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Effect of tissue scaffold topography on protein structure monitored by fluorescence spectroscopy
Journal of Biotechnology 189 (2014) 166–174
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Effect of tissue scaffold topography on protein structure monitored by fluorescence spectroscopy Carla A.M. Portugal a,∗ , Roman Truckenmüller b , Dimitrios Stamatialis c , João G. Crespo a a
REQUIMTE/CQFB, Departamento de Química, FCT-UNL, Campus da Caparica, 2829-516 Caparica, Portugal MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Faculty of Science and Technology, Department of Tissue Regeneration, P.O. Box 217, 7500 AE Enschede, The Netherlands c MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Faculty of Science and Technology, Department of Biomaterials Science & Technology, P.O. Box 217, 7500 AE Enschede, The Netherlands b
a r t i c l e
i n f o
Article history: Received 2 May 2014 Received in revised form 7 August 2014 Accepted 8 September 2014 Available online 19 September 2014 Keywords: Tissue scaffold Protein adhesion Protein conformation Surface topography Fluorescence emission
a b s t r a c t The impact of surface topography on the structure of proteins upon adhesion was assessed through noninvasive fluorescence monitoring. This study aimed at obtaining a better understanding about the role of protein structural status on cell–scaffold interactions. The changes induced upon adsorption of two model proteins with different geometries, trypsin (globular conformation) and fibrinogen (rod-shaped conformation) on poly-l-lactic acid (PLLA) scaffolds with different surface topographies, flat, fibrous and surfaces with aligned nanogrooves, were assessed by fluorescence spectroscopy monitoring, using tryptophan as structural probe. Hence, the maximum emission blue shift and the increase of fluorescence anisotropy observed after adsorption of globular and rod-like shaped proteins on surfaces with parallel nanogrooves were ascribed to more intense protein–surface interactions. Furthermore, the decrease of fluorescence anisotropy observed upon adsorption of proteins to scaffolds with fibrous morphology was more significant for rod-shaped proteins. This effect was associated to the ability of these proteins to adjust to curved surfaces. The additional unfolding of proteins induced upon adsorption on scaffolds with a fibrous morphology may be the reason for better cell attachment there, promoting an easier access of cell receptors to initially hidden protein regions (e.g. RGDS sequence), which are known to have a determinant role in cell attaching processes. © 2014 Elsevier B.V. All rights reserved.
1. Introduction The physiologic behaviour and morphology of cell tissues depend largely on the degree of interaction between cells and cells with the extracellular media during growth (Ge et al., 2013). In supported cell growth, the interaction of cells with the involving media is mediated by the solid supporting matrices, so called tissue scaffolds, with the interaction being largely influenced by the chemical and topographical characteristics of the scaffold surfaces (Jiang and Papoutsakis, 2013). Optimal tissue scaffolds are those that guarantee a stable cell attachment (Zonca et al., 2013), allowing efficient interactions between cell receptors and specific regions of the adhesion proteins (Imen et al., 2008), which pre-adsorb at the scaffold surface, and simultaneously promote a controlled,
∗ Corresponding author. Tel.: +351 21 2948314; fax: +351 21 2948500. E-mail address: [email protected] (C.A.M. Portugal). http://dx.doi.org/10.1016/j.jbiotec.2014.09.009 0168-1656/© 2014 Elsevier B.V. All rights reserved.
non-limited access of the cell to extracellular nutrients and the removal of excreted waste products. The impact exhibited by the topography of scaffold surfaces on cell growth has been reported by several researchers. It is known that the use of cell supports with regular and aligned channels favours cell orientation, a phenomenon often referred to as contact ‘guidance’, leading to the formation of well-organized tissues (Morelli et al., 2010; Papenburg et al., 2010a; Truckenmüller et al., 2012). Moreover, several studies have revealed that the use of solid matrices with aligned patterns is also essential to assure adequate cell maturation and the regulation of diverse cell mechanisms (Chew et al., 2008). The presence of topographical induced cell orientation effects has been supported by confocal laser scanning microscopy studies using specifically labelled proteins (Braber et al., 1998; Morelli et al., 2010) and scanning electron microscopy (SEM) analysis (Zhong et al., 2006; Morelli et al., 2010) which revealed that cellular growth and proliferation occur parallel to the long axes of the surface patterns or, in case of scaffold surfaces with fibrous morphology, along the fibres. In contrast, flat or randomly
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patterned surfaces generate non-oriented cell growth leading to less structured cell tissues. Different hypothesis have been proposed to explain the diverse behaviours observed. Some authors suggested that they may be related to differences in local surface free energy (or surface wettability) that restrains protein and cell attachment to specific surface regions. Others have attributed the preferential cell orientation to the presence of a minimal length for cell attachment of 2 m that forces cells to attach along the longer axes of surface patterns (Izzard and Lochner, 1976). Alternatively, preferential cell attachment may be correlated with the anisotropic geometry of surfaces (Papenburg et al., 2010b) which forces the orientation of cells due to a required energetic balance between surface stresses and shear free planes (Chen, 2008; Guilak et al., 2009). This effect generates mechanotransduction processes which rule primarily cell shape (Guilak et al., 2009; Horbett, 1994) and then the intrinsic cell signalling mechanisms. These mechanisms are driven by the tensional forces between extracellular matrix (ECM) and cells upon cell attachment to scaffolds, which subsequently regulate cell proliferation and differentiation paths. Also, a recent study from Fujita et al. (2009) revealed that cell protrusions which expand perpendicularly to the nanogroove direction retract more quickly than those that move parallel to it. This effect led these authors to conclude that the cell retraction process may contribute to the cell elongation and alignment in nanogrooved surfaces. The relationship between surface topography and cell growth profile is still unclear; however, it has also been hypothesized that this cell contact guiding effect exerted by specific surface topographies may be directed by the structural status of the different adhesion proteins (e.g. fibrinogen, fibronectin, vibronectin, etc.) that primarily adsorb at the scaffold surface during the growth process. Based on this effect, it is reasonable to speculate that the efficiency of cell anchorage to scaffolds is intimately related to the mode how surface impacts on the structure and molecular flexibility of the adhesion proteins since it may influence the accessibility of cell receptors to protein regions (Imen et al., 2009), such as arginine-glycine-), such as arginine–glycine–aspartic acid (RGD) sequences, which are known to be determinant in cell attaching processes (Horbett, 1994). The ability and the way proteins adsorb at a given surface depend on their conformation and also the structural status of an adsorbed protein relies on the surface topography. This relationship was clearly demonstrated in the work developed by Roach et al. (2006) which highlights the contrasting effects induced on globular and rod-like proteins upon contacting surfaces with different patterns. These authors have shown that the contact of a rod-like protein, such as fibrinogen, to silica surfaces with accentuated curvature results in a significant change of its secondary structure, characterized by a loss of the ␣-helix structure in benefit of random coil chain segments, which is related to its capacity to fit the surface. Contrastingly, and due to their inability to adapt to surfaces with high curvatures, globular proteins exhibit an opposite structural behaviour when adsorbed at an identical surface, registering a decrease of the ␣-helix content with the decrease of surface curvature. Studies of the impact of surface topography in cell growth have shown that cell spreading, adhesion and differentiation also depend on the dimensions of the surface topographical features (Dalby et al., 2002, 2003), and fibre diameter (Elias et al., 2002) when fibrous scaffolds are used. However, a clear relationship between structural changes induced upon attachment of plasma proteins to the scaffold surface and the degree of cell orientation achieved with different surface topographies has not been established so far. Fluorescence spectroscopy is a highly sensitive technique capable of providing simultaneously valuable information about protein structure and mobility, which has not been sufficiently explored so
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far for monitoring of cell growth at tissue scaffolds. Therefore, in this work, a fluorescence monitoring methodology combining data from steady-state fluorescence and fluorescence anisotropy is used to assess the impact of surface topography on the structure and molecular flexibility of proteins. Protein structural changes can be inferred from the fluorescence emission of a protein intrinsic fluorescence probe, tryptophan. The use of tryptophan as a structural reporter excludes the need of protein labelling with an external chromophore. Based on the high sensitivity of the fluorescence emission of tryptophan to changes of the physicochemical properties in its surrounding environment, fluorescence emission allows for in situ monitoring of the changes of protein structure (Portugal et al., 2006, 2007; Guedidi et al., 2012) and the mode that each geometrically different protein interacts to the surface. Fluorescence anisotropy measurements allow for eliciting information about the influence of the different surface topographies on molecular flexibility. The present study was conducted using model proteins with opposite conformational geometries: a globular protein, trypsin (largely used in adsorption studies, presenting a significant fluorescence emission) and a rod-like protein, fibrinogen, which integrates the group of adhesion proteins directly involved in the cell–tissue scaffold interaction process. The two proteins were selected in order to mimic the conformational diversity (globular and rod-like proteins) of the proteins involved in the cell attachment process. This work aims at showing the potential impact of the structural status of proteins induced by scaffold surface topography on cell attachment using a non-invasive fluorescence monitoring approach. To achieve this goal, a correlation between the topographically induced structural changes of the adhesion proteins and the growth of C2C12 mouse pre-myoblast cells at these scaffolds was established. As described in previous papers from some of the authors of the present work (Haneveld, 2006; Papenburg et al., 2010a; Bettahalli et al., 2012) the use of fibrous scaffolds improves cell–cell interactions while allowing a good spreading of the C2C12 cells. 2. Experimental 2.1. Preparation of protein solutions Protein solutions of trypsin from bovine pancreas (EC. 3.4.21.4, Ref. T9935 from Sigma–Aldrich), a globular protein with MW of 20.3 kDa (Fig. 1(a)), and fibrinogen from bovine plasma (EC 232598-6, Ref. F8630, from Sigma–Aldrich), a rod-like protein with MW of 340 kDa (Fig. 1(b)), with respective concentrations of 2 g/L and 0.7 g/L were prepared in 0.1 M Trizma® buffer at pH 8 containing 0.02 M of CaCl2 , added to prevent trypsin autolysis. Trizma buffer was prepared using Trizma® hydrochloride (Sigma Aldrich) and Trizma® base (Sigma Aldrich). To avoid protein structural changes due to the physicochemical conditions, both protein solutions were prepared using the solvent media with identical characteristics. Protein concentration was selected in order to be in the range of the typical cell culture media protein content and to assure the detection of fluorescence emission from the protein molecules adsorbed at the scaffold surfaces. Differences in concentrations of both protein solutions were due to the low solubility of fibrinogen in the Trizma buffer used. 2.2. Protein adsorption on PLLA scaffolds with distinct topographies Trypsin and fibrinogen were separately adsorbed at poly-llactic acid (PLLA) scaffolds with different surface patterns: flat, aligned nanogroove pattern and random fibrous surfaces. Surfaces with aligned nano-patterning were obtained by using nano imprint
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Fig. 1. Schematic representations of (a) the three-dimensional globular structure of trypsin from bovine pancreas and (b) the three-dimensional rod-like structure of bovine fibrinogen (adapted from Brown et al., 2000) highlighting the location of tryptophans in red colour (structural probe). Both schematic representations were obtained and adapted from the Protein Data Bank at http://www.rcsb.org/pdb/home/home.do.
Fig. 2. Thermal NIL process for patterning a PLLA film with the following process steps: (a) insertion of the PLLA film, (b) evacuating the space between the film and an imprinting stamp from silicon, heating up the stamp and with it the film, and applying a gas pressure to imprint the stamp into the film, and (c) cooling down the stamp and the film, releasing the gas pressure, removing the vacuum and demoulding the imprinted film by peeling it off from the stamp ((1) sealing foil separating the vacuum section of the nano imprint lithography machine from its high-pressure section, (2) PLLA film to be imprinted, (3) imprint stamp from silicon (blue and red corresponds to cold and heated), (4) compressed gas. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
lithography. The nanogroove patterns were imprinted (Fig. 2) in a FDA (U.S. Food and Drug Administration) compliant, 35 m thin, biaxially oriented blown film (nominal thickness: 30 m) from a blend of PLA (Plastic Suppliers/Sidaplax; EarthFirst Packaging Film, PF). The film had a glass transition temperature of 58 ◦ C, a melting temperature of approximately 160 ◦ C and the surface tension of the untreated surface was 38 mN m−1 . Imprinting was carried out in a machine for hot embossing nano imprint lithography (Obducat, Malmö, Sweden; NIL 6 ). The imprinting stamp from silicon carried a periodical pattern of parallel ridges with a width of 150 nm, a periodicity of 500 nm and a height of 100 nm. The stamp was fabricated by laser interference lithography and wet chemical etching (Haneveld, 2006). To facilitate demoulding, the stamp was functionalized with a perfluorodecyltrichlorosilane (FDTS; ABCR, Karlsruhe, Germany) anti-stiction layer by vapour deposition (Haneveld, 2006). In particular, the surface with aligned nano-pattern is composed of well-oriented channels with 150 nm width and 100 nm depth and an inter-channel spacing of 350 nm (Fig. 3(a) and (b)).
Fibrous scaffolds were prepared by dissolution of 12 wt% PLLA solution (PLLA mol wt. 1.6 × 105 g/mol) in 1,3-dioxane which was then spun using an in-house-built electrospinning set-up. SEM images were made to determine the surface topography and fibre diameter (Fig. 3(c) and (d)). For this, the sheets were sputter-coated with gold. Analysis of SEM results showed that the fibrous PLLA scaffold is composed by fibres with an average diameter of 2.5 m. The fibrous PLLA scaffolds obtained had a thickness of 200 m and a porosity of 71 ± 3% (Bettahalli et al., 2012). The adsorption of the proteins to the various scaffolds was induced by immersion of the scaffolds, for 3 h in 5 mL of trypsin or fibrinogen solutions under permanent external soft stirring, using a multi-mixer system (mistral, Lab-line Instruments). Upon this procedure, initial protein solutions (i.e. before the adsorption process) and the scaffold surfaces with adsorbed proteins were inspected using the fluorescence methodology described below to evaluate the influence of the surface topography on the structural status of proteins. The amount of the adsorbed proteins at the different surfaces was determined by inductively coupled plasma (ICP-AES) analysis using a Jobin Yvon-Ultima ICP spectrometer (Edison, NJ, USA). This analysis allow for spectroscopic quantification of proteins in samples acquired before and after adsorption based in the sulphur content, using its characteristic emission line at 180.676 nm. 2.3. Monitoring structural changes of the proteins induced upon interaction with the scaffold surfaces The structural changes underwent by the two proteins upon adsorption at the scaffold surfaces were assessed by using a fluorescence methodology, which combined the information provided by steady-state fluorescence emission and steady-state fluorescence anisotropy. 2.3.1. Steady-state fluorescence analysis The fluorescence scans were acquired at an excitation wavelength, exc , of 290 nm and at an emission wavelength, em , ranging from 300 to 550 nm, for the solutions of proteins before the adsorption and for proteins adsorbed at each scaffold surface. The fluorescence spectra were acquired using a SPEX Fluorolog spectrofluorimeter, equipped with excitation and emission polarizers positioned at the magic angle (54.7◦ ) and using excitation and emission slits of 5 nm bandpass. Steady-state fluorescence analysis enables us at inferring about changes of the three-dimensional structure of proteins induced upon adsorption at the different surfaces. The fluorescence emission obtained at exc of 290 nm is essentially due to the contribution of the indole group of the tryptophan residues present at each protein chain. Changes of the fluorescence characteristics of the
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Fig. 3. SEM images of the PLLA scaffolds with nanogrooved surface produced by nano imprint lithography obtained with magnifications of: (a) x 47.15K and (b) x 17.06K 47.15× K and (b) 17.06× K; electrospun PLLA scaffolds obtained with magnifications of (c) x 75 and (d) x 500, prepared by dissolution of 12 wt% PLLA solution in 1,3-dioxane.
tryptophan residues may result from a different proximity to quenchers present in the protein chain, such as aspartate, disulfidric and hydrogen bridges (Chen and Barkley, 1998), differences on the exposure degree of these fluorophores to the solvent, e.g. as result of the protein folding/unfolding processes, leading to displacements of the maximum emission. Shifts of the maximum emission to higher emission wavelengths (maximum emission red shift) may be ascribed to a higher exposure of the tryptophan residues to an environment with higher dielectric constant (or higher polarity) (Portugal et al., 2007; Faria et al., 2004). In contrast, a decrease of the dielectric constant in the microenvironment surrounding the fluorophores, resultant from a protein folding process, may lead to the dislocation of the maximum emission to lower emission wavelengths (maximum emission blue shift) followed by an increase of the fluorescence intensity (Portugal et al., 2007).
protein fluorescence spectra, leading to an increase of the fluorescence emission of the system and to an apparent dislocation of the maximum emission, which could prevent an accurate perception of the fluorescence emission changes induced by the adsorption process. To overcome this limitation, the fluorescence emission spectra of the adsorbed proteins were elicited by subtracting the fluorescence emission spectra obtained for scaffold surfaces before protein adsorption from the fluorescence emission spectra obtained for scaffold surface with the adsorbed protein, resulting in the dashdot line spectra shown in Fig. 4 (a). Fluorescence anisotropy from the adsorbed proteins, shown in Fig. 4 (b) was also determined according to the methodology described by Guedidi et al. (2012).
2.3.2. Steady-state fluorescence anisotropy analysis Fluorescence anisotropy provides information regarding the mobility of the fluorophores present in the protein. Changes of the mobility of fluorophores, either that concerning to tryptrophans intrinsic rotation ability or that due to protein mobility, can be correlated with the impact of protein interactions with its surrounding environment (i.e. folding/unfolding processes or protein–surface interactions) on their structure and molecular flexibility. The fluorescence anisotropy, r, was acquired using the spectrofluorimeter referred above using the same slits and spectral ranges, by varying the position of excitation and emission polarizers. Determination of fluorescence anisotropy, r, values is described in detail in previous works (Portugal et al., 2006, 2007; Guedidi et al., 2012).
A fluorescence monitoring methodology combining steadystate fluorescence emission and fluorescence anisotropy analysis was used in order to gather a better understanding of the role of the structural status of the adhesion proteins in the cell attachment process. In this work, fluorescence monitoring was applied to probe the impact of PLLA surfaces with flat, aligned nano-patterning and random fibrous pattern designs on the structure of two model proteins with distinct conformational geometries: trypsin, with a globular geometry (Fig. 1(a)) and fibrinogen, a rod-shaped protein (Fig. 1(b)), which were selected to mimic the diversity of proteins’ conformational geometries present in cell media, either they are directly involved in cellular adhesion or not. The number of tryptophans per protein chain was also taken as a model protein selection criterion, since it dictates the success of fluorescence monitoring. Fibrinogen is an adhesion protein, used as mediator of cell adsorption at tissue scaffolds, with a high number of tryptophans. Although, trypsin is not a cell binding protein, its tryptophan content facilitates the detection of this biomolecule in conditions where the perception of tryptophan signal is perturbed by external elements in proteins’ vicinity. This aspect is critical when monitoring the fluorescence of adsorbed proteins, in case
2.3.3. Determination of the fluorescence emission properties of the adsorbed proteins As illustrated in Fig. 4, the fluorescence emission of PLLA scaffold surfaces is characterized by the presence of a large emission band of low intensity with a maximum emission between 390 and 450 nm (depending on the scaffold surface topography). The emission band of the PLLA scaffolds superimposes at the red edge of the
3. Results and discussion
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Fig. 4. (a) Fluorescence emission spectra and (b) fluorescence anisotropy spectra obtained at exc of 290 nm for trypsin in solution before adsorption (1), PLLA fibrous surface (2), PLLA fibrous surface with adsorbed trypsin (3) and trypsin adsorbed at fibrous PLLA surface upon subtraction of the contribution of the PLLA surface (4).
of the contribution of the intrinsic fluorescence of the surface (e.g. tissue scaffold) at the protein emission region (Guedidi et al., 2012). The amount of trypsin and fibrinogen adsorbed to the surfaces with different topographies was determined by ICP-AES analysis. The results obtained revealed that the amount of trypsin adsorbed at flat and fibrous surfaces were 2.12 × 10−9 mol/cm2 (0.043 mg/cm2 ) and 3.99 × 10−11 mol/cm2 (0.81 g/cm2 ), respectively. When compared in molar terms, it was observed that the amount of fibrinogen adsorbed at flat and fibrous surfaces of 4.51 × 10−10 mol/cm2 (0.153 mg/cm2 ) and 7.22 × 10−12 mol/cm2 (2.45 g/cm2 ), respectively, was smaller, which is attributable to the significantly different protein sizes. 3.1. Fluorescence monitoring of the influence of surface topography on the structural behaviour of globular proteins Fig. 5(a) shows the fluorescence spectra obtained, at exc of 290 nm, for the globular protein, trypsin, before and after the adsorption at scaffolds with distinct surface topographies. For the easiness of the analysis of the results and for cancelling the dependence of fluorescence emission on the protein concentration, fluorescence changes were analyzed based on the comparison of normalized fluorescence emission spectra. As can be observed, the adsorption of trypsin at flat surfaces causes a clear displacement of the maximum emission to higher wavelengths, a red shift, with an amplitude of ∼6 nm, followed by the enlargement of the emission bandwidth (Fig. 5(a)). A maximum emission red shift of lower amplitude (∼2 nm) is also visible after the adsorption of this protein at surfaces with the fibrous morphology (Fig. 5(a)). The fluorescence emission changes detected reflect the increase of the dielectric constant at the local microenvironment of tryptophans due to their higher exposure to the external aqueous phase. Hence, this behaviour clearly indicates that the adsorption of trypsin at PLLA flat and fibrous surfaces generates the presence of molecular unfolding processes which, based on the amplitude of the fluorescence emission changes, seems to be more significant upon contact with flat surfaces. Trypsin exhibited a distinct structural behaviour upon adsorption at PLLA surface with an aligned nano-patterning. In this case, the adsorption is followed by the dislocation of the maximum emission to lower wavelengths, a blue shift (Fig. 5(a)). The blue shift observed is attributable to the increase of the hydrophobic character of the environment in the vicinity of the tryptophans. This effect may be due to a higher contact of the tryptophans with the
scaffold surface, which has a dielectric constant lower than water or to protein structural folding processes induced by their contact with the surface with such topography. The information about the structural changes of trypsin induced by the adsorption process was complemented by the analysis of the rotational ability of tryptophans provided by steady-state fluorescence anisotropy. From the anisotropy spectra shown in Fig. 5(b), it is possible to conclude that the rotational ability of tryptophans is also influenced by the surface topography. The attachment of trypsin to flat surfaces and surfaces with aligned patterning lead to an increase of its fluorescence anisotropy, expressing a hindered rotational ability of tryptophan residues upon protein adsorption, compared to that observed for trypsin in solution. There is an obvious agreement between the increase of fluorescence anisotropy and the maximum emission blue shift observed upon adsorption at the surface with aligned pattern. Indeed, the molecular processes which explain the presence of a maximum emission blue shift, i.e. an additional molecular folding or an increase of the contact degree of tryptophans towards the surface, may both justify the restricted tryptophan mobility. Contrastingly, the maximum emission red shift, i.e. trypsin unfolding, followed by a reduction of tryptophans’ mobility, as monitored upon trypsin adsorption at the flat surface, seems contradictory at a first glance. However, both effects may result from a combination of different molecular events inherent to the adsorption process, which generate apparently opposite alterations on the fluorescence emission and anisotropy spectra. The total mobility of tryptophans results from the sum of different tryptophan rotational components: the intrinsic rotational ability of tryptophans, which is hampered by the protein’s threedimensional structure; and the mobility of the protein as a whole, which contributes to an additional mobility of the tryptophan residues. Based on this explanation, the loss of tryptophans’ mobility observed upon attachment at the flat surface may be mainly attributed to a decrease of the whole protein mobility after attaching to the scaffold surface. However, the tryptophan imprisonment, which should result in a maximum emission blue shift, was probably not enough to surpass the protein maximum emission red shift induced by unfolding. An opposite structural behaviour was observed upon adsorption of trypsin at random fibrous surfaces. The decrease of fluorescence anisotropy (Fig. 5(b)) and the small maximum emission red shift (Fig. 5(a)) observed in this case supports the presence of a slight trypsin unfolding, with the consequent increase of tryptophans
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Fig. 5. Fluorescence emission (a) and anisotropy spectra (b) of trypsin in solution (1), trypsin adsorbed at PLLA flat surface (2), trypsin adsorbed at PLLA fibrous surface (3) and trypsin adsorbed at PLLA aligned nano-patterned surface (4).
mobility. Together, these results suggest the absence of significant structural changes upon trypsin attachment to fibrous surfaces. Still, fluorescence data allowed us to speculate that the trypsin structural changes are potentially followed by an incomplete or less efficient attachment of trypsin at the fibrous scaffold. The fluorescence emission changes observed for the trypsin adsorbed at flat and fibrous surfaces are in concordance with the results reported in the literature. Our results show that the adhesion of globular proteins to flat or fibrous surfaces is clearly preceded by an unfolding step, which is compatible with the increase of random-coil character of protein structure (Roach et al., 2006). Moreover, the proportion of random coil segments increases with the increase of the radius of the surface curvature, which is also in agreement with the highly intense unfolding achieved upon adsorption of globular protein at flat surfaces. Additionally, it was observed that the adsorption of globular proteins at flat surfaces is followed by a structural spreading of the adsorbed protein (Agnihotri and Siedlecki, 2004; Roach et al., 2006) whereas minor alterations are expected due to their adsorption at surfaces with high curvatures (such is the case for fibrous scaffolds). The molecular spreading of globular proteins at flat surfaces results from the presence of a higher contacting surface area available, and explains the trypsin unfolding as well as the improved contact of the tryptophans with the flat surface. The ability of trypsin to spread at surfaces with higher curvatures is limited due to the reduced contacting area available. Although in a lower extent, trypsin unfolding still occurs, resulting in minor maximum emission red shift amplitudes (as shown in Fig. 5 (a)), contributing to the increase tryptophan mobility. The increase of tryptophans rotational ability also evidences the presence of random coil protein regions, which are unable to bind at the scaffold surface. 3.2. Fluorescence monitoring of the effect of topography on the structure of rod-like proteins A similar study was conducted to elicit information about the influence of scaffold surfaces with different topographies on the conformation of a rod-like protein, fibrinogen (Fig. 6). Analysis of the fluorescence emission spectra obtained for the fibrinogen before and after adsorption at surfaces with flat, aligned nano-channels and fibrous morphology (Fig. 6(a)) reveals that the adsorption of fibrinogen does not cause displacements of its maximum emission as intense as those observed after adsorption of
trypsin at identical surfaces. Only a subtle blue shift, with no more than to 2 nm, followed by the enlargement of the emission band, was observed after the adsorption of this protein at the surface with aligned nanogrooves. Complementary fluorescence anisotropy results, depicted in Fig. 6 (b), show that despite the absence of significant fluorescence emission changes, the adsorption of fibrinogen at scaffold surfaces is followed by the alteration of fluorescence anisotropy. Indeed, the adsorption of fibrinogen at flat surfaces leads to an increase of the fluorescence anisotropy, relatively to the anisotropy of fibrinogen in solution. As referred previously, the increase of fluorescence anisotropy corresponds to a decrease of tryptophans mobility, attributable either to the contact of tryptophans to the surface or to an additional folding of the protein regions containing the tryptophans. In this case, tryptophan–surface interactions are facilitated by the elongated conformation of fibrinogen, which allows for an improved contact of amino acids to the flat surface without involving a significant change of the protein structure. The higher increase of fluorescence anisotropy and the maximum emission blue shift observed for fibrinogen adsorbed at the nanogrooved surface suggest a more intense contact of tryptophans to the nanogrooved surface, which may be related to a highly stable protein adhesion and to a good cell attachment and growth observed at these scaffolds (Papenburg et al., 2007; Truckenmüller et al., 2012). Nevertheless, in this case, the anisotropy reached values higher than 0.4, the maximum limit theoretically possible for fluorescence anisotropy (Valeur, 2002). The high fluorescence anisotropy might be related to the presence of light scattering and reflection phenomena by the surface, disabling us from taking clear conclusions from these results. The adsorption of fibrinogen at fibrous surfaces generates a decrease of the fluorescence anisotropy (Fig. 6(b)), a similar behaviour to that found upon adsorption of trypsin (globular protein) at PLLA surfaces with the same morphology (Fig. 5(b)). Identically to that discussed for trypsin, the decrease of the fluorescence anisotropy of fibrinogen adsorbed at fibrous surface expresses higher tryptophan mobility, attributable to a higher fibrinogen unfolding degree upon adsorption. In the native conformation of fibrinogen, tryptophans are already highly exposed to the aqueous media, which is confirmed by the average lifetime of tryptophans in free fibrinogen, which is equal to 2.71 ns (result obtained by time-resolved fluorescence),
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Fig. 6. (a) Fluorescence emission and (b) fluorescence anisotropy spectra obtained at exc of 290 nm for fibrinogen in solution (1), adsorbed at flat PLLA surface (2), adsorbed at fibrous PLLA surface (3) and adsorbed at aligned nano-patterned surface (4).
that corresponds to a lifetime close to that expected for tryptophans totally exposed to the aqueous environment, which is equal to ∼3 ns (Petrich et al., 1983). As discussed above, the absence of fluorescence emission changes suggests that the adsorption of fibrinogen does not result in a significant increase of the exposure degree of tryptophans to the external aqueous environment. Hence, the increase of the protein unfolding degree may be explained based on the ability of rod-like proteins, such as fibrinogen, to mould to surface curvatures (Roach et al., 2006). Although the curvature radius of the surface used in this work was not significantly accentuated regarding the protein size, it may be enough to force the amino acids to pull apart upon fibrinogen adsorption. Also it may be speculated that the adsorption is associated to an additional unfolding of the nodular regions of fibrinogen, which contains most of the tryptophans present in the fibrinogen chain (Fig. 1(c)). This effect would justify the higher mobility of tryptophans and a better accessibility to protein regions with a relevant role in the cell adhesion, which were initially hidden within the protein structural core. According to literature, RGD sequences or the dodecapeptide at the carboxyl terminus of the ␥-chain were found to be one of the preferential cell binding sites at fibrinogen (Horbett, 1994). These segments are located in different fibrinogen domains, in particular they are placed within D domain ␥ 400–411, C domain at A␣ 98–101 and at the coiled-coil regions A␣ 255–258, which in turn are placed at the extreme nodular regions of fibrinogen where the accessibility of the amino acids in the native protein may be limited (Brown et al., 2000; Mosesson et al., 2001). The partial unfolding of proteins, specifically fibrinogen, upon adsorption promotes the exposure of these chain segments improving cell–protein interactions, and would justify the contribution of molecular unfolding to the enhancement of cell adhesion performance, as observed at fibrous scaffolds. In fact, recent studies performed by some of the authors of the present work showed that there was higher cell–cell contact on the fibrous PLLA sheets compared to the PLLA flat sheets and tissue culture polystyrene (TCPS) (Papenburg et al., 2010a). This is in agreement with literature, where it was reported that in various cell types, integrins associated with cell–cell as well as cell–ECM contact were up-regulated on nano-fibrous scaffolds compared to flat surfaces as response to increased protein adsorption (Kumar, 2006). Also, SEM images supported this observation as the C2C12 cells spread well on the fibrous PLLA sheets and show good connection between the cells’ filopodia as well as between the cells and the nano-fibrous surface. The DNA assay data also
indicated slightly increased cell proliferation on the fibrous PLLA compared to the flat PLLA, although only significant at day 4 for the higher cell density (Kumar, 2006). Moreover, as shown by Pagliara et al. (2010), electrospinning and nano imprint lithography induces molecular orientation within polymer structure, resulting in the increase of light polarization and consequently an increase of fluorescence anisotropy. Therefore, the adsorption of proteins to structurally aligned or random surfaces may promote preferential or disordered orientations of the transition dipole moments of tryptophans towards the excitation light, also contributing to the increase or decrease of fluorescence anisotropy, respectively. This behaviour suggests the alignment of protein molecules at the nanogrooved surface, which corroborates the improved cell orientation of tissues obtained using such scaffold surfaces (Truckenmüller et al., 2012; Papenburg et al., 2007). 4. Concluding remarks A non-invasive fluorescence monitoring methodology was used to access the impact of different surface topographies on the structural status of proteins, aiming at establishing a correlation between the structural behaviour of adhesion proteins adsorbed at the scaffold surfaces and the cell growth process. Fluorescence analysis simultaneously provided information about the folding/unfolding status of the adsorbed proteins as well as their molecular mobility level. Together, this information allowed for inferring about protein binding intensity, accessibility to hindered protein regions and the presence of anisotropic structural features which may support the presence of topographically induced oriented cell attachment. The fluorescence results obtained confirmed that the structural behaviour of proteins when interacting with surfaces depends not only on the surface pattern design but also on the protein conformational geometry (Roach et al., 2006). The latter dictates the ability of proteins to fit the surface and, together with the surface chemistry, it rules the impact of the adsorption process on the protein conformation. As expressed by the increase of fluorescence anisotropy, the adsorption of globular or rod-shaped proteins at flat surfaces and surfaces with aligned nano-patterning leads to a high restriction of protein mobility, comparatively to that produced by the adsorption at fibrous surfaces. This structural behaviour stands for an intense surface–protein interaction, which may justify a stable protein
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attachment. The increase of fluorescence anisotropy may also comprise the contribution of a preferential alignment of the adsorbed proteins driven by the surface channel pattern. This behaviour is corroborated by the alignment of FITC labelled proteins adsorbed at grooved surfaces monitored by Braber et al. (1998) using confocal laser scanning microscopy. In contrast, the adsorption of both globular and rod-shaped proteins to random fibrous scaffolds leads to a decrease of fluorescence anisotropy to values even smaller than that obtained for proteins in solution. These results may be interpreted by an amplification of the rotational ability of tryptophans, which is higher upon adsorption of rod-shaped than globular proteins, and may reflect the ability of these molecules to mould to curved surfaces (Roach et al., 2006). Such structural alterations offer an improved exposure of the amino acids to the external media and support the contact of important protein chain segments, e.g. RGDs sequences (Horbett, 1994) to the receptors of membrane cell proteins, explaining the better cell growth efficiency obtained when fibrous scaffolds are used as cellular supports (Papenburg et al., 2010a; Woo et al., 2003). Also, it may be rationalized as the presence of non-oriented adsorbed proteins at the surface of randomly aligned fibrous scaffold. Hence, the in situ fluorescence monitoring approach used in this work shows clearly that cell attachment efficiency is ruled by a combined action of two effects: the preferential adsorption and alignment of proteins according to specific surface topographical features and the topographically induced protein structural status. Additionally, fluorescence monitoring has shown to be a powerful tool that can be used as a single technique to guide the development of tissue scaffolds with improved designs and also the configuration of new engineered adhesion peptides (Sprio et al., 2011), contributing in this mode to the improvement of cell attachment processes. Acknowledgements The authors would like to acknowledge to Prof. Fernando Pina, Prof. Maria João Melo and Prof. João Carlos Lima the access to the spectrofluorimeter. The authors also acknowledge the funding from Fundac¸ão para a Ciência e a Tecnologia (FCT-MEC), Portugal, through the project PEstC/EQB/LA0006/2011. References Agnihotri, A., Siedlecki, C., 2004. Time-dependent conformational changes in fibrinogen measured by atomic force microscopy. Langmuir 20, 8846–8852, http://dx.doi.org/10.1021/la049239+. Bettahalli, N.M.S., Groen, N., Steg, H., Unadkat, H., de Boer, J., van Blitterswijk, C.A., Wessling, M., Stamatialis, D., 2012. Development of multilayer constructs for tissue engineering. J. Tissue Eng. Regen. Med., http://dx.doi. org/10.1002/term.1504. Braber, E.T., de Ruijter, J.E., Ginsel, L.A., von Recum, A.F., Jansen, J.A., 1998. Orientation of ECM protein deposition, fibroblast cytoskeleton, and attachment complex components on silicone microgrooved surfaces. J. Biomed. Mater. Res. 40, 291–300, http://dx.doi.org/10.1002/(SICI)1097-4636(199805)40: 2<291::AID-JBM14>3.0.CO;2-P. Brown, J.H., Volkmann, N., Jun, G., Henschen-Edman, A.H., Cohen, C., 2000. The crystal structure of modified bovine fibrinogen. Proc. Natl. Acad. Sci. U.S.A. 97, 85–90, http://dx.doi.org/10.1073/pnas.97.1.85. Chen, C.S., 2008. Mechanotransduction – a field pulling together? J. Cell Sci. 121, 3285–3292, http://dx.doi.org/10.1242/jcs.023507. Chen, Y., Barkley, M.D., 1998. Toward understanding tryptophan fluorescence in proteins. Biochemistry 37, 9976–9982, http://dx.doi.org/10.1021/bi980274n. Chew, S.Y., Mi, R., Hoke, A., Leong, K.W., 2008. The effect of the alignment of electrospun fibrous scaffolds on Schwann cell maturation. Biomaterials 29, 653–661, http://dx.doi.org/10.1016/j.biomaterials.2007.10.025. Dalby, M.J., Riehle, M.-O., Johnstone, H.J.H., Affrossman, S., Curtis, A.S.G., 2002. Polymer demixed nanotopography control of fibroblast spreading and proliferation. Tissue Eng. 8, 1099–1108, http://dx.doi.org/10.1089/107632702320934191. Dalby, M.J., Riehle, M.-O., Johnstone, H.J.H., Affrossman, S., Curtis, A.S.G., 2003. Non-adhesive nanotopography: fibroblast response to poly(n-butyl methacrylate)-poly(styrene) demixed surface features. J. Biomed. Mater. Res. 67A, 1025–1032, http://dx.doi.org/10.1002/jbm.a.10139.
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Journal of Biotechnology journal homepage: www.elsevier.com/locate/jbiotec
Effect of tissue scaffold topography on protein structure monitored by fluorescence spectroscopy Carla A.M. Portugal a,∗ , Roman Truckenmüller b , Dimitrios Stamatialis c , João G. Crespo a a
REQUIMTE/CQFB, Departamento de Química, FCT-UNL, Campus da Caparica, 2829-516 Caparica, Portugal MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Faculty of Science and Technology, Department of Tissue Regeneration, P.O. Box 217, 7500 AE Enschede, The Netherlands c MIRA Institute for Biomedical Technology and Technical Medicine, University of Twente, Faculty of Science and Technology, Department of Biomaterials Science & Technology, P.O. Box 217, 7500 AE Enschede, The Netherlands b
a r t i c l e
i n f o
Article history: Received 2 May 2014 Received in revised form 7 August 2014 Accepted 8 September 2014 Available online 19 September 2014 Keywords: Tissue scaffold Protein adhesion Protein conformation Surface topography Fluorescence emission
a b s t r a c t The impact of surface topography on the structure of proteins upon adhesion was assessed through noninvasive fluorescence monitoring. This study aimed at obtaining a better understanding about the role of protein structural status on cell–scaffold interactions. The changes induced upon adsorption of two model proteins with different geometries, trypsin (globular conformation) and fibrinogen (rod-shaped conformation) on poly-l-lactic acid (PLLA) scaffolds with different surface topographies, flat, fibrous and surfaces with aligned nanogrooves, were assessed by fluorescence spectroscopy monitoring, using tryptophan as structural probe. Hence, the maximum emission blue shift and the increase of fluorescence anisotropy observed after adsorption of globular and rod-like shaped proteins on surfaces with parallel nanogrooves were ascribed to more intense protein–surface interactions. Furthermore, the decrease of fluorescence anisotropy observed upon adsorption of proteins to scaffolds with fibrous morphology was more significant for rod-shaped proteins. This effect was associated to the ability of these proteins to adjust to curved surfaces. The additional unfolding of proteins induced upon adsorption on scaffolds with a fibrous morphology may be the reason for better cell attachment there, promoting an easier access of cell receptors to initially hidden protein regions (e.g. RGDS sequence), which are known to have a determinant role in cell attaching processes. © 2014 Elsevier B.V. All rights reserved.
1. Introduction The physiologic behaviour and morphology of cell tissues depend largely on the degree of interaction between cells and cells with the extracellular media during growth (Ge et al., 2013). In supported cell growth, the interaction of cells with the involving media is mediated by the solid supporting matrices, so called tissue scaffolds, with the interaction being largely influenced by the chemical and topographical characteristics of the scaffold surfaces (Jiang and Papoutsakis, 2013). Optimal tissue scaffolds are those that guarantee a stable cell attachment (Zonca et al., 2013), allowing efficient interactions between cell receptors and specific regions of the adhesion proteins (Imen et al., 2008), which pre-adsorb at the scaffold surface, and simultaneously promote a controlled,
∗ Corresponding author. Tel.: +351 21 2948314; fax: +351 21 2948500. E-mail address: [email protected] (C.A.M. Portugal). http://dx.doi.org/10.1016/j.jbiotec.2014.09.009 0168-1656/© 2014 Elsevier B.V. All rights reserved.
non-limited access of the cell to extracellular nutrients and the removal of excreted waste products. The impact exhibited by the topography of scaffold surfaces on cell growth has been reported by several researchers. It is known that the use of cell supports with regular and aligned channels favours cell orientation, a phenomenon often referred to as contact ‘guidance’, leading to the formation of well-organized tissues (Morelli et al., 2010; Papenburg et al., 2010a; Truckenmüller et al., 2012). Moreover, several studies have revealed that the use of solid matrices with aligned patterns is also essential to assure adequate cell maturation and the regulation of diverse cell mechanisms (Chew et al., 2008). The presence of topographical induced cell orientation effects has been supported by confocal laser scanning microscopy studies using specifically labelled proteins (Braber et al., 1998; Morelli et al., 2010) and scanning electron microscopy (SEM) analysis (Zhong et al., 2006; Morelli et al., 2010) which revealed that cellular growth and proliferation occur parallel to the long axes of the surface patterns or, in case of scaffold surfaces with fibrous morphology, along the fibres. In contrast, flat or randomly
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patterned surfaces generate non-oriented cell growth leading to less structured cell tissues. Different hypothesis have been proposed to explain the diverse behaviours observed. Some authors suggested that they may be related to differences in local surface free energy (or surface wettability) that restrains protein and cell attachment to specific surface regions. Others have attributed the preferential cell orientation to the presence of a minimal length for cell attachment of 2 m that forces cells to attach along the longer axes of surface patterns (Izzard and Lochner, 1976). Alternatively, preferential cell attachment may be correlated with the anisotropic geometry of surfaces (Papenburg et al., 2010b) which forces the orientation of cells due to a required energetic balance between surface stresses and shear free planes (Chen, 2008; Guilak et al., 2009). This effect generates mechanotransduction processes which rule primarily cell shape (Guilak et al., 2009; Horbett, 1994) and then the intrinsic cell signalling mechanisms. These mechanisms are driven by the tensional forces between extracellular matrix (ECM) and cells upon cell attachment to scaffolds, which subsequently regulate cell proliferation and differentiation paths. Also, a recent study from Fujita et al. (2009) revealed that cell protrusions which expand perpendicularly to the nanogroove direction retract more quickly than those that move parallel to it. This effect led these authors to conclude that the cell retraction process may contribute to the cell elongation and alignment in nanogrooved surfaces. The relationship between surface topography and cell growth profile is still unclear; however, it has also been hypothesized that this cell contact guiding effect exerted by specific surface topographies may be directed by the structural status of the different adhesion proteins (e.g. fibrinogen, fibronectin, vibronectin, etc.) that primarily adsorb at the scaffold surface during the growth process. Based on this effect, it is reasonable to speculate that the efficiency of cell anchorage to scaffolds is intimately related to the mode how surface impacts on the structure and molecular flexibility of the adhesion proteins since it may influence the accessibility of cell receptors to protein regions (Imen et al., 2009), such as arginine-glycine-), such as arginine–glycine–aspartic acid (RGD) sequences, which are known to be determinant in cell attaching processes (Horbett, 1994). The ability and the way proteins adsorb at a given surface depend on their conformation and also the structural status of an adsorbed protein relies on the surface topography. This relationship was clearly demonstrated in the work developed by Roach et al. (2006) which highlights the contrasting effects induced on globular and rod-like proteins upon contacting surfaces with different patterns. These authors have shown that the contact of a rod-like protein, such as fibrinogen, to silica surfaces with accentuated curvature results in a significant change of its secondary structure, characterized by a loss of the ␣-helix structure in benefit of random coil chain segments, which is related to its capacity to fit the surface. Contrastingly, and due to their inability to adapt to surfaces with high curvatures, globular proteins exhibit an opposite structural behaviour when adsorbed at an identical surface, registering a decrease of the ␣-helix content with the decrease of surface curvature. Studies of the impact of surface topography in cell growth have shown that cell spreading, adhesion and differentiation also depend on the dimensions of the surface topographical features (Dalby et al., 2002, 2003), and fibre diameter (Elias et al., 2002) when fibrous scaffolds are used. However, a clear relationship between structural changes induced upon attachment of plasma proteins to the scaffold surface and the degree of cell orientation achieved with different surface topographies has not been established so far. Fluorescence spectroscopy is a highly sensitive technique capable of providing simultaneously valuable information about protein structure and mobility, which has not been sufficiently explored so
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far for monitoring of cell growth at tissue scaffolds. Therefore, in this work, a fluorescence monitoring methodology combining data from steady-state fluorescence and fluorescence anisotropy is used to assess the impact of surface topography on the structure and molecular flexibility of proteins. Protein structural changes can be inferred from the fluorescence emission of a protein intrinsic fluorescence probe, tryptophan. The use of tryptophan as a structural reporter excludes the need of protein labelling with an external chromophore. Based on the high sensitivity of the fluorescence emission of tryptophan to changes of the physicochemical properties in its surrounding environment, fluorescence emission allows for in situ monitoring of the changes of protein structure (Portugal et al., 2006, 2007; Guedidi et al., 2012) and the mode that each geometrically different protein interacts to the surface. Fluorescence anisotropy measurements allow for eliciting information about the influence of the different surface topographies on molecular flexibility. The present study was conducted using model proteins with opposite conformational geometries: a globular protein, trypsin (largely used in adsorption studies, presenting a significant fluorescence emission) and a rod-like protein, fibrinogen, which integrates the group of adhesion proteins directly involved in the cell–tissue scaffold interaction process. The two proteins were selected in order to mimic the conformational diversity (globular and rod-like proteins) of the proteins involved in the cell attachment process. This work aims at showing the potential impact of the structural status of proteins induced by scaffold surface topography on cell attachment using a non-invasive fluorescence monitoring approach. To achieve this goal, a correlation between the topographically induced structural changes of the adhesion proteins and the growth of C2C12 mouse pre-myoblast cells at these scaffolds was established. As described in previous papers from some of the authors of the present work (Haneveld, 2006; Papenburg et al., 2010a; Bettahalli et al., 2012) the use of fibrous scaffolds improves cell–cell interactions while allowing a good spreading of the C2C12 cells. 2. Experimental 2.1. Preparation of protein solutions Protein solutions of trypsin from bovine pancreas (EC. 3.4.21.4, Ref. T9935 from Sigma–Aldrich), a globular protein with MW of 20.3 kDa (Fig. 1(a)), and fibrinogen from bovine plasma (EC 232598-6, Ref. F8630, from Sigma–Aldrich), a rod-like protein with MW of 340 kDa (Fig. 1(b)), with respective concentrations of 2 g/L and 0.7 g/L were prepared in 0.1 M Trizma® buffer at pH 8 containing 0.02 M of CaCl2 , added to prevent trypsin autolysis. Trizma buffer was prepared using Trizma® hydrochloride (Sigma Aldrich) and Trizma® base (Sigma Aldrich). To avoid protein structural changes due to the physicochemical conditions, both protein solutions were prepared using the solvent media with identical characteristics. Protein concentration was selected in order to be in the range of the typical cell culture media protein content and to assure the detection of fluorescence emission from the protein molecules adsorbed at the scaffold surfaces. Differences in concentrations of both protein solutions were due to the low solubility of fibrinogen in the Trizma buffer used. 2.2. Protein adsorption on PLLA scaffolds with distinct topographies Trypsin and fibrinogen were separately adsorbed at poly-llactic acid (PLLA) scaffolds with different surface patterns: flat, aligned nanogroove pattern and random fibrous surfaces. Surfaces with aligned nano-patterning were obtained by using nano imprint
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Fig. 1. Schematic representations of (a) the three-dimensional globular structure of trypsin from bovine pancreas and (b) the three-dimensional rod-like structure of bovine fibrinogen (adapted from Brown et al., 2000) highlighting the location of tryptophans in red colour (structural probe). Both schematic representations were obtained and adapted from the Protein Data Bank at http://www.rcsb.org/pdb/home/home.do.
Fig. 2. Thermal NIL process for patterning a PLLA film with the following process steps: (a) insertion of the PLLA film, (b) evacuating the space between the film and an imprinting stamp from silicon, heating up the stamp and with it the film, and applying a gas pressure to imprint the stamp into the film, and (c) cooling down the stamp and the film, releasing the gas pressure, removing the vacuum and demoulding the imprinted film by peeling it off from the stamp ((1) sealing foil separating the vacuum section of the nano imprint lithography machine from its high-pressure section, (2) PLLA film to be imprinted, (3) imprint stamp from silicon (blue and red corresponds to cold and heated), (4) compressed gas. (For interpretation of the references to colour in this figure legend, the reader is referred to the web version of this article.)
lithography. The nanogroove patterns were imprinted (Fig. 2) in a FDA (U.S. Food and Drug Administration) compliant, 35 m thin, biaxially oriented blown film (nominal thickness: 30 m) from a blend of PLA (Plastic Suppliers/Sidaplax; EarthFirst Packaging Film, PF). The film had a glass transition temperature of 58 ◦ C, a melting temperature of approximately 160 ◦ C and the surface tension of the untreated surface was 38 mN m−1 . Imprinting was carried out in a machine for hot embossing nano imprint lithography (Obducat, Malmö, Sweden; NIL 6 ). The imprinting stamp from silicon carried a periodical pattern of parallel ridges with a width of 150 nm, a periodicity of 500 nm and a height of 100 nm. The stamp was fabricated by laser interference lithography and wet chemical etching (Haneveld, 2006). To facilitate demoulding, the stamp was functionalized with a perfluorodecyltrichlorosilane (FDTS; ABCR, Karlsruhe, Germany) anti-stiction layer by vapour deposition (Haneveld, 2006). In particular, the surface with aligned nano-pattern is composed of well-oriented channels with 150 nm width and 100 nm depth and an inter-channel spacing of 350 nm (Fig. 3(a) and (b)).
Fibrous scaffolds were prepared by dissolution of 12 wt% PLLA solution (PLLA mol wt. 1.6 × 105 g/mol) in 1,3-dioxane which was then spun using an in-house-built electrospinning set-up. SEM images were made to determine the surface topography and fibre diameter (Fig. 3(c) and (d)). For this, the sheets were sputter-coated with gold. Analysis of SEM results showed that the fibrous PLLA scaffold is composed by fibres with an average diameter of 2.5 m. The fibrous PLLA scaffolds obtained had a thickness of 200 m and a porosity of 71 ± 3% (Bettahalli et al., 2012). The adsorption of the proteins to the various scaffolds was induced by immersion of the scaffolds, for 3 h in 5 mL of trypsin or fibrinogen solutions under permanent external soft stirring, using a multi-mixer system (mistral, Lab-line Instruments). Upon this procedure, initial protein solutions (i.e. before the adsorption process) and the scaffold surfaces with adsorbed proteins were inspected using the fluorescence methodology described below to evaluate the influence of the surface topography on the structural status of proteins. The amount of the adsorbed proteins at the different surfaces was determined by inductively coupled plasma (ICP-AES) analysis using a Jobin Yvon-Ultima ICP spectrometer (Edison, NJ, USA). This analysis allow for spectroscopic quantification of proteins in samples acquired before and after adsorption based in the sulphur content, using its characteristic emission line at 180.676 nm. 2.3. Monitoring structural changes of the proteins induced upon interaction with the scaffold surfaces The structural changes underwent by the two proteins upon adsorption at the scaffold surfaces were assessed by using a fluorescence methodology, which combined the information provided by steady-state fluorescence emission and steady-state fluorescence anisotropy. 2.3.1. Steady-state fluorescence analysis The fluorescence scans were acquired at an excitation wavelength, exc , of 290 nm and at an emission wavelength, em , ranging from 300 to 550 nm, for the solutions of proteins before the adsorption and for proteins adsorbed at each scaffold surface. The fluorescence spectra were acquired using a SPEX Fluorolog spectrofluorimeter, equipped with excitation and emission polarizers positioned at the magic angle (54.7◦ ) and using excitation and emission slits of 5 nm bandpass. Steady-state fluorescence analysis enables us at inferring about changes of the three-dimensional structure of proteins induced upon adsorption at the different surfaces. The fluorescence emission obtained at exc of 290 nm is essentially due to the contribution of the indole group of the tryptophan residues present at each protein chain. Changes of the fluorescence characteristics of the
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Fig. 3. SEM images of the PLLA scaffolds with nanogrooved surface produced by nano imprint lithography obtained with magnifications of: (a) x 47.15K and (b) x 17.06K 47.15× K and (b) 17.06× K; electrospun PLLA scaffolds obtained with magnifications of (c) x 75 and (d) x 500, prepared by dissolution of 12 wt% PLLA solution in 1,3-dioxane.
tryptophan residues may result from a different proximity to quenchers present in the protein chain, such as aspartate, disulfidric and hydrogen bridges (Chen and Barkley, 1998), differences on the exposure degree of these fluorophores to the solvent, e.g. as result of the protein folding/unfolding processes, leading to displacements of the maximum emission. Shifts of the maximum emission to higher emission wavelengths (maximum emission red shift) may be ascribed to a higher exposure of the tryptophan residues to an environment with higher dielectric constant (or higher polarity) (Portugal et al., 2007; Faria et al., 2004). In contrast, a decrease of the dielectric constant in the microenvironment surrounding the fluorophores, resultant from a protein folding process, may lead to the dislocation of the maximum emission to lower emission wavelengths (maximum emission blue shift) followed by an increase of the fluorescence intensity (Portugal et al., 2007).
protein fluorescence spectra, leading to an increase of the fluorescence emission of the system and to an apparent dislocation of the maximum emission, which could prevent an accurate perception of the fluorescence emission changes induced by the adsorption process. To overcome this limitation, the fluorescence emission spectra of the adsorbed proteins were elicited by subtracting the fluorescence emission spectra obtained for scaffold surfaces before protein adsorption from the fluorescence emission spectra obtained for scaffold surface with the adsorbed protein, resulting in the dashdot line spectra shown in Fig. 4 (a). Fluorescence anisotropy from the adsorbed proteins, shown in Fig. 4 (b) was also determined according to the methodology described by Guedidi et al. (2012).
2.3.2. Steady-state fluorescence anisotropy analysis Fluorescence anisotropy provides information regarding the mobility of the fluorophores present in the protein. Changes of the mobility of fluorophores, either that concerning to tryptrophans intrinsic rotation ability or that due to protein mobility, can be correlated with the impact of protein interactions with its surrounding environment (i.e. folding/unfolding processes or protein–surface interactions) on their structure and molecular flexibility. The fluorescence anisotropy, r, was acquired using the spectrofluorimeter referred above using the same slits and spectral ranges, by varying the position of excitation and emission polarizers. Determination of fluorescence anisotropy, r, values is described in detail in previous works (Portugal et al., 2006, 2007; Guedidi et al., 2012).
A fluorescence monitoring methodology combining steadystate fluorescence emission and fluorescence anisotropy analysis was used in order to gather a better understanding of the role of the structural status of the adhesion proteins in the cell attachment process. In this work, fluorescence monitoring was applied to probe the impact of PLLA surfaces with flat, aligned nano-patterning and random fibrous pattern designs on the structure of two model proteins with distinct conformational geometries: trypsin, with a globular geometry (Fig. 1(a)) and fibrinogen, a rod-shaped protein (Fig. 1(b)), which were selected to mimic the diversity of proteins’ conformational geometries present in cell media, either they are directly involved in cellular adhesion or not. The number of tryptophans per protein chain was also taken as a model protein selection criterion, since it dictates the success of fluorescence monitoring. Fibrinogen is an adhesion protein, used as mediator of cell adsorption at tissue scaffolds, with a high number of tryptophans. Although, trypsin is not a cell binding protein, its tryptophan content facilitates the detection of this biomolecule in conditions where the perception of tryptophan signal is perturbed by external elements in proteins’ vicinity. This aspect is critical when monitoring the fluorescence of adsorbed proteins, in case
2.3.3. Determination of the fluorescence emission properties of the adsorbed proteins As illustrated in Fig. 4, the fluorescence emission of PLLA scaffold surfaces is characterized by the presence of a large emission band of low intensity with a maximum emission between 390 and 450 nm (depending on the scaffold surface topography). The emission band of the PLLA scaffolds superimposes at the red edge of the
3. Results and discussion
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Fig. 4. (a) Fluorescence emission spectra and (b) fluorescence anisotropy spectra obtained at exc of 290 nm for trypsin in solution before adsorption (1), PLLA fibrous surface (2), PLLA fibrous surface with adsorbed trypsin (3) and trypsin adsorbed at fibrous PLLA surface upon subtraction of the contribution of the PLLA surface (4).
of the contribution of the intrinsic fluorescence of the surface (e.g. tissue scaffold) at the protein emission region (Guedidi et al., 2012). The amount of trypsin and fibrinogen adsorbed to the surfaces with different topographies was determined by ICP-AES analysis. The results obtained revealed that the amount of trypsin adsorbed at flat and fibrous surfaces were 2.12 × 10−9 mol/cm2 (0.043 mg/cm2 ) and 3.99 × 10−11 mol/cm2 (0.81 g/cm2 ), respectively. When compared in molar terms, it was observed that the amount of fibrinogen adsorbed at flat and fibrous surfaces of 4.51 × 10−10 mol/cm2 (0.153 mg/cm2 ) and 7.22 × 10−12 mol/cm2 (2.45 g/cm2 ), respectively, was smaller, which is attributable to the significantly different protein sizes. 3.1. Fluorescence monitoring of the influence of surface topography on the structural behaviour of globular proteins Fig. 5(a) shows the fluorescence spectra obtained, at exc of 290 nm, for the globular protein, trypsin, before and after the adsorption at scaffolds with distinct surface topographies. For the easiness of the analysis of the results and for cancelling the dependence of fluorescence emission on the protein concentration, fluorescence changes were analyzed based on the comparison of normalized fluorescence emission spectra. As can be observed, the adsorption of trypsin at flat surfaces causes a clear displacement of the maximum emission to higher wavelengths, a red shift, with an amplitude of ∼6 nm, followed by the enlargement of the emission bandwidth (Fig. 5(a)). A maximum emission red shift of lower amplitude (∼2 nm) is also visible after the adsorption of this protein at surfaces with the fibrous morphology (Fig. 5(a)). The fluorescence emission changes detected reflect the increase of the dielectric constant at the local microenvironment of tryptophans due to their higher exposure to the external aqueous phase. Hence, this behaviour clearly indicates that the adsorption of trypsin at PLLA flat and fibrous surfaces generates the presence of molecular unfolding processes which, based on the amplitude of the fluorescence emission changes, seems to be more significant upon contact with flat surfaces. Trypsin exhibited a distinct structural behaviour upon adsorption at PLLA surface with an aligned nano-patterning. In this case, the adsorption is followed by the dislocation of the maximum emission to lower wavelengths, a blue shift (Fig. 5(a)). The blue shift observed is attributable to the increase of the hydrophobic character of the environment in the vicinity of the tryptophans. This effect may be due to a higher contact of the tryptophans with the
scaffold surface, which has a dielectric constant lower than water or to protein structural folding processes induced by their contact with the surface with such topography. The information about the structural changes of trypsin induced by the adsorption process was complemented by the analysis of the rotational ability of tryptophans provided by steady-state fluorescence anisotropy. From the anisotropy spectra shown in Fig. 5(b), it is possible to conclude that the rotational ability of tryptophans is also influenced by the surface topography. The attachment of trypsin to flat surfaces and surfaces with aligned patterning lead to an increase of its fluorescence anisotropy, expressing a hindered rotational ability of tryptophan residues upon protein adsorption, compared to that observed for trypsin in solution. There is an obvious agreement between the increase of fluorescence anisotropy and the maximum emission blue shift observed upon adsorption at the surface with aligned pattern. Indeed, the molecular processes which explain the presence of a maximum emission blue shift, i.e. an additional molecular folding or an increase of the contact degree of tryptophans towards the surface, may both justify the restricted tryptophan mobility. Contrastingly, the maximum emission red shift, i.e. trypsin unfolding, followed by a reduction of tryptophans’ mobility, as monitored upon trypsin adsorption at the flat surface, seems contradictory at a first glance. However, both effects may result from a combination of different molecular events inherent to the adsorption process, which generate apparently opposite alterations on the fluorescence emission and anisotropy spectra. The total mobility of tryptophans results from the sum of different tryptophan rotational components: the intrinsic rotational ability of tryptophans, which is hampered by the protein’s threedimensional structure; and the mobility of the protein as a whole, which contributes to an additional mobility of the tryptophan residues. Based on this explanation, the loss of tryptophans’ mobility observed upon attachment at the flat surface may be mainly attributed to a decrease of the whole protein mobility after attaching to the scaffold surface. However, the tryptophan imprisonment, which should result in a maximum emission blue shift, was probably not enough to surpass the protein maximum emission red shift induced by unfolding. An opposite structural behaviour was observed upon adsorption of trypsin at random fibrous surfaces. The decrease of fluorescence anisotropy (Fig. 5(b)) and the small maximum emission red shift (Fig. 5(a)) observed in this case supports the presence of a slight trypsin unfolding, with the consequent increase of tryptophans
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Fig. 5. Fluorescence emission (a) and anisotropy spectra (b) of trypsin in solution (1), trypsin adsorbed at PLLA flat surface (2), trypsin adsorbed at PLLA fibrous surface (3) and trypsin adsorbed at PLLA aligned nano-patterned surface (4).
mobility. Together, these results suggest the absence of significant structural changes upon trypsin attachment to fibrous surfaces. Still, fluorescence data allowed us to speculate that the trypsin structural changes are potentially followed by an incomplete or less efficient attachment of trypsin at the fibrous scaffold. The fluorescence emission changes observed for the trypsin adsorbed at flat and fibrous surfaces are in concordance with the results reported in the literature. Our results show that the adhesion of globular proteins to flat or fibrous surfaces is clearly preceded by an unfolding step, which is compatible with the increase of random-coil character of protein structure (Roach et al., 2006). Moreover, the proportion of random coil segments increases with the increase of the radius of the surface curvature, which is also in agreement with the highly intense unfolding achieved upon adsorption of globular protein at flat surfaces. Additionally, it was observed that the adsorption of globular proteins at flat surfaces is followed by a structural spreading of the adsorbed protein (Agnihotri and Siedlecki, 2004; Roach et al., 2006) whereas minor alterations are expected due to their adsorption at surfaces with high curvatures (such is the case for fibrous scaffolds). The molecular spreading of globular proteins at flat surfaces results from the presence of a higher contacting surface area available, and explains the trypsin unfolding as well as the improved contact of the tryptophans with the flat surface. The ability of trypsin to spread at surfaces with higher curvatures is limited due to the reduced contacting area available. Although in a lower extent, trypsin unfolding still occurs, resulting in minor maximum emission red shift amplitudes (as shown in Fig. 5 (a)), contributing to the increase tryptophan mobility. The increase of tryptophans rotational ability also evidences the presence of random coil protein regions, which are unable to bind at the scaffold surface. 3.2. Fluorescence monitoring of the effect of topography on the structure of rod-like proteins A similar study was conducted to elicit information about the influence of scaffold surfaces with different topographies on the conformation of a rod-like protein, fibrinogen (Fig. 6). Analysis of the fluorescence emission spectra obtained for the fibrinogen before and after adsorption at surfaces with flat, aligned nano-channels and fibrous morphology (Fig. 6(a)) reveals that the adsorption of fibrinogen does not cause displacements of its maximum emission as intense as those observed after adsorption of
trypsin at identical surfaces. Only a subtle blue shift, with no more than to 2 nm, followed by the enlargement of the emission band, was observed after the adsorption of this protein at the surface with aligned nanogrooves. Complementary fluorescence anisotropy results, depicted in Fig. 6 (b), show that despite the absence of significant fluorescence emission changes, the adsorption of fibrinogen at scaffold surfaces is followed by the alteration of fluorescence anisotropy. Indeed, the adsorption of fibrinogen at flat surfaces leads to an increase of the fluorescence anisotropy, relatively to the anisotropy of fibrinogen in solution. As referred previously, the increase of fluorescence anisotropy corresponds to a decrease of tryptophans mobility, attributable either to the contact of tryptophans to the surface or to an additional folding of the protein regions containing the tryptophans. In this case, tryptophan–surface interactions are facilitated by the elongated conformation of fibrinogen, which allows for an improved contact of amino acids to the flat surface without involving a significant change of the protein structure. The higher increase of fluorescence anisotropy and the maximum emission blue shift observed for fibrinogen adsorbed at the nanogrooved surface suggest a more intense contact of tryptophans to the nanogrooved surface, which may be related to a highly stable protein adhesion and to a good cell attachment and growth observed at these scaffolds (Papenburg et al., 2007; Truckenmüller et al., 2012). Nevertheless, in this case, the anisotropy reached values higher than 0.4, the maximum limit theoretically possible for fluorescence anisotropy (Valeur, 2002). The high fluorescence anisotropy might be related to the presence of light scattering and reflection phenomena by the surface, disabling us from taking clear conclusions from these results. The adsorption of fibrinogen at fibrous surfaces generates a decrease of the fluorescence anisotropy (Fig. 6(b)), a similar behaviour to that found upon adsorption of trypsin (globular protein) at PLLA surfaces with the same morphology (Fig. 5(b)). Identically to that discussed for trypsin, the decrease of the fluorescence anisotropy of fibrinogen adsorbed at fibrous surface expresses higher tryptophan mobility, attributable to a higher fibrinogen unfolding degree upon adsorption. In the native conformation of fibrinogen, tryptophans are already highly exposed to the aqueous media, which is confirmed by the average lifetime of tryptophans in free fibrinogen, which is equal to 2.71 ns (result obtained by time-resolved fluorescence),
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Fig. 6. (a) Fluorescence emission and (b) fluorescence anisotropy spectra obtained at exc of 290 nm for fibrinogen in solution (1), adsorbed at flat PLLA surface (2), adsorbed at fibrous PLLA surface (3) and adsorbed at aligned nano-patterned surface (4).
that corresponds to a lifetime close to that expected for tryptophans totally exposed to the aqueous environment, which is equal to ∼3 ns (Petrich et al., 1983). As discussed above, the absence of fluorescence emission changes suggests that the adsorption of fibrinogen does not result in a significant increase of the exposure degree of tryptophans to the external aqueous environment. Hence, the increase of the protein unfolding degree may be explained based on the ability of rod-like proteins, such as fibrinogen, to mould to surface curvatures (Roach et al., 2006). Although the curvature radius of the surface used in this work was not significantly accentuated regarding the protein size, it may be enough to force the amino acids to pull apart upon fibrinogen adsorption. Also it may be speculated that the adsorption is associated to an additional unfolding of the nodular regions of fibrinogen, which contains most of the tryptophans present in the fibrinogen chain (Fig. 1(c)). This effect would justify the higher mobility of tryptophans and a better accessibility to protein regions with a relevant role in the cell adhesion, which were initially hidden within the protein structural core. According to literature, RGD sequences or the dodecapeptide at the carboxyl terminus of the ␥-chain were found to be one of the preferential cell binding sites at fibrinogen (Horbett, 1994). These segments are located in different fibrinogen domains, in particular they are placed within D domain ␥ 400–411, C domain at A␣ 98–101 and at the coiled-coil regions A␣ 255–258, which in turn are placed at the extreme nodular regions of fibrinogen where the accessibility of the amino acids in the native protein may be limited (Brown et al., 2000; Mosesson et al., 2001). The partial unfolding of proteins, specifically fibrinogen, upon adsorption promotes the exposure of these chain segments improving cell–protein interactions, and would justify the contribution of molecular unfolding to the enhancement of cell adhesion performance, as observed at fibrous scaffolds. In fact, recent studies performed by some of the authors of the present work showed that there was higher cell–cell contact on the fibrous PLLA sheets compared to the PLLA flat sheets and tissue culture polystyrene (TCPS) (Papenburg et al., 2010a). This is in agreement with literature, where it was reported that in various cell types, integrins associated with cell–cell as well as cell–ECM contact were up-regulated on nano-fibrous scaffolds compared to flat surfaces as response to increased protein adsorption (Kumar, 2006). Also, SEM images supported this observation as the C2C12 cells spread well on the fibrous PLLA sheets and show good connection between the cells’ filopodia as well as between the cells and the nano-fibrous surface. The DNA assay data also
indicated slightly increased cell proliferation on the fibrous PLLA compared to the flat PLLA, although only significant at day 4 for the higher cell density (Kumar, 2006). Moreover, as shown by Pagliara et al. (2010), electrospinning and nano imprint lithography induces molecular orientation within polymer structure, resulting in the increase of light polarization and consequently an increase of fluorescence anisotropy. Therefore, the adsorption of proteins to structurally aligned or random surfaces may promote preferential or disordered orientations of the transition dipole moments of tryptophans towards the excitation light, also contributing to the increase or decrease of fluorescence anisotropy, respectively. This behaviour suggests the alignment of protein molecules at the nanogrooved surface, which corroborates the improved cell orientation of tissues obtained using such scaffold surfaces (Truckenmüller et al., 2012; Papenburg et al., 2007). 4. Concluding remarks A non-invasive fluorescence monitoring methodology was used to access the impact of different surface topographies on the structural status of proteins, aiming at establishing a correlation between the structural behaviour of adhesion proteins adsorbed at the scaffold surfaces and the cell growth process. Fluorescence analysis simultaneously provided information about the folding/unfolding status of the adsorbed proteins as well as their molecular mobility level. Together, this information allowed for inferring about protein binding intensity, accessibility to hindered protein regions and the presence of anisotropic structural features which may support the presence of topographically induced oriented cell attachment. The fluorescence results obtained confirmed that the structural behaviour of proteins when interacting with surfaces depends not only on the surface pattern design but also on the protein conformational geometry (Roach et al., 2006). The latter dictates the ability of proteins to fit the surface and, together with the surface chemistry, it rules the impact of the adsorption process on the protein conformation. As expressed by the increase of fluorescence anisotropy, the adsorption of globular or rod-shaped proteins at flat surfaces and surfaces with aligned nano-patterning leads to a high restriction of protein mobility, comparatively to that produced by the adsorption at fibrous surfaces. This structural behaviour stands for an intense surface–protein interaction, which may justify a stable protein
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attachment. The increase of fluorescence anisotropy may also comprise the contribution of a preferential alignment of the adsorbed proteins driven by the surface channel pattern. This behaviour is corroborated by the alignment of FITC labelled proteins adsorbed at grooved surfaces monitored by Braber et al. (1998) using confocal laser scanning microscopy. In contrast, the adsorption of both globular and rod-shaped proteins to random fibrous scaffolds leads to a decrease of fluorescence anisotropy to values even smaller than that obtained for proteins in solution. These results may be interpreted by an amplification of the rotational ability of tryptophans, which is higher upon adsorption of rod-shaped than globular proteins, and may reflect the ability of these molecules to mould to curved surfaces (Roach et al., 2006). Such structural alterations offer an improved exposure of the amino acids to the external media and support the contact of important protein chain segments, e.g. RGDs sequences (Horbett, 1994) to the receptors of membrane cell proteins, explaining the better cell growth efficiency obtained when fibrous scaffolds are used as cellular supports (Papenburg et al., 2010a; Woo et al., 2003). Also, it may be rationalized as the presence of non-oriented adsorbed proteins at the surface of randomly aligned fibrous scaffold. Hence, the in situ fluorescence monitoring approach used in this work shows clearly that cell attachment efficiency is ruled by a combined action of two effects: the preferential adsorption and alignment of proteins according to specific surface topographical features and the topographically induced protein structural status. Additionally, fluorescence monitoring has shown to be a powerful tool that can be used as a single technique to guide the development of tissue scaffolds with improved designs and also the configuration of new engineered adhesion peptides (Sprio et al., 2011), contributing in this mode to the improvement of cell attachment processes. Acknowledgements The authors would like to acknowledge to Prof. Fernando Pina, Prof. Maria João Melo and Prof. João Carlos Lima the access to the spectrofluorimeter. The authors also acknowledge the funding from Fundac¸ão para a Ciência e a Tecnologia (FCT-MEC), Portugal, through the project PEstC/EQB/LA0006/2011. References Agnihotri, A., Siedlecki, C., 2004. Time-dependent conformational changes in fibrinogen measured by atomic force microscopy. Langmuir 20, 8846–8852, http://dx.doi.org/10.1021/la049239+. Bettahalli, N.M.S., Groen, N., Steg, H., Unadkat, H., de Boer, J., van Blitterswijk, C.A., Wessling, M., Stamatialis, D., 2012. Development of multilayer constructs for tissue engineering. J. Tissue Eng. Regen. Med., http://dx.doi. org/10.1002/term.1504. Braber, E.T., de Ruijter, J.E., Ginsel, L.A., von Recum, A.F., Jansen, J.A., 1998. Orientation of ECM protein deposition, fibroblast cytoskeleton, and attachment complex components on silicone microgrooved surfaces. J. Biomed. Mater. Res. 40, 291–300, http://dx.doi.org/10.1002/(SICI)1097-4636(199805)40: 2<291::AID-JBM14>3.0.CO;2-P. Brown, J.H., Volkmann, N., Jun, G., Henschen-Edman, A.H., Cohen, C., 2000. The crystal structure of modified bovine fibrinogen. Proc. Natl. Acad. Sci. U.S.A. 97, 85–90, http://dx.doi.org/10.1073/pnas.97.1.85. Chen, C.S., 2008. Mechanotransduction – a field pulling together? J. Cell Sci. 121, 3285–3292, http://dx.doi.org/10.1242/jcs.023507. Chen, Y., Barkley, M.D., 1998. Toward understanding tryptophan fluorescence in proteins. Biochemistry 37, 9976–9982, http://dx.doi.org/10.1021/bi980274n. Chew, S.Y., Mi, R., Hoke, A., Leong, K.W., 2008. The effect of the alignment of electrospun fibrous scaffolds on Schwann cell maturation. Biomaterials 29, 653–661, http://dx.doi.org/10.1016/j.biomaterials.2007.10.025. Dalby, M.J., Riehle, M.-O., Johnstone, H.J.H., Affrossman, S., Curtis, A.S.G., 2002. Polymer demixed nanotopography control of fibroblast spreading and proliferation. Tissue Eng. 8, 1099–1108, http://dx.doi.org/10.1089/107632702320934191. Dalby, M.J., Riehle, M.-O., Johnstone, H.J.H., Affrossman, S., Curtis, A.S.G., 2003. Non-adhesive nanotopography: fibroblast response to poly(n-butyl methacrylate)-poly(styrene) demixed surface features. J. Biomed. Mater. Res. 67A, 1025–1032, http://dx.doi.org/10.1002/jbm.a.10139.
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