Resemblance of Electrospun Collagen Nanofibers to Their Native

Dec 20, 2012 - Abby Chainani , Kirk J. Hippensteel , Alysha Kishan , N. William Garrigues , David Simms Ruch , Farshid Guilak , Dianne Little...
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Resemblance of Electrospun Collagen Nanofibers to Their Native Structure Jochen Bürck,† Stefan Heissler,‡ Udo Geckle,§ Mohammad Fotouhi Ardakani,∥ Reinhard Schneider,∥ Anne S. Ulrich,†,⊥ and Murat Kazanci*,# †

Institute of Biological Interfaces IBG2, ‡Institute of Functional Interfaces IFG, §Institute of Applied Materials, Energy St. Sys. IAM-ESS, #Institute of Biological Interfaces IBG1, Karlsruhe Institute of Technology (KIT), D-76021 Karlsruhe, Germany ∥ Laboratory for Electron Microscopy LEM, ⊥Institute of Organic Chemistry, Karlsruhe Institute of Technology (KIT), D-76131 Karlsruhe, Germany ABSTRACT: Electrospinning is a promising method to mimic the native structure of the extracellular matrix. Collagen is the material of choice, since it is a natural fibrous structural protein. It is an open question how much the spinning process preserves or alters the native structure of collagen. There are conflicting results in the literature, mainly due to the different solvent systems in use and due to the fact that gelatin is employed as a reference state for the completely unfolded state of collagen in calculations. Here we used circular dichroism (CD) and Fourier-transform infrared spectroscopy (FTIR) to investigate the structure of regenerated collagen samples and scanning electron microscopy (SEM) and transmission electron microscopy (TEM) to illuminate the electrospun nanofibers. Collagen is mostly composed of folded and unfolded structures with different ratios, depending on the applied temperature. Therefore, CD spectra were acquired as a temperature series during thermal denaturation of native calf skin collagen type I and used as a reference basis to extract the degree of collagen folding in the regenerated electrospun samples. We discussed three different approaches to determine the folded fraction of collagen, based on CD spectra of collagen from 185 to 260 nm, since it would not be sufficient to obtain simply the fraction of folded structure θ from the ellipticity at a single wavelength of 221.5 nm. We demonstrated that collagen almost completely unfolded in fluorinated solvents and partially preserved its folded structure θ in HAc/EtOH. However, during the spinning process it refolded and the PP-II fraction increased. Nevertheless, it did not exceed 42% as deduced from the different secondary structure evaluation methods, discussed here. PP-II fractions in electrospun collagen nanofibers were almost same, being independent from the initial solvent systems which were used to solubilize the collagen for electrospinning process. formed by two α1 (I) chains and one α2 (I) chain.3,4 Among these collagens, type I collagen is the main structural element of the extracellular matrix. Collagen type I accounts for up to 70− 90% of the collagen found in the body, and it is present in the form of elongated fibers in various tissues. These building blocks are rod-like triple helices that are stabilized by intramolecular hydrogen bonds between Gly and Hyp in adjacent chains.5 Extracted type I collagen is favored for biomedical applications, since in vitro under appropriate conditions it will spontaneously self-assemble to form biodegradable and biocompatible insoluble fibrils of high mechanical strength, low immunogenicity, and with a Dperiodicity that is indistinguishable from that of native fibers.6−9 Electrospinning is a popular scaffold fabrication strategy in tissue engineering. In the electrospinning process, high voltage is applied to a liquid droplet and the body of the liquid becomes charged. The liquid deforms into a cone shape, called a “Taylor cone”, and emits a charged jet of liquid. The charged

1. INTRODUCTION The extracellular matrix (ECM) is the extracellular part of animal tissue that provides mechanical support and regulates cellular communication. It also segregates tissues from one another, supports and defines the intercellular spaces, and maintains the normal state of differentiation within the cellular compartment.1 The fabrication of synthetic biocompatible scaffolds that can mimic the native ECM for tissue engineering has attracted significant attention in recent years. In the past there were considerable efforts to develop suitable scaffolds for tissue engineering applications using different biodegradable polymers, collagen and polymer/collagen blends. Since collagen is a basic structural element in native extracellular matrices, its abundant presence in natural tissues, composing 30% by weight of body protein tissues,2 makes it a natural choice as a polymer for biomedical materials and tissue-engineering matrices. Collagen is also biodegradable and biocompatible and enhances cellular penetration and wound repair. Molecular collagen is made of three α chains, and the properties of these α chains define the material properties of the collagen. For example, type III collagen fibrils are composed of three α1 (III) chains, whereas type I collagen fibrils are © 2012 American Chemical Society

Received: August 16, 2012 Revised: December 20, 2012 Published: December 20, 2012 1562

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the stability of the triple helix.25 There is a need for a hydrophobic component that is miscible with water and yet has strong hydrophobic parts to separate triple helices from each other in order to break the collagen bundles into single triple helices. It is usually accepted that hydrophobic interactions are favored in positive entropy changes resulting from the release of structured water when nonpolar groups interact with one another. The effect of hydrophobic hydration is to enhance the solubility of nonpolar species and to disfavor their aggregation.26 Short-chain aliphatic alcohols seem to be good candidates as they are partially or totally miscible with water and exhibit a strong hydrophobic part. Aliphatic alcohols have also rather positive impact on collagen processing when used under the correct conditions.27 We investigated the effect of electrospinning solvent systems and of the spinning process on the structure of the electrospun collagen nanofibers. Moreover, by observing the structure of collagen at intermediate stages, we separated these two steps (preparation of the collagen solution, and formation of electrospun nanofibers) to distinguish their possible contribution to the denaturation of collagen. We included two further solvent systems, in addition to the fluoroalcohol solvents mentioned above. One of them was a phosphate buffer saline (PBS)/ethanol mixture that was introduced by Wnek and coworkers.28 The other one, which was used previously by Szentivanyi et al.29 and Buttafoco et al.,30 was employed with some modifications. Since, the electrospinning of natural molecules has proven to be somewhat of a challenge because they do not behave like classical polymers lacking the viscoelastic properties essential for stable electrospinning. The stability of their tertiary structure reduces their capacity to unravel in an extensional flow field, which in return prevents the viscoelastic response necessary for jet stabilization. Consequently, biomolecules are often blended and cospun with synthetic polymers.13,14 Buttafoco et al.30 report that fibers cannot be spun from 1 to 2% collagen in weak acidic solutions and that fiber formation is accomplished by adding high molecular weight PEO (900 kDa), which increases the viscosity of the solution, thus leading to the stable jets. In our application, we used a diluted acetic acid (0.5M) and ethanol mixture (1:1) to dissolve collagen and a polyethylene oxide blend (1:1) to prepare the electrospinning solution. We used circular dichroism (CD) spectropolarimetry, supported by IRspectroscopy, to observe the secondary structure of the collagen molecules. Previously, Zeugolis et al.5 has argued that the electrospun collagen scaffolds are not crystalline, triple-helical, and do not possess a high degree of axial alignment and exhibit a characteristic D-periodicity (the fingerprint of fibrous collagens). This is due to alternating overlap and gap zones, produce by the specific packing arrangement of the 300 nm long and 1.5 nm wide collagen molecules.8,31−35 They suggest that fluoroalcohols are the major cause of denaturation and conclude that electrospinning of collagen or cospinning of collagen-synthetic polymers out of fluoroalcohols results in the creation of gelatin, a protein derived from denatured collagen. In the present study, we investigated the effect of four different common solvent systems and of the electrospinning parameters on the secondary structure of collagen. Three different approaches for secondary structure estimation were applied to the CD spectra over the spectral range of 185−260 nm to obtain detailed quantitative information about the folded fraction of collagen, both in solution and after producing the

jet is then elongated in a whipping process and moves toward a grounded collector. Electrospinning is a suitable way to produce fibers with diameters smaller than 1 μm and has a number of advantages. Also electrospinning allows control over the fiber diameter and orientation of the fibers in a mesh.10 Such materials aim to mimic extracellular matrix components, such as collagen fibrils, whose diameter in vivo range from 20 nm to 40 μm.11,12 In order to regenerate collagen in the form of nanofibers, it is necessary to dissolve them in suitable solvents. Extensive research has been performed regarding why a particular solution will be spinnable or not. It has been concluded that a spinnable solution is one where the forming jet is sufficiently stable; that is, the filament will not break up before the final dry fiber is formed.13,14 Collagen can be easily dissolved in acidic conditions to produce an electrospinning solution. However, the slow evaporation rate of the acid and its strong affinity with collagen leads to the deposition of wet fibers on the target, which can partially weld together and lessen the porosity of the mat. Therefore, fluoroalcohols, such as 1,1,1,3,3,3-hexafluoro-2-propanol (HFIP)15−17 and 2,2,2trifluoroethanol (TFE),18 are proposed as alternative electrospinning solvents. The high evaporation rate and moderate affinity of these solvents make them very good solvents for electrospinning.19 Extracted fibrillar collagen maintains a fibrillar structure due to hydrophobic interactions as well as a network of water molecules around each triple helix bound to it by hydrogen bonds. It is proposed that fluoroalcohols act directly via hydrophobic and hydrophilic interactions to separate the triple helixes, hence dissolving the crystalline structure of collagen. Furthermore, the spacing between each polyproline-II (PP-II) helix in the coiled coil is rather large and makes it easy for small molecules such as fluoroalcohols to break hydrophobic interactions between PP-II helices. This bulk effect causes unfolding of the tertiary structure of the proteins, while concurrently strengthening some intramolecular hydrogen bonds involved with secondary structures, such as αhelixes. This combined effect results in the formation of what is called the “open-helical structure” in which the interaction between helix segments is weak, while the hydrophobic segments are mostly exposed to the solvents.14 Opening of the intramolecular disulfide bridges, together with the partial denaturing created by the TFE environment, produce a pronounced expansion of the protein. This, in turn, affected both the rheological properties of the solution and its spinnability.14 In addition, very high shear forces and strain rates (on the order of 103 s−1) acting during electrospinning process, induce conformational changes (strain induced crystallization). This dominant factor is believed to cause stretching and orientation/rearrangement of polymer chains, as indicated by in-process measurements of jets and limits the natural folding of the electrospun collagen nanofibers.20 It has been also shown earlier with different proteins that fluoroalcohols do not only denature the native structure but they also lower the denaturation temperature.21,22 Since gelatin is the water-soluble degradation product of the originally waterinsoluble collagen fibril, the observed water solubility of the electrospun collagen scaffolds might point to an extensive conformational change.5,23 It has been demonstrated that salt from buffer can replace the water network around the triple helix and enhance its stability24 when the collagen is dissolved in an acidic/buffer mixture. Salt is then a favorable element to stabilize the triple helix and does not have any negative impact on its solubility in acid. Increasing ionic strength even increases 1563

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grids of 3 mm in diameter. Due to the possible contrast tuning by energy-filtered TEM, electrospun collagen nanofibers were analyzed without staining, fixation and embedding. Circular Dichroism. CD spectra were recorded using a J-815 spectropolarimeter (Jasco Co., Tokyo, Japan). The instrument was routinely calibrated with a 0.06% (w/v) aqueous solution of ammonium D-10-(+)-camphor sulfonate at 290.5 nm. The spectra were scanned between 260 and 190 at 0.1 nm intervals. Three repeat scans at a scan rate of 10 nm min−1, 8 s response time, and 1 nm bandwidth were averaged for each sample and its respective blank. Samples of calf skin collagen type I in 0.05 M acetic acid, HFIP, TFE, and a blend with 0.1 mg of PEO in 1:1 (v/v) 0.05 M acetic acid/ ethanol, were prepared by weighing 1 mg of collagen and vortexing it with an appropriate volume of the respective solvent to get a 1 mg/mL stock solution. The collagen nanofiber solutions after electrospinning were prepared in a similar way by dissolving the weighed-in material in 0.05 M acetic acid. The final concentrations of all CD samples were adjusted to 0.1 mg/mL collagen by diluting the freshly prepared stock solution with an appropriate aliquot of the corresponding solvent and subsequent vortexing. The collagen concentration in the solutions was verified via the absorption of the respective samples measured at 192 nm to ensure the same concentration in all samples. CD spectra were collected using rectangular quartz glass cuvettes with 0.1 cm optical path length. Before testing collagen solution in TFE, HFIP, and HAc/ ethanol, the baseline spectra of pure TFE, HFIP, and 0.05 M acetic acid/ethanol (1:1, v/v) including 0.1 mg of PEO were used as a blank control for the corresponding native collagen samples. The averaged blank spectrum was subtracted from the averaged sample spectrum to get the corrected line shape. All spectra were recorded at 15 °C, using a rectangular cell holder, connected to a water thermostat. The thermal denaturation temperature series of native collagen in 0.05 M HAc was performed using the automated temperature scan program contained in the JASCO software. The heating rate was fixed to 0.5 K/ min, and at each selected temperature a 3 min delay was implemented to ensure thermal equilibrium in the cell. CD spectra were smoothened by the adaptive smoothing method, which is part of the Jasco Spectra Analysis software. Spectral deconvolution of the CD spectra of native collagen in 0.05 M acetic acid/ethanol, HFIP and TFE as well as of the electrospun collagen nanofibers, which were dissolved in 0.05 M HAc, was performed using the convex constraint algorithm (CCA plus).37,38 Attenuated Total Reflection Fourier-Transform Infrared Spectroscopy (ATR-FTIR). A Bruker Tensor 27 Fourier transform IR spectrometer (Bruker Optik GmbH, Ettlingen, Germany), with a Bruker Platinum ATR cell, was employed to obtain the IR spectra of these samples which were available in adequate amount. The cell was equipped with an ATR crystal made from diamond, onto which the samples were pressed directly. A total of 32 scans in the 4000−400 cm−1 spectral range were recorded with a scan velocity of 10 kHz and a spectral resolution of 4 cm−1. The reference spectra were acquired with the unloaded diamond crystal. To characterize very thin layers of nanofibers on a micrometer scale, the IR measurement was performed using a Bruker Hyperion 3000 FTIR microscope that provides a special ATR objective with a crystal tip made from germanium. A total of 32 scans in the 4000−600 cm−1 spectral range were recorded with a scan velocity of 20 kHz and a spectral resolution of 4 cm−1. The reference spectra were acquired with the unloaded germanium crystal. A Bruker Tensor 27 equipped with a Bio ATR 2 cell was used for the temperature dependent FTIR measurements of the collagen solutions. The special design of the cell provides temperature control of the samples. Spectra of each solution were measured at 20 and 40 °C. In order to obtain comparable/uniform data, all spectra were baseline corrected and normalized using the Bruker OPUS software, version 6.5.

nanofibers. The electrospun collagen nanofibers were illustrated by using SEM and TEM methods.

2. EXPERIMENTAL SECTION 2.1. Materials. Collagen type I was the gift from the Kensey Nash Corporation. Potassium chloride (KCl, ≥99.5%) was purchased from Carl Roth GmbH. Hydrochloric acid (HCl, 32%) and acetic acid (HAc, 99%) were purchased from Acros organics. Ethanol (≥99.8%), sodium chloride (NaCl, ≥99.5%, impurities were insoluble matter), sodium phosphate dibasic heptahydrate (Na2HPO4·7H2O, ≥99%), potassium phosphate monobasic (KH2PO4, 99.99%), 1,1,1,3,3,3hexafluoro-2-propanol (HFIP, ≥99.8%), 2,2,2-trifluoroethanol (TFE, ≥99%), and poly(ethylene oxide) (PEO, Mv ≈ 600 000) were all purchased from Sigma-Aldrich and used as received. 2.2. Preparation of Collagen Solutions. The collagen used in these experiments was a water-insoluble lyophilized foam powder consisting of tropocollagen extracted from bovine dermis (generously donated by the Kensey Nash Corporation, USA) and was used without further purification. Four different samples were prepared by dissolving collagen in different solvent systems: (1) 10% w/v collagen solution was prepared by dissolving collagen in HFIP (electrical conductivity: 111 μS/cm, @25 °C); (2) 10% w/v collagen solution was prepared by dissolving collagen in TFE (electrical conductivity: 250 μS/cm, @25 °C); (3) 15% w/v collagen solution was prepared by dissolving collagen in a mixture of phosphate buffer saline (PBS) and ethanol (3:2). PBS was prepared as described in ref 36 by dissolving 160 g of NaCl, 28.8 g of Na2HPO4, 4 g of KCl, and 4.8 g of KH2PO4 in 800 mL of deionized water, and distilled H2O was added to make 1 L (electrical conductivity: 5000 μS/cm, @25 °C); and finally (4) 2% collagen and 2% PEO blend was dissolved in 0.5 M HAc and ethanol mixture at 1:1 ratio (electrical conductivity: 200 μS/cm, @25 °C). 2.3. Electrospinning Procedure. The prepared electrospinning solutions were loaded into a 2 mL syringe (Omnifix, B. Braun, Melsungen, Germany) with a blunt end nozzle, controlled by a syringe pump (kd Scientific, Holliston, USA). The solution was pushed through a capillary blunt steel needle (21 gauge, 0.7 mm i.d. × 50 mm length) at a constant speed (between 0.2 and 0.5 μL/min). The steel needle was connected to a high voltage source (Spellman Bertan Series 205B High Voltage Electronics, NY, USA) by tungsten electrodes. The electric potential is needed to start the spinning process and thus form a jet. The applied DC voltage was held at 20−22 kV. Three steel rings with the same (positive) charge were placed at the same height and perpendicular to the needle tip in order to stabilize the jet and direct it downward. A Cu collector was placed 15 cm from the needle tip to collect the electrospun collagen nanofibers. The nanofiber meshes were collected on cover-glasses placed onto the collector. Collagen films were produced by casting 1% collagen solution in 0.5 M HAc on silicon molds. They were dried either at room temperature or 70 °C in an oven overnight. The films were used to observe thermal effects on the structure, using FTIR spectroscopy. 2.4. Characterization. Circular dichroism (CD) spectropolarimetry was employed to characterize the structure of collagen and nanofiber meshes and supported by Fourier-transform infrared spectroscopy (FTIR). Scanning Electron Microscopy (SEM). The morphology of the produced electrospun nanofibers was evaluated using Zeiss Supra 55 Scanning Electron Microscope (Carl Zeiss SMT AG, Oberkochen, Germany) after Au/Pd sputtering. Transmission Electron Microscopy (TEM). The microstructure of the electrospun nanofiber samples was characterized by means of transmission electron microscopy on a Zeiss LEO 922 Omega microscope which is equipped with an imaging in-column energy filter. The microscope was operated at 200 kV accelerating voltage, and TEM bright-field (BF) images were recorded with a 2K CCD camera (ProScan). To enhance the image contrast only elastically scattered electrons were used for imaging. TEM samples were directly prepared from the as-prepared electrospun collagen nanofibers by taking a piece of the fibers from the fiber mesh on the glass support by means of a pair of tweezers and putting it in between a sandwich of two copper

3. THEORETICAL SECTION One of the long-held criticisms of electrospinning has been the potential denaturing effect of organic solvents. Since electrospinning is a very popular method to produce scaffolds, we 1564

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Figure 1. SEM images of electrospun collage nanofibers from (a) HFIP, (b) TFE, (c) HAc/EtOH, and (d) PBS/EtOH.

(III) Convex constraint algorithm deconvolution with a reference set of collagen in the folded and full (thermally) unfolded state and dissolved in various solvents (CCA set-II). In the second and third procedure, we used the Convex constraint algorithm,37,38 which has been developed to deduce the spectral contribution of base spectra representing common secondary structure fold motifs directly and only using experimental CD curves of a representative set number of protein spectra. CCA is a general deconvolution method for a CD spectra set of any variety of conformational type. The algorithm, based on a set of three constraints, is able to deconvolute a set of CD curves to its common “pure” component curves and conformational weights. The following three constraints are used for the deconvolution: (a) The sum of the weight coefficients, C(i, j), for each conformer must be unity;

decided to investigate how the common electrospinning solvents and electrospinning process would affect the collagen structure and how much the native structure of collagen would be preserved. To be able to understand the behavior of collagen in different popular electrospinning solvents and to determine the degree of folding of the produced electrospun nanofibers, we compared three different secondary structure estimation procedures to analyze the CD spectra of native collagen and electrospun collagen nanofibers in relevant solvents. We called the first method: (I) Single wavelength estimation of collagen folding described in Ackerman et al.:39 We employed this method to extract the fraction of triple helix, mainly relying on the equilibrium state of collagen spectra. The fraction of folded collagen (θ) was defined as θ = (θobs − θu)/(θt − θu)

(1)

where θobs, θt, and θu represented the ellipticities of observed, triple helix, and unordered state, respectively, measured at a wavelength of 221.5 nm. θt and θu were measured at 10 and 90 °C, respectively, in 0.05 M HAc. The PP-II peak at 221.5 nm exhibited positive values at 10 °C and transformed to a negative ellipticity at 90 °C, which indicated that collagen completely unfolded at that temperature. Obviously, ellipticities at temperatures between 10 and 90 °C represented a combination of triple helix and random coil structures. (II) Convex constraint algorithm deconvolution with a reference (base) data set of CD spectra of natively folded and gradually denatured collagen samples obtained by heating the sample with distinct steps until 90 °C (CCA set-I).

p

∑ C(i , j) = 1 i=1

(2)

where j = 1, 2, ..., N, representing the number of the analyzed CD spectra. (b) C(i , j) ≥ 0.

(3)

(c) The points of {C(i, j), i = 1, ..., P}, j = 1, ..., N, must be embedded in a simplex of the P-dimensional Euclidean space with the smallest volume.38 1565

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4.1. Scanning Electron Microscopy (SEM) Results. Figure 1 demonstrates the SEM images of the electrospun nanofibers obtained from different solvents in this study. Electrospinning yielded randomly oriented and interconnected fibrous meshes with the fiber diameter in the nanometer range. The electrospun nanofiber diameters were mainly determined by the collagen concentrations and electrospinning solvents. For example, 10% collagen solution in HFIP and TFE produced electrospun nanofiber meshes with the fiber diameter ranging between 150 and 200 nm (Figure 1a,b), 2% collagen + 2% PEO in HAc/EtOH produced electrospun nanofiber meshes with the fiber diameter ranging between 50 and 100 nm (Figure 1c) and 15% collagen in PBS/EtOH produced electrospun nanofiber meshes in a wide range of fiber diameters and with many bead formations (Figure 1d). The low collagen concentration in this solvent could be the main reason for bead formations, and the fiber diameters varied along their lengths. The collagen nanofibers in Figure 1a−c had smoother surfaces than the collagen nanofibers in Figure 1d had. In general, the increased collagen concentrations in HFIP, TFE and collagen + PEO mixtures increased the nanofiber diameters. However, the collagen in PBS/EtOH solvent even at relatively high concentrations produced thinner nanofibers. 4.2. Transmission Electron Microscopy (TEM) Results. A TEM bright-field image of electrospun collagen nanofiber samples is given in Figure 2. They did not exhibit the characteristic D-periodicity pattern of native collagen. The collagen concentration and the solvent types determined the electrospun collagen nanofibers̀ diameters. TEM results confirmed the SEM results; electrospinning produces randomly oriented and interconnected nanofibers with varied fiber diameters. We did not include the image of the nanofibers

The CCA algorithm operates in a reciprocal fashion and determines the conformational weights, Cij, and curves, gi(λ). If P is the pure components chosen and Cij, at a given j, is the weight of the ith pure component curve, gi(λ), the measured CD curve ∫ hj (λ) can be fit by a calculated curve, ∫ cj (λ) that has the form p

f jc (λ) =

∑ Cij*gi(λ)

(4)

i=1

For the estimation model CCA set-I, we measured 12 temperature series of CD spectra (at 10, 15, 20, 25, 30, 35, 40, 50, 60, 70, 80, and 90 °C) and the spectra, that would be evaluated, were run altogether. Deconvolution was performed to obtain the theoretical base spectra. For the estimation model CCA set-II, we ran 8 CD spectra. They were (1) native collagen at 10 °C (completely folded structure); (2) collagen at 90 °C (completely unfolded structure); collagen in (3) HFIP, (4) TFE, and (5) HAc/ EtOH solvents; and electrospun collagen nanofibers in (6) HFIP, (7) TFE, and (8) HAc/EtOH solvents. Using these spectra, CCA deconvolution was performed. After running the algorithm program, we obtained two base spectra for CCA set-I and four base spectra for CCA set-II, which could be linearly combined to create the measured experimental spectra. Fourier self-deconvolution method for FT-IR spectra was employed, which was developed by Kaupinnen et al.40 to increase the resolution of the amide I bands. The method provides a way of computationally resolving overlapped lines that cannot be instrumentally resolved due to their intrinsic line width. Any experimental spectrum, E(υ), can be expressed as a convolution of a line shape function, G(υ), and a spectrum, E′(υ), that is ∞

E(ν) = G(ν)*E′(ν) =

∫−∞ G(ν′)E′(ν − ν′) dν′

(5)

where * indicates the deconvolution operation. In order to deconvolute G(υ) from E(υ), one should take the inverse Fourier transform of both sides of eq 5, giving I(x) = ξ −1{G(ν)} I ′(x)

(6)

And the interferogram corresponding to the deconvoluted spectrum is given by I ′(x) =

1 I (x ) ξ −1{G(ν)}

(7) −1

where ξ{ } is the Fourier transform, ξ { } is the inverse Fourier transform, and x has units of centimeters. E′(υ) is then simply obtained by taking the Fourier transform of I′(x). Linear baseline correction, smoothing at 10% and gauss + Lorentz peak type were selected for all FTIR spectra to obtain amide I band components in the frequency range of 1600− 1700 cm−1.

4. RESULTS AND DISCUSSION In this paper, we report the effects of solvent systems on the native collagen structure and of the electrospinning process on the secondary structures of the electrospun nanofibers. We estimated the modifications on secondary structures by using FTIR and CD spectropolarimetry and illuminated the electrospun nanofibers by using SEM and TEM.

Figure 2. TEM images of electrospun collage nanofibers from (a) HFIP, (b) TFE, and (c) HAc/EtOH. 1566

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from PBS/EtOH solvents. Since during preparation for TEM experiments, the fibers folded on each other and were overlaid. However, it did not display any D-periodicity either. 4.3. Fourier Transform Infrared Spectroscopy (FTIR) Results. FTIR was employed to analyze the structure of the collagen solutions, electrospun collagen nanofibers, and dried collagen films. Figure 3 displays the native structure of

Figure 4. FTIR spectra of dried collagen films at room temperature and 70 °C.

Figure 3. FTIR spectrum of native collagen powder.

commercial collagen powder, which was used in our experimental work. The amide I vibration band had a peak position at 1635 cm−1 and a shoulder at 1653 cm−1. The peak position at 1635 cm−1 indicated that our native collagen structure had mainly unfolded or extended structures.41,42 Amide II and amide III adsorption peaks, typical of collagen, had peak positions at 1543 and 1240 cm−1.42 Muyonga et al.43 reportes that the differences in the amide III region of the bone gelatins compared to acid-soluble collagen and skin gelatins are noteworthy, since the intensity of the amide III band has been associated with the triple helical structure. In order to detect the heat induced denaturation process, we analyzed the FTIR spectra of dried collagen films, which were obtained at room temperature and 70 °C. The major differences were observed at amide III band region and the ratio of amide I band to amide II band. The decrease of the amide III band intensity could be attributed to the heat induced unwinding of the collagen triplehelical structure.41,44 Muyonga et al.43 argues that heating the collagen over 50 °C will cause new heat induced bonds in the structure and more intermolecular associations. The denaturation also caused a shift of the amide I band position, and the shoulder at 1653 cm−1 almost disappeared (Figure 4). A broader peak was formed at 1638 cm−1. Figure 5 shows the spectra of electrospun collagen nanofibers, obtained by dissolving in HFIP and TFE for the spinning process. The two spectra were almost identical, with a peak position at 1637 cm−1 for amide I. Amide II peaks were shifted to lower frequency values at 1528 cm−1. Disappearance of the shoulder at 1653 cm−1 and the peak position at 1637 cm−1 suggested that the regenerated electrospun collagen nanofibers were in the denatured state. The shift of Amide II peaks to the lower frequencies was another strong indication of the unfolded structure.45 Amide III bands became almost invisible for both spectra, suggesting an unfolded structure. Since these two fluorinated solvents produced electrospun nanofiber meshes that showed FTIR spectra with identical amide I, II and III band positions, we decided to use the HFIP spectra to represent the samples obtained by fluorinated solvents. All

Figure 5. FTIR spectra of electrospun collagen nanofibers obtained from HFIP and TFE.

other electrospun collagen nanofiber spectra, obtained in other solvents, gave very similar spectral line-shapes. The spectra of these electrospun nanofibers were obtained, using (a) 1:1 (w/ w) collagen and PEO blends dissolved in 0.1 M HCI with the addition of salt (34 mM NaCl); (b) 1:1 (w/w) collagen and PEO blends dissolved in 1:1 (v/v) 0.1 M HCI and EtOH; (c) pure collagen spectra were obtained upon dissolving 15% (w/ v) collagen in 1:1 (v/v) PBS and EtOH; and (d) 1:1 (w/w) collagen and PEO blends dissolved in 1:1 (v/v) 0.05 M HAc and EtOH. All four spectra displayed the amide I band at 1647 cm−1 and amide II band at 1541 cm−1, which are the indication of the folded and unfolded structures̀ existence together in mixed forms (Figure 6). We observed substantial decrease in the amide III band intensity, compared to native collagen spectra. However, it scored higher intensity values than the collagen spectra obtained from HFIP. Figure 7 shows the amide I components of native collagen solution in 0.05 M HAc at 20 and 40 °C, electrospun collagen nanofibers obtained from HFIP, and electrospun collagen/PEO nanofiber blend obtained from HAc/EtOH. Deconvolution results of the amide I bands of native collagen solutions at 20 and 40 °C, electrospun collagen nanofibers obtained from HFIP, and electrospun collagen/PEO nanofiber blends showed that the bands consist 1567

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4.4.2. Native Collagen in Different Solvents. Figure 10 shows the CD spectra of collagen in four different electrospinning solvents. In solution collagen consisted mainly of folded structures in 0.05 M HAc at 15 °C and of unfolded structures in 0.05 M HAc at 90 °C (temperature induced gelation). Native collagen spectra in fluorinated solvents (HFIP and TFE) gave spectra that were more similar to the spectrum of denatured collagen at 90 °C than to the native folded collagen at 15 °C. However, there was a significant difference between the solution spectra in HFIP and TFE at lower wavelengths (between 185 and 190 nm). That was collagen in TFE had a positive ellipticity in this range and a negative band at wavelength of ∼203 nm. One could thus assume a small fraction of an α-helical fold in this solvent, which is known to be a helix-inducing environment for proteins.47 We used three different models to estimate the structural behavior of collagen molecules in the different solvents, as summarized in Figure 11. The calculated PP-II fractions were based on eq 1. CollagenPEO blends in HAc/EtOH solvent scored highest for the folded fraction θ with a value of 0.28, while collagen in HFIP scored 0.07. Since the spectrum in TFE represented itself a principal component, as shown in Figure 10, we could not calculate the folded fraction θ in TFE with this approach. Equation 1, which gave the fraction of folded collagen (θ) simply based on the evaluation at the single wavelength of 221.5 nm, could not be applied in this case. In order to evaluate the full spectral information contained in the CD spectra and to understand the behavior of collagen in the relevant solvents, we determined the PP-II fraction by applying the convex constraint algorithm. Spectral deconvolutions based on reference spectra contained in the CCA set-I and CCA set-II (see Theoretical Section), resulted in two and four base spectra, which could be linearly combined to describe the spectral variation in the measured experimental spectra. According to the estimation based on CCA set-I, the PP-II fraction was 0.015 for the collagen-PEO blend dissolved in EtOH/HAc, 0.051 for native collagen dissolved in HFIP, and 0 for native collagen in TFE. The PP-II fraction from the CCA set-II model gave very similar results for all samples, with a fraction of around 0.001. Thus, we could not detect any significant PP-II fraction in the collagen solutions according to model CCA set-II. 4.4.3. Structural Characterization of Electrospun Collagen Nanofiber Meshes. Each electrospun collagen nanofiber mesh, as obtained by using different solvents, was solubilized in 0.05 M HAc and then characterized by CD. Figure 12 shows the spectra of native collagen in HAc at 15 and 90 °C; and of three different electrospun collagen nanofibers, prepared from different solvents. Native collagen solution at 15 °C displayed the highest amount of PP-II fraction, whereas 90 °C displayed the lowest amount of PP-II, as discussed above. The electrospinning solvents and blending with PEO did not cause any significant structural differences in the obtained electrospun collagen nanofiber spectra, i.e. the three electrospun nanofiber spectra were quite similar. Basically, we observed the same folding behavior, self- and reassembly of the electrospun collagen nanofibers, being independent from the electrospinning solvents. The θ fraction is 0.37 for electrospun nanofibers obtained from HFIP, 0.38 for electrospun nanofibers from TFE, and 0.42 for collagen-PEO blends obtained from EtOH/HAc (using eq 1). CCA set-I and set-II predictions gave less PP-II fractions compared to the single wavelength method, but with very similar results for all three samples. The PP-II fraction derived from CCA set-I gave 0.17

Figure 6. FTIR spectra of electrospun nanofibers (a) a collagen/PEO blend from HCl with salt, (b) a collagen/PEO blend from an HCl/ EtOH mixture, (c) native collagen, dissolved at 15% in PBS and EtOH, and (d) a collagen/PEO blend from HAc/EtOH.

of seven, six, nine, and nine components, respectively. It is reported that protein segments with similar structures do not necessarily show band components with the same frequencies.43 We observed the decrease in amide I band intensity after heating the collagen solution from 20 to 40 °C. The component at 1660.4 cm−1, which is accepted as an indicator of triple helical structure, disappeared when the solution was heated to 40 °C. The denatured collagen structure, which is characterized from a band position centered at 1643 cm−1,45 became the main component for the collagen solution at 40 °C. Amide I bands had peak positions at 1639 cm−1 for electrospun collagen nanofibers and 1649 cm−1 for electrospun collagen/ PEO nanofiber blends. Due to the unfolding of the triple helical structure, we observed that the amide I band component at 1660 cm−1 either completely diminished (collagen solution at 40 °C), or its relative intensity was significantly reduced compared to other components. The remaining amide I components reflected mixed stretching modes. 4.4. Circular Dichroism Results. CD utilizes the differential absorption of left and right handed circular polarized light in an asymmetric environment to assess secondary structure. The amide bonds of a protein in highly ordered conformations such as α-helices, β-sheets, or the PP-II helix exhibit characteristic spectral line shapes due to the specific orientations46 of the chromophores contained in the protein backbone. 4.4.1. Temperature Dependent Folding/Unfolding of Collagen. The CD spectra of native collagen in 0.05 M HAc at different temperatures are presented in Figure 8. We obtained the temperature series by starting from 10 °C and increasing the temperature by 5 °C, up to 40 °C. We then increased the temperature in steps of 10 °C, up to 90 °C. The fraction of PP-II helix and unordered structure of collagen in 0.05 M HAc was calculated using eq 1. Between 10 and 30 °C, the PP-II fractions exceed 90%, and the melting temperature in this solvent was determined to be around 30 °C. Above 35 °C, the structure was dominated by unordered conformations, and collagen finally unfolds completely at 90 °C. Figure 9 shows the calculated fractions of PP-II and unordered collagen conformations. 1568

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Figure 7. FTIR spectra of (a) collagen solution at 20 °C, (b) a collagen solution at 40 °C, (c) collagen nanofibers from HFIP, and (d) collagen/PEO blend from HAc/EtOH.

Figure 9. Calculated fraction of PP-II conformation in thermally denatured native collagen. Figure 8. CD spectra showing the thermal denaturation of 0.1 mg/mL native collagen in 0.05 M HAc.

for electrospun nanofibers from HFIP, 0.17 for electrospun nanofibers from TFE, and 0.19 for electrospun collagen/PEO 1569

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Figure 12. CD spectra of native collagen at room temperature and 90 °C and electrospun nanofiber meshes obtained from different solvents.

Figure 10. CD spectra of native collagen dissolved in different electrospinning solvents (all data apart from the spectra measured in TFE and HFIP have been truncated below 190 nm due to strong background noise).

(Figure 13). The results indicated that the electrospinning process decreased the θ fraction just 8−18% for the first

Figure 11. Folded fraction (PP-II fraction) of native collagen in different solvents (based on the three different evaluation approaches, as described in the Theoretical Section). Figure 13. Folded fraction of electrospun collagen nanofibers and films, obtained from different solvents (based on the three different evaluation approaches, as described in the Theoretical Section).

nanofibers from EtOH/HAc. The PP-II fractions according to CCA set-II resulted in 0.14 for electrospun nanofibers from HFIP, 0.15 from TFE, and 0.16 for electrospun collagen-PEO nanofibers from EtOH/HAc. Within the error limits of the measurements, we could not detect any significant difference of PP-II fraction in the nanofibers obtained from electrospinning of collagen from different solvents. We also compared the CD spectra of collagen film samples with those of electrospun nanofibers to examine the structural effect of the electrospinning process. The films were obtained by casting the collagen solutions from the respective solvents. The collagen films were dissolved in 0.05 M HAc and characterized by CD. We observed a significant increase in the θ fraction of collagen films for all three approaches. The collagen films that were casted from HFIP scored θ fraction values of 0.495, 0.372, and 0.367 for the first, second, and third approach respectively. The collagen films that were casted from TFE scored values of 0.529, 0.423, and 0.423, and collagen PEO blend scored values of 0.57, 0.493, and 0.491, respectively

approach. This showed that the negative effect of spinning process on the fibril structure was not that drastic. However, II. and III. approaches indicated 3−4 folds increase in the θ fractions of collagen films. Very high shear forces and strain rates acting during the electrospinning process, which limit the natural folding of the electrospun collagen nanofibers, were the dominant effect to limit the folded fraction in the electrospun collagen nanofibers. In case of collagen film formation, rapid evaporation first leads to formation of a solid skin, follows by further evaporation of liquid core, leaving voids previously occupied by solvents allowing partial relaxation of the matrix. This induces generation of heterogeneous and porous structures. Therefore the solvent effect is more pronounced in the collagen film formation.20 Moreover, it was important to take into account the whole spectrum range, using II. and III. 1570

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distinct base spectra for the second approach, and by four distinct base spectra for the third approach. The calculated RMSD error shows (0.0138) that the estimated models fit well with the experimental results. Nevertheless, the PP-II fraction was not significantly different among electrospun nanofiber collagen samples. Since the unfolded collagen (gelatin) is susceptible to water and mechanically weak, it is required to cross-link electrospun collagen nanofibers to improve the end use properties. Alternative ways to control the spin-ability and the mechanical properties of the produced electrospun protein nanofibers are to manipulate the protein conformation, control protein aggregation and exchange of intra/intermolecular disulfide bonds.14 These alternative methods and solutions provide the possibility that engineering proteins into biocompatible fibrous structures still could be used in a wide range of biomedical applications such as suturing, wound dressing and wound closure.

approaches, in order to evaluate the secondary structures of the regenerated collagen TEM results displayed no D-periodicity patterns. Therefore, the calculated PP-II fraction by the first approach did not provide information about D-periodicity of the collagen nanofibers. Even electrospun collagen nanofibers from PBS/ EtOH with 42% PP-II fraction did not display any D-bands. It was not sufficient to obtain simply the fraction of folded structure θ from the ellipticity at a single wavelength of 221.5 nm and relatively high folded fraction ratios from the first approach do not represent the real D-periodicity ratios.

5. CONCLUSIONS Electrospinning of collagen is a complex process, and many parameters could affect the final structure of the electrospun nanofibers. In this paper, we investigated the structure of collagen in different electrospinning solvents and in the final electrospun nanofiber mesh products, using FTIR and CD spectroscopy. The experimental results were compared with the structure of native collagen, and data evaluation procedures to determine the PP-II fraction in the electrospun nanofibers were established. Since the ratio between folded and unfolded structures in collagen varies with temperature, we employed this characteristic feature to investigate collagen structures and assess our estimation models. We demonstrated that collagen was completely unfolded in fluorinated solvents, while the native structure was partially preserved in PEO blends due to the HAc/EtOH solvent system. After the spinning process we observed refolding to a certain degree, but the PP-II fraction did not exceed 42% as revealed by the different models and solvent systems discussed here. Thus, we speculated that refolding fraction was limited due to the acting high shear forces during the electrospinning process. We also evaluated the structure of collagen films, which were obtained by casting the same collagen solutions systems. The results were compared with the electrospun nanofiber meshes̀ results. It was shown that even simple casting processes reduced the folded fraction in native structure by 40%. The final PP-II fraction in the collagen films from different solutions were ranged in between 50 and 60%. Therefore it is not just electrospinning process, but also other protein regeneration processes will result with certain amount of unfolding structure in the end product and sacrifice over the native structure. In the case of electrospinning, the high shear forces and the high strain rates (on the order 103 s−1) acting during electrospinning, cause stretching and reorientation of polymer chains (e.g., strain induced crystallization). In parallel, extremely rapid evaporation of solvents affects the macrostructure of electrospun nanofibers.20 Therefore one should expect the formation of supermolecular unfolded structures (i.e., ordered structures).48 The temperature dependent transition from folded to unfolded structure (gelation) occurs around 30 °C in 0.05 M HAc (Figures 8 and 9). We also demonstrated that calculating the folded fraction from a simple evaluation of θ at 221.5 nm in a CD spectrum may not be sufficient to obtain information about the PP-II fraction in electrospun collagen nanofiber samples. Even 42% PP-II fraction from first approach did not show any D-periodicity in TEM images. Therefore, we used two further evaluation procedures based on the CCA algorithm, which included the information from the full spectral range, to simulate each experimental spectrum. The results showed that the spectral variation in the experimental spectra could be described by two

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AUTHOR INFORMATION

Corresponding Author

*E-mail: [email protected]. Phone: +49-721-60823225. Fax: +49-721-608-25546. Notes

The authors declare no competing financial interest.

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ACKNOWLEDGMENTS We would like to thank Dr. Stefan Giselbrecht for his help and discussion in the design and construction of custom-made electrospinning device, the technicians at machine shop facilities at IBG1, and Bianca Posselt IBG2 for excellent technical assistance with the CD measurements.

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REFERENCES

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