values (from 25±1 % to 18±1 %) was accompanied by an increase in β-strands (from 26±1 % to 30±1 %). Interestingly, we identified a transition regime in the secondary structure content between pH 7 and 8. This regime was found to coincide with our morphological SEM analysis of fibers on gold, which showed an onset of fiber formation from pH 7 onwards (see Figure S-7) that was well in agreement with our previous study on glass. 37

The FTIR spectra of fibrinogen nanofibers, which were prepared under varying pH conditions, are shown in Figure 3B together with the evaluation of the secondary structure content obtained from deconvolution of the amide I band. While the shape of the amide I bands was not found to depend on the pH value we observed an increase in peak intensity from low to high pH values. For all pH values, the secondary structure analysis yielded structure contents between 16 and 21% for α-helices and between 35 and 43% for β-strands, both being

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Figure 3: Comparison of pH-dependent conformational changes of fibrinogen nanofibers prepared with 2.5x PBS. A: Results from CD spectroscopy with a bar chart summarizing the secondary structure analysis obtained from BeStSel. 52,53 B: Results from FTIR spectroscopy with a bar chart summarizing the secondary structure analysis using the deconvolution procedure described in the SI, S1: Methods and Figure S-1. characteristic for the formation of nanofibers (see Figure 1B). Although CD spectroscopy and SEM analysis revealed a conformational and morphological transition state between pH 7 and 8, this transition towards fibrous features was not confirmed by FTIR analysis. Similar to the previous concentration dependence, this can be correlated with the different analysis modes and varying regions of interest used in CD and FTIR spectroscopy.

The isoelectric point (IEP) of fibrinogen was reported to be at pH 5.8. 61 Interestingly, if the pH value during fiber assembly was below the IEP of fibrinogen or close to it, no fiber formation was obtained. On the other hand, with pH values above the IEP, fiber formation could be induced. Thus, the observed onset of fibrillogenesis might be correlated to conformational changes in the fibrinogen molecule, which occur upon change of the pH value in the presence of salt. Previously, it was established from numerical simulations and hydrodynamic measurements that fibrinogen changes from a compact into a more elongated shape when the pH value increases from neutral to more basic pH. 62 Hence, it is well possible that this change in the molecular shape promotes the tendency of individual fibrinogen molecules

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to align into nanofibers.

Influence of cross-linking on the secondary structure of nanofibrous fibrinogen scaffolds For future studies of cell adhesion, which will be carried out in aqueous environment, non-soluble fibrinogen fibers are required. In contrast to extruded nanofibers of fibrinogen and other ECM proteins, which were stable in aqueous solutions, 12,16 we reported recently that self-assembled fibrinogen nanofibers dissolve again after being rehydrated. 37 Nevertheless, to achieve non-soluble protein scaffolds cross-linking is a well-established method, which has previously been shown to increase the strength and lifespan of fibrous fibrinogen scaffolds. 63 Hence, we introduced an additional cross-linking step after the fiber assembly. Nanofibrous scaffolds were cross-linked with formaldehyde vapour and afterwards thoroughly washed with water to remove salt crystals and, in a second step, dried again. During this process, the morphology of the non cross-linked and cross-linked samples remained largely unchanged and the nanofibrous fibrinogen scaffolds were maintained as SEM analysis revealed (c.f. Figure 4A and B). A comparison of the associated CD spectra in Figure 4C revealed an almost perfect overlap, which was obtained by a careful baseline correction considering the removal of salt crystals during washing. Moreover, deconvolution yielded deviations of only 1 % between non cross-linked and cross-linked fibrinogen scaffolds. The corresponding FTIR results are depicted in Figure 4D. In agreement with the CD results, amide I and II peaks maintained their shape and peak positions, thus indicating no structural changes of the fibers after cross-linking. This fact is further approved by the deconvolution analysis of the amide I peak where no differences in the structural content were found between non cross-linked and cross-linked fibrinogen fibers.

Secondary structure of dried and wet fibrinogen nanofibers In the last step towards the application of nanofibrous fibrinogen scaffolds for wound healing, we investi-

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Figure 4: Influence of cross-linking on the morphology and conformation of nanofibrous fibrinogen scaffolds. SEM images of non cross-linked fibers and fibers, which were crosslinked with formaldehyde (FA) vapour (A and B). C: Results from CD spectroscopy with a bar chart summarizing the secondary structure analysis obtained from BeStSel. 52,53 D: Results from FTIR spectroscopy with a bar chart summarizing the secondary structure analysis using the deconvolution procedure described in the SI.

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gated the morphology and conformation of fixated nanofibers upon rehydration in an aqueous environment. For this purpose, we introduced an additional silanization step with (3Aminopropyl)triethoxysilane (APTES) on the glass slides prior to fiber assembly to immobilize the resulting scaffolds for subsequent rehydration. We could previously show that this step does not have any influence on the nanofiber morphology. 37 Using CD spectroscopy we could now establish that the presence of the underlying APTES layer did not change the secondary structure of self-assembled fibrinogen nanofibers (c.f. Figure S-8).

The morphology of dried fibrinogen scaffolds obtained from AFM imaging (c.f. Figure 5A) agrees well with our previous results obtained via SEM analysis. When fibrinogen nanofibers were rehydrated in water we observed a swelling of the scaffolds while the nanofibrous morphology was preserved (c.f. Figure 5B). In Figure 5C we compare the CD spectra of fibrinogen in solution, nanofibrous fibrinogen and rehydrated cross-linked fibers. After rehydration we obtained signal intensities in the wavelength region above 205 nm, which were more similar to the CD spectrum in solution than to nanofibrous fibrinogen. Yet, minor deviations in signal intensities were found between 205 to 215 nm. However, in the wavelength region below 205 nm we found an overlap of the CD signals from non-rehydrated and rehydrated fibrinogen nanofibers. This wavelength region is generally attributed to changes in the π → π ∗ transitions along the C=O bond of the protein backbone. Nevertheless, also adjacent bonding in close proximity to the chromophores can influence the CD signal. 39,64 Hence, the observed changes below 205 nm might indicate that the conformational transitions associated with drying in salt solutions are reversible to a certain extent. The remaining changes could potentially be attributed to protein/surface or protein/protein interaction. To compare the secondary structure of self-assembled fibrinogen nanofibers we also measured the CD spectrum of native fibrin, i.e. fibrinogen, which was enzymatically converted by thrombin. This thrombin-induced transition leads to a CD spectrum very similar to the spectrum of native fibrinogen in solution. The main difference can be found in the wave-

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Figure 5: Comparison of rehydration effects on the morphology and conformation of crosslinked fibrinogen fibers. A: AFM image of the the dried fibers and B: of the rehydrated fibers, identical scale bar for both images. C: Comparison of the CD spectra for fibrinogen in solution, nanofibrous fibrinogen, rehydrated cross-linked fibers and fibrin. length region below 195 nm (c.f. Figure 5C). This finding is in good agreement with Dutta et al., 33 who proposed the absence of conformational changes in fibrin based on FTIR measurements. Hence, we ensured that our well-controlled method to self-assemble fibrinogen into nanofibrous scaffolds leads to secondary structures close to the physiological conditions observed for native fibrin.

Towards a new generation of nanofibrous wound healing scaffolds it will be highly important to study the interaction of self-assembled fibrinogen fibers with cell types, which are key players in blood coagulation and tissue regeneration. Based on our important result that salt-induced self-assembly does not induce any major changes in the protein conformation we assume that our novel fibrinogen scaffolds will be well-suited to promote the adhesion of thrombocytes, 36 fibroblasts 14 or endothelial cells 15 in future cell culture studies. Moreover, self-assembled fibrinogen scaffolds could serve as well-controllable in vitro model system for drug screening in hemostatic therapy or future studies on amyloid-related disease mechanisms.

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In conclusion, by combining CD and FTIR spectroscopy we showed that salt-induced selfassembly of fibrinogen nanofibers leads to partial transitions from α-helices to β-strands in the protein conformation. This powerful combination of CD and FTIR spectroscopy enabled us to reveal structural changes in fibrinogen nanofibers on distinct length scales and with varying resolution. Moreover, we could correlate the conformational trend in the presence of salt with a morphological transition from planar to nanofibrous fibrinogen scaffolds. Despite the observed structural changes no amyloid formation was observed during salt-induced selfassembly of fibrinogen nanofibers. Towards tailored fibrinogen scaffolds for wound healing applications we studied the influence of protein concentration and pH on structural changes during self-assembly. Furthermore, we introduced cross-linking in formaldehyde vapor and rehydration to obtain nanofibrous scaffolds, which were stable in aqueous environment. This post-treatment preserved the nanoarchitecture while structural changes of fibrinogen were partially reversed to a more conformationally native state. In summary, we were able to tailor the nanomorphology and the secondary structure of fibrinogen nanofibers using salt-induced self-assembly without inducing any pathogenic amyloid transitions. Hence, we established a multiscale wound healing platform, which can be well controlled from the molecular level via the nanomorphology to the macroscale.

Author information +

K.S. and S.S. contributed equally to this work.

Acknowledgement The authors thank Prof. Lucio Colombi Ciacchi for access to the CD spectroscope and Prof. Manfred Radmacher for his support with atomic force microscopy analysis. Annette 21


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Peter is gratefully acknowledged for her support in confocal microscopy. K.S., N.S. and D.B. gratefully acknowledge funding by the Emmy Noether program of the German Research Council under grant number BR5043/1-1.

Supporting Information Available S1: Methods; 1. CD and 2. FTIR S2: Results; including 1. reference structure of fibrinogen, 2. CD spectra of fibrinogen in solution, 3. Staining with thioflavin , 4. UV-vis absorbance 5. concentration-dependent SEM images, 6. pH-dependent SEM images and 7. APTES influence 8. Thrombin and fibrinopeptides.

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Controlling the multiscale structure of nanofibrous fibrinogen scaffolds

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