Biointerfaces Designed Through Directed Collagen

Copyright © 2014 American Scientific Publishers All rights reserved Printed in the United States of America

Journal of Bionanoscience Vol. 8, 407–418, 2014

Biointerfaces Designed Through Directed Collagen Assembly Sara Mauquoy, Emilienne Zuyderhoff, Jessem Landoulsi† , Deepak Kalaskar‡ , Sophie Demoustier-Champagne, and Christine Dupont-Gillain∗ Institute of Condensed Matter and Nanosciences, Université catholique de Louvain, Croix du Sud 1 (Bte L7.04.01), 1348 Louvain-la-Neuve, Belgium

REVIEW

Creating substrates mimicking the extracellular matrix (ECM) is a promising approach to better control cell-material interactions. The aim of this research is to use collagen, a major component of the ECM, to elaborate nanostructured biointerfaces. Different strategies were implemented and are reviewed here. First, chemically patterned substrates were used to confine collagen assembly in given domains. Then, layer-by-layer (LbL) assembly was used to incorporate collagen in thin multilayered films, using assembly with either poly(styrene sulfonate) or fibronectin, another ECM component involved in cell adhesion. Finally, collagen-based nanotubes were synthesized using LbL assembly within the pores of a membrane. Electrophoretic deposition was used to immobilize these nanotubes at the surface of indium-tin oxide-coated glass, and preosteoblast cells were shown to interact with the nanotubes. Taken together, these methods improve our ability to direct collagen adsorption, thereby producing tailored biointerfaces for applications in biomaterials science and tissue engineering.

Keywords: Biointerfaces, Biomimetic Surfaces, Protein Adsorption, Collagen, Heterogeneous Surfaces, Layer-By-Layer Assembly, Nanotubes, Preosteoblasts.

CONTENTS 1. 2. 3. 4.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Materials and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Collagen Adsorption on Patterned Substrates . . . . . . . . . . . . . . Layer-by-Layer Assemblies Based on Collagen . . . . . . . . . . . . . 4.1. Collagen/Poly(Styrene Sulfonate) Films . . . . . . . . . . . . . . . 4.2. Collagen/Fibronectin Films . . . . . . . . . . . . . . . . . . . . . . . . 5. Surface-Immobilized Collagen Nanotubes . . . . . . . . . . . . . . . . . 6. Conclusion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

407 410 410 412 413 413 417 417 418 418

1. INTRODUCTION In vivo, cells rest on or are embedded in the extracellular matrix (ECM), which is a complex and highly structured assembly of proteins, polysaccharides and proteoglycans.1 Cells are connected to the ECM through receptor-mediated interactions. In particular, dimeric transmembrane proteins ∗

Author to whom correspondence should be addressed. Present address: Laboratoire de Réactivité de Surface, Université Pierre et Marie Curie, Case Courrier 178, Place Jussieu 4, 75252 Paris cedex 05, France ‡ Present address: Division of Surgery and Interventional Science, Faculty of Medical Sciences, University College London Royal Free NHS Trust Hospital, Pond St, Hampstead, NW3 2QG, United Kingdom †

J. Bionanosci. 2014, Vol. 8, No. 6

called integrins are involved in the anchorage of cells to ECM adhesion proteins (including collagens, fibronectin, vitronectin, laminin). The integrin-adhesion protein interaction has further consequences on cytoskeleton organization and hence influences many cell processes such as cell attachment, adhesion, spreading, growth, migration and differentiation.2 Creating substrates which mimic the ECM is thus an adequate strategy to better control cellmaterial interactions, which is of outermost importance in biomaterials science and tissue engineering. While layers of ECM components are commonly used to culture cells in vitro, providing them with anchoring sites and allowing cell growth,3 it becomes evident that controlling the organization of these components would lead to a better tuning of cell behavior.4–6 Collagen is the most abundant protein in the animal kingdom and is a major component of ECM where it forms insoluble fibrils. There are actually more than 10 types of collagen molecules. Type I collagen, made of three helical polypeptides gathered into a superhelix (molar mass ∼300 kg/mol; length ∼300 nm; diameter ∼1.5 nm), is predominant in skin, tendons and bones7 and offers many recognition sequences for cell integrins and for other adhesive proteins of the ECM.8 It is a self-assembling molecule which aggregates into fibrils in vivo as well as in vitro.

1557-7910/2014/8/407/012

doi:10.1166/jbns.2014.1270

407

Biointerfaces Designed Through Directed Collagen Assembly

Mauquoy et al.

Sara Mauquoy graduated in Bioengineering in 2012 at Université catholique de Louvain (UCL). She is currently doing her Ph.D. thesis in the team of Professor C. Dupont-Gillain at UCL, in the Institute of Condensed Matter and Nanosciences. She works on the creation of biomimetic surfaces, based on collagen and fibronectin, to influence stem cells behavior.

REVIEW

Emilienne Zuyderhoff obtained her Ph.D. degree in agricultural sciences and bioengineering in 2012, from Université catholique de Louvain (Belgium), where she worked under the supervision of Professor Christine Dupont-Gillain on the development of organized interfacial collagen layers and on their influence on cell behavior.

Jessem Landoulsi received his Ph.D. in materials science at the Technical University of Compiégne (UTC, France) in 2008. After a postdoctoral work at the Institute of Condensed Matter and Nanosciences (UCL, Belgium), he joined the Laboratory of Surface Reactivity of the University of Pierre & Marie Curie (UPMC Paris VI, France) to work on the interfaces between materials and biological systems. His current research interest is in probing the interactions between (bio)macromolecules and nanostructured surfaces at the molecular level, with the aim to design biomimetic surfaces and to explore relevant biological events at the nanoscale, such as protein adsorption, molecular recognition and cell adhesion. He has published over 33 peer-reviewed journal articles and 3 book chapters. Deepak Kalaskar has a multidisciplinary background in engineering, chemistry and biology. He has worked in both industry and academia. He did his Ph.D. in the field of Biomedical Materials from the University of Manchester, UK. Following his Ph.D., he worked at various universities in UK and Belgium on number of developmental research projects in the field of Bionanomaterials and Stem cells. Currently, Dr. Kalaskar is Lecturer in Cellular Engineering at UCL, London. He is deputy director of the postgraduate course in Burns, Plastic and Reconstructive surgery at UCL and actively involved in research on nanotechnology and tissue engineering solutions for artificial organs such as synthetic nose, ear, skin and bone & tendon tissue engineering. Sophie Demoustier-Champagne received her Ph.D. degree in Chemistry Sciences from the Université catholique de Louvain (UCL) in 1995. In 1999, she got a research associate position at the F.N.R.S. (Fonds National de la Recherche Scientifique, Belgium) and she is currently Full Professor at UCL. Her current main research topics are the elaboration, characterization and assembly of various functional nanowires and nanotubes, as well as, the bio-functionalization of surfaces and nano-objects for applications in the biomedical field. She is author of several book chapters, 90 papers and 3 patents.

408

J. Bionanosci. 8, 407–418, 2014

Mauquoy et al.


Christine Dupont-Gillain holds a Ph.D. in agricultural sciences and bioengineering. She is currently lecturer at the Faculty of Bioengineering, Université catholique de Louvain (Belgium). She leads a research group specialized in the physical chemistry of biointerfaces.

J. Bionanosci. 8, 407–418, 2014

named layer-by-layer (LbL) assembly.18 Such construction is based on the overcompensation of the surface charge at each polyelectrolyte adsorption step. This overcompensation both regulates the adsorption of the polyelectrolyte by preventing its self-association and allows the adsorption of the oppositely charged layer.19 LbL assembly opened the way to the combination of the properties of two or more polyelectrolytes on surfaces with various shapes.18 This simple, versatile, environment-friendly and low-cost method products films with different structures and growth behaviors depending on the deposition conditions (pH, concentration, adsorption time, ionic strength, solvent, temperature, etc).20 21 Incorporating biomolecules in LbL assemblies is an attractive strategy to produce biomimetic interfaces. The assembly of proteins can be considered, although it is challenging to predict their behavior since proteins are weak polyelectrolytes and their charge thus depends on pH. Moreover, they are polyampholytes and, while their global charge can be predicted based on their isoelectric point (iep), which can be computed using the amino acid sequence, local domains can be found in which the charge differs from the average one. This is especially the case for collagen, in reason of the very anisotropic shape of the molecule. The theoretical iep of collagen is close to 9, while experimental determination of collagen iep gave values ranging from 5.5 to 9.3.22 It is thus necessary to determine the conditions of collagen assembly at interfaces experimentally. Several groups have combined collagen and hyaluronic acid (HA) through LbL assembly, with a view to mimic the ECM. Johansson observed good build-up at pH 4.7, and the obtained films presented fibrils.23 Such fibrils were also observed by Zhang24 for similar assemblies built starting from an anchoring layer of polyethyleneimine (PEI), which gave thicker films in similar conditions but with longer adsorption times. Collagen/HA films were shown to increase the proliferation and to stimulate the differentiation of preosteoblasts cells on titanium substrate.25 Multilayered films made of collagen and antithrombogenic heparin are also studied to increase the blood compatibility of materials.26–30 Such combination is expected to favor endothelization while imparting antithrombogenicity to devices such as stents. A decreased platelet adhesion and a 409

REVIEW

Type I collagen is thus the ideal building block to create biointerfaces mimicking the ECM. In previous works, we have shown that the supramolecular organization of adsorbed type I collagen layers depends on the substrate surface properties (chemical nature, topography), on the characteristics of the collagen solution (concentration, state of aggregation) and on the details of the preparation procedure (duration of adsorption, mode of drying).9 10 The formation of fibrillar structures in the adsorbed phase was favored by the use of smooth and hydrophobic substrates and by increasing adsorption duration and/or collagen concentration. For example, multimolecular collagen assemblies were formed at the surface of polystyrene, a hydrophobic polymer, while a mat of individual collagen molecules was found at the hydrophilic surface of plasma-oxidized polystyrene.11 Moreover, discontinuous collagen layers could be obtained by dewetting.12 More generally, many observations have been made on the ability of collagen to form diverse supramolecular structures in the adsorbed state. Atomic force microscopy (AFM) was an essential tool to unravel collagen organization at the supra/molecular scale. Particularly, this technique allows the interface to be probed in aqueous phase with minimal disturbance. The distribution of adsorbed proteins in the plane of the interface can be arranged into nanometer-scale patterns, mainly using two approaches. On the one hand, proteins can be selectively deposited on defined areas of a homogeneous substrate by means of nanoscale physical contact between this substrate and the tip of an atomic force microscope13 14 or an elastomeric stamp.15 On the other hand, substrates presenting chemical heterogeneities can be used, with defined areas favoring protein adsorption while the remaining areas are designed to repel proteins.16 A variety of methods are nowadays available to produce such chemically heterogeneous substrates, as reviewed elsewhere.17 In most cases, poly(ethylene glycol) (PEG) is used to functionalize the domains preventing protein adsorption. ECM being organized in three dimensions, methods allowing the vertical organization of ECM compounds to be controlled are desired as well. In 1992, Decher suggested to alternate the adsorption of a polycation and a polyanion to build multilayered ultrathin films, in a process

Mauquoy et al.

REVIEW


better adhesion, spreading and proliferation of endothelial cells was indeed observed on stainless steel coated with collagen/heparin assemblies compared to virgin stainless steel.28 Collagen was also combined through LbL assembly with chondroitin sulfate30 and with alginate.31 32 To the best of your knowledge, collagen is always used as a polycation for LbL assembly. Landoulsi tried to use collagen as a polyanion in assemblies with PEI by working at pH 8.3. However, the growth of the multilayered film stopped after the adsorption of three layers, possibly because the high pH promoted collagen fibrillation.22 Assembling collagen and fibronectin together would constitute a further step towards the development of biomimetic interfaces. Collagen is indeed the most important structural component of ECM, and fibronectin (Fn) is an adhesion protein involved in many cell processes. These two proteins could possibly be assembled based on electrostatic interactions, although this is difficult to predict given the uncertainty regarding collagen electrical properties (iep ∼5 for Fn; iep ∼5.5–9.3 for collagen, see here above). A specific interaction is however known to occur between these two proteins,8 33 which could be exploited for the build-up of LbL assemblies. To the best of our knowledge, Fn/collagen LbL assemblies have not been reported in the literature. LbL assembly based on specific interactions between Fn and gelatin, a compound obtained by collagen hydrolysis, was however described.34–49 QCM results showed that the construction is efficient on a phospholipid bilayer.34 Fn/gelatin multilayers were also deposited on cell layers to obtain three-dimensional cell multilayers.38 The same assemblies made directly around hepatocyte carcinoma cells protected them from physical stress,35 and if made around human aortic smooth muscle cells and human umbilical vein endothelial cells, they allowed artificial multilayered blood vessels to be constructed.39 This paper reviews approaches aiming at preparing biointerfaces with an improved control of collagen organization. Different strategies were explored. First, patterned substrates were prepared with a view to confine collagen and to trigger its aggregation in defined areas. Then, LbL assembly was used to incorporate collagen in multilayered films. In particular, we report first attempts to build Fn/collagen LbL assemblies. Finally, interfaces were decorated with collagen-based nanotubes, which were built through LbL assembly within the pores of a template. Throughout the paper, the usefulness of AFM to observe the supramolecular organization of biointerfaces and to unravel mechanisms of layer formation is highlighted.

UK) from solution at 100 g/ml and fibronectin (Invitrogen, Life Technologies, Gent, Belgium) from solution at 25 g/ml. Denatured collagen is produced by heating of native collagen solution during 1 h at 90 C. Poly(ethyleneimine) (Fluka, Sigma-aldrich, Steinheim, Germany) is used at a concentration of 1 mg/ml. All the solutions are prepared in phosphate saline buffer (PBS; pH = 7.4), prepared with tablets (Gibco, Life technologies, Paisley, UK) dissolved in ultrapure water. All the protein solutions are prepared and stored at 4 C and are brought to working temperature 15 min before use. QCM (Q-sense, Goteborg, Sweden) experiments: The LbL assembly is performed in the liquid cell of the QCM, with the flow fixed at 30 l/min and temperature set at 25 C. The gold-covered quartz crystals (Standard Gold QSX 301,Q-sense, Goteborg, Sweden) used as sensors were spin-coated during 20 s at 2000 rpm with polystyrene (Aldrich, Milwaukee, USA) 0.5% w/w in toluene (Prolabo, VWR, Leuven, Belgique). After establishing the baseline in PBS, the adsorption of PEI is performed during 30 min (when required), followed by a rinsing step in PBS of 30 min. The adsorption of each protein layer is performed during 10 min, followed by a rinsing step in PBS of 30 min. Film preparation outside QCM: LbL films were prepared on spin-coated polystyrene substrates. Spin-coating is performed with 20 g/L PS in toluene at 5000 rpm during 40 s on previously washed (H2 O2 /H2 SO4 1:2; UV/O3 15 min) silicon wafers. PEI, n-Col, d-Col and Fn are adsorbed during 30 min. After each adsorption step, three rinsings with PBS are performed during 1 min. The temperature is fixed at 37 C. At the end of the construction, the films are rinsed 7 times with water, dried under a nitrogen flow and stored in a desiccator before analysis. Atomic force microscopy (Nanoscope V multimode, Digital Instruments, Santa Barbara, USA): AFM topographic images are acquired in tapping mode with AFM tips with a spring constant of 20 to 80 N·m−1 and resonant frequency in air from 358 to 382 Hz (Veeco, Camarillo, Canada). The amplitude set point was fixed to 250 mv. Image sizes were 1 × 0.5 m2 , 10 × 5 m2 , 50 × 25 m2 . Images are treated with the Nanoscope analysis software for flattening and structure size measurements. Water contact angle measurement: Contact angles are measured using the sessile drop method. A 0.3 l drop of ultrapure water is deposited on the surface with a syringe and the angle value is read after 5s. Plotted data are an average on 5 or 6 measurements on the same sample and error bars are confidence intervals.

2. MATERIALS AND METHODS

3. COLLAGEN ADSORPTION ON PATTERNED SUBSTRATES

This section presents materials and methods related to Section 4.2 (unpublished results). Experimental methods of the other sections can be found in given references. Materials: LbL assembly is performed between collagen type I (Biochrom AG, Source Bioscience, Nottingham, 410

Collagen was adsorbed on chemically heterogeneous substrates with a view to trigger its aggregation in defined areas, thereby producing biointerfaces mimicking ECM. These substrates were designed based on the knowledge J. Bionanosci. 8, 407–418, 2014

Mauquoy et al.

Biointerfaces Designed Through Directed Collagen Assembly (a) CH3/PEG

(b) PS/PMMA

PEG + col

PMMA + col

NXPS = 9 %

NXPS = 4 %

CH3 + col

PS + col

NXPS = 16%

PS/PMMA 35/65

CH3/PEG 50/550

(friction)

(phase)

REVIEW

CH3/PEG 50/550

NXPS = 16 %

PS/PMMA 35/65 + col

NXPS = 12 %

CH3/PEG 50/550 + col

PS/PMMA 65/35

CH3/PEG 95/55 + col

PS/PMMA 65/35 + col

(phase)

NXPS = 14 %

Fig. 1. (a) CH3 /PEG system: AFM images (z-range = 10 nm, except for friction image for which z-range = 0.3 V) of collagen (+col) adsorbed on homogeneous PEG and CH3 substrates (5 m × 2.5 m; insets: 750 nm × 375 nm), of naked heterogeneous CH3 /PEG substrates produced by combination of electron beam lithography and silanation (2 m × 1 m), and of heterogeneous CH3 /PEG substrates after collagen adsorption (+ col) (2 m × 1 m, scale bar = 200 nm for zoom on collagen assembly). The width of CH3 and PEG tracks is given in nm. (b) PS/PMMA system: AFM images (5 m × 2.5 m, z-range = 30 nm except for phase images for which z-scale = 10 ) of collagen (+col) adsorbed on homogeneous PMMA and PS substrates, of naked mixed PS/PMMA substrates produced by polymer demixing in thin films (PS domains appear in dark color while PMMA domains appear in lighter color on phase images), and of PS/PMMA substrates after collagen adsorption (+col). The fraction of PS and PMMA in the surface layer of heterogeneous films is given. NXPS refers to the molar fraction of nitrogen detected by XPS. Panel (a) is adapted with permission from Ref. [40], F. A. Denis, et al., Small 1, 984 (2005). © 2005, Wiley. Panel (b) is adapted with permission from [41], E. M. Zuyderhoff and C. C. Dupont-Gillain, Langmuir 28, 2007 (2012). © 2012, American Chemical Society.

acquired when studying the organization of adsorbed collagen on homogeneous substrates. Figure 1 presents AFM images obtained after type I collagen adsorption on (i) a monolayer of PEG-terminated alkylsilanes assembled on a silicon wafer (hereafter named PEG) (Fig. 1(a)), (ii) a monolayer of methyl-terminated alkylsilanes assembled on a silicon wafer (hereafter named CH3 (Fig. 1(a)), (iii) a thin spin-coated layer of poly(methyl methacrylate) (PMMA) (Fig. 1(b)), J. Bionanosci. 8, 407–418, 2014

(iv) a thin spin-coated layer of polystyrene (PS) (Fig. 1(b)). Experimental details regarding substrate preparation and collagen adsorption were described previously (for PEG and CH3 , see Ref. [40]; for PMMA and PS, see Ref. [41]). On PEG, a sparse network of individual collagen molecules is observed. On PMMA, a relatively smooth collagen layer is found, with small aggregates whose dimensions (height ∼4 nm; length ∼300–600 nm) roughly correspond to the ones of pentamers known to be formed 411

REVIEW


at the onset of collagen aggregation.42 On CH3 , a dense layer of short but a bit thicker collagen fibrils (height ∼4–8 nm; length ∼300–600 nm) is formed. Finally, on PS, much larger collagen structures are formed, showing several arms of ∼300 nm long and a height of ∼10 nm, associated together via an anchoring knot (height ∼20 nm). X-ray photoelectron spectroscopy (XPS) was also used to determine the surface chemical composition of the same samples. The nitrogen content can be related to the amount of collagen detected at the interface since this element is not found in the naked substrates. The nitrogen content is the lowest on PEG (4%), in agreement with the expected protein repellency of this substrate, and with the observation of isolated molecules at the interface. Note that the same PEG layer totally prevented albumin adsorption, but collagen is a very large molecule and it is more difficult to avoid its adsorption. An intermediate value (9%) is found on PMMA, pointing to the formation of a layer with a thickness lower than the probed depth (i.e., ∼10 nm). The nitrogen content measured on CH3 and PS (∼16%) is not very far from the theoretical nitrogen content of collagen (∼19% if hydrogen, an element not detected by XPS, is excluded, as calculated from the amino acid sequence of collagen). This indicates that the obtained collagen layers almost fully cover the substrate and reach a thickness at least similar to the probed depth. Starting from the knowledge of the organization of adsorbed collagen on these homogeneous substrates, two types of heterogeneous substrates were created to trigger collagen aggregation in defined domains at the interface. A first strategy (Fig. 1(a)) consisted in drawing CH3 tracks, with a width in the nanometer range, in a PEG matrix. This was achieved using a combination of electron beam lithography and silane monolayer grafting, as detailed previously.43 A second strategy (Fig. 1(b)) was based on the phase separation of PS and PMMA in thin films obtained by spin-coating a mixture of these two polymers in different proportions, as detailed previously.41 Inclusions of PS in a PMMA matrix and, conversely, inclusions of PMMA in a PS matrix were obtained. These latter patterned substrates are less ordered than the ones obtained using electron beam lithography, but they offer the advantage of a more easy preparation process, which can be applied over large surface areas. The quality of the obtained substrates was examined using a combination of AFM (shape, dimensions and distribution of patterns) and XPS (surface chemical composition). AFM used in the lateral force mode was used to evidence differences in friction, pointing to the different chemical composition of CH3 and PEG domains.40 There was indeed almost no topographical contrast for this system (see Fig. 1(a)). Images based on phase contrast obtained in tapping mode AFM were useful to identify PS and PMMA domains41 (see Fig. 1(b)). Collagen was adsorbed on these heterogeneous substrates. On CH3 /PEG substrates (Fig. 1(a)), while almost 412

Mauquoy et al.

no contrast was found on AFM height images between the CH3 tracks and the PEG matrix in absence of collagen (height ∼0.2 nm), a marked contrast was observed after collagen adsorption, demonstrating the accumulation of collagen in the CH3 areas. The height of the obtained collagen bundles was 6.5 ± 0.6 nm and was independent of track width. Collagen molecules, which are not much flexible, must be aligned along the tracks when the track width is small (50 nm) compared to the length of the molecule (∼300 nm). When track width increases to 95 nm, it can be observed that part of this alignment is lost, and some molecules escape from the tracks. Hence, the chemical contrast between tracks and matrix, and the anisotropy of the CH3 domains are at the origin of collagen assembly in defined nanotracks. On PS/PMMA substrates (Fig. 1(b)), aggregates were clearly formed starting from PS domains. When PS domains were under the form of inclusions in the PMMA matrix (PS/PMMA 35/65 substrate), each inclusion served as an anchoring point for a long and large collagen fibril, while only small aggregates were formed on the PMMA matrix, in agreement with the behavior observed on the homogeneous polymers. A similar behavior was found when the matrix was made of PS (PS/PMMA 65/35 substrate): collagen fibrils were formed in this matrix, leaving the PMMA inclusions only covered with individual molecules or smaller collagen aggregates. Note that the dimensions of the obtained collagen fibrils were similar to the ones found on pure PS when PS formed the matrix of the system, while much longer fibrils were formed when PS formed inclusions. This is attributed to the small diameter of inclusions, which is in the range of the length of collagen molecules. The nitrogen content measured by XPS on these heterogeneous PS/PMMA substrates after collagen adsorption roughly corresponds to the average, weighted by the surface coverage of each polymer, of the nitrogen content found in similar conditions on the pure polymers. Again, the chemical nature and the geometry of the PS and PMMA domains govern collagen organization at the interface. Chemical patterns at the nanometer scale are thus shown to be useful to control collagen organization and aggregation at interfaces.

4. LAYER-BY-LAYER ASSEMBLIES BASED ON COLLAGEN With a view to introduce elements of vertical organization in biointerfaces, and to later use LBL assembly to produce collagen-based nanotubes (see Section 5 of this paper), we have investigated the build-up of collagen/poly(styrene sulfonate) (PSS) films. PSS, a strong polyanion, was chosen to ensure the stability and robustness of the created layers. In a second approach, collagen/fibronectin assemblies were studied, to reproduce associations of adhesion proteins found in the ECM. J. Bionanosci. 8, 407–418, 2014

Mauquoy et al.

Fig. 2. QCM results recorded for the assembly of n-Col (plain blue line) and d-Col (red broken line) with PSS. A layer of PAH was deposited at t = 0 min, prior to further assembly. Broken arrows: addition of n-Col (blue) or d-Col (red); plain arrows: addition of PSS. Adapted with permission from [22], J. Landoulsi, et al., Soft Matter 7, 3337 (2011). © 2011, The Royal Society of Chemistry.

J. Bionanosci. 8, 407–418, 2014

d-Col allows force-distance curves to be obtained, which reveal the adhesion force between collagen and the previously deposited PAH/PSS film. Force-distance curves were recorded repeatedly using the same modified tip on different locations at the interface, as presented in Figure 3. Initially, the interaction is higher between PSS and d-Col than between PSS and n-Col. This behavior could be explained by the fact that d-Col is not anymore in the triple helical conformation typical of n-Col. This conformation may mask or maintain a larger distance between charged residues which could interact electrostatically with PSS. Additionally, d-Col is more flexible, allowing a closer contact with PSS. After a short time of interaction between the tip and the film, a marked decrease of the adhesion force is observed for d-Col, while a weaker decrease is found with n-Col. This decrease may be attributed to the attraction of PSS molecules from the PAH/PSS film by the modified tip. The Col-modified tip becomes covered with PSS desorbed from the film, while there is still PSS remaining at the extreme surface of the film. The tip and the substrate are then negatively charged and interact electrostatically in a repulsive way. The more marked decrease recorded for d-Col compared to n-Col may be related to a stronger d-Col/PSS interaction compared to the n-Col/PSS interaction, giving a higher coverage of the tip with PSS in the case of d-Col. This in line with the formation of soluble d-Col/PSS complexes and the strong d-Col adsorption on PSS as observed with QCM (see Fig. 2). 4.2. Collagen/Fibronectin Films The LbL assembly of Fn with collagen is performed in phosphate buffered saline (PBS, pH = 7.4) on polystyrene substrates, and the effect of an anchoring PEI layer and of the denaturation of collagen is examined. At this pH, the global charge of Fn is negative and collagen is theoretically positively charged, although this must be considered with caution (see Introduction section). The buffer was chosen with a view to be close to physiological conditions, thereby preventing destabilization of the film in biological medium. QCM results are presented in Figure 4(a). A decrease of frequency is observed in all cases after each Fn/collagen bilayer adsorption. It appears clearly that the PEI anchoring layer greatly improves the assembly efficiency, with a higher frequency shift observed at almost each step. The total frequency decrease observed for PEI/(Fn/n-Col)6 is about 200 Hz, while a value of about 70 Hz is obtained for (Fn/n-Col)6. The difference is thus much larger than the decrease of frequency provoked by PEI adsorption itself (∼45 Hz). This behavior was also reported in the literature for other assemblies such as PSS/PAH. It was attributed to the high degree of surface coverage by the uniform PEI anchoring layer, reducing the number of layers and the thickness necessary for growth to become independent of the substrate.45 46 In absence of the PEI anchoring layer, assembly of d-Col gives thicker films 413

REVIEW

4.1. Collagen/Poly(Styrene Sulfonate) Films Native collagen (n-Col)22 44 as well as denatured collagen (d-Col; produced by heating the n-Col solution at 90 C for 1 h)22 were assembled with PSS at pH 4.7 (acetate buffer). A first anchoring layer of poly(allylamine hydrochloride) (PAH) was used. The film build-up was monitored by quartz crystal microbalance with dissipation monitoring (QCM). Figure 2 presents the recorded frequency shifts (f ) after each step of the assembly process. Each addition of n-Col generates a higher f compared to d-Col, probably because the native molecules are longer and thus more extended in solution than denatured ones, which tend to form a coil. The very high dissipation shift (D) recorded after n-Col deposition steps confirms this hypothesis.22 Addition of PSS on n-Col gives rise to a slight gain of mass (weak decrease of frequency) and to a marked decrease of dissipation, attributed to contraction of the film. On d-Col, a positive f , i.e., a loss of hydrated mass, is observed after PSS addition, which could be due either to compaction or reorganization of the film, or to the loss of adsorbed molecules. The overall trend of the QCM results points to the successful assembly of n-Col or d-Col with PSS into multilayered films. Ellipsometry data22 confirm the efficiency of the assembly: the thickness of the dry films is approximately 18 nm for PAH/(PSS/n-Col)6 and PAH/(PSS/d-Col)6 . A decrease of the dry thickness is however observed after each addition of PSS in the case of assembly with d-Col, in line with the positive f recorded by QCM. The two species would form a soluble complex at the interface, resulting in a partial dissolution of the film. AFM images show that no collagen fibrils are formed in the multilayered films, and that films based on n-Col are denser compared to films based on d-Col.22 To further understand the mechanism of LbL assembly, the interaction of n-Col and d-Col with a PAH/PSS film was investigated using AFM in force spectroscopy mode. The use of AFM tips functionalized with n-Col or


Mauquoy et al.

REVIEW


Fig. 3. AFM in force spectroscopy mode: (a) schematic representation of a n-Col modified tip interacting with a PAH/PSS film and force-distance curves recorded as a function of time, (b) schematic representation of a d-Col modified tip interacting with a PAH/PSS film and force-distance curves recorded as a function of time. Adapted with permission from [22], J. Landoulsi, et al., Soft Matter 7, 3337 (2011). © 2011, The Royal Society of Chemistry.

and more regular build-up, with an increase of f at each adsorption step, than the one of n-Col. It is well known that the interaction of Fn is better with d-Col than with n-Col in physiological conditions, possibly because interaction sites could be masked by the triple helical conformation of n-Col.33 Finally, dissipation generally increases for collagen adsorption steps while it tends to decrease after Fn adsorption, probably because collagen is a larger and more anisotropic molecule, which tends to extend in solution, keeping the film more hydrated, while Fn adsorption would make the film stiffer. Figure 4(b) presents tapping mode AFM images of the obtained Fn/collagen films. The morphology observed 414

for 1 × 0.5 m2 images (insets) is quite similar for the three films and rather featureless, though a bit rougher in presence of PEI. On 10 × 5 m2 images, it clearly appears that the PEI anchoring layer produces a much rougher film, with aggregates of a diameter of a few hundreds of nanometers, in the range of the length of the collagen molecule. Moreover, very large fibrils are randomly observed when n-Col is used. 50 × 25 m2 images (Fig. 4(c)) confirmed the presence of large fibrils with a diameter from 0.5 m to a few m, as often observed for n-Col in physiological conditions.47 Some fibrils showed the characteristic banding pattern of collagen fibers. The absence of fibrils when collagen is denatured was expected J. Bionanosci. 8, 407–418, 2014

Mauquoy et al.


because the triple helical conformation of collagen is necessary for collagen assembly.48 Fibrils with a diameter of maximum 200 nm were previously observed in multilayered films of collagen and HA24 and fibrils with a diameter of about 10–20 nm were observed on films made of collagen and heparin or chondroitin sulfate.30 These thin fibrils are probably due to fibrillation after adsorption because collagen was deposited from an acidic solution that prevents fibrillation in solution. On the contrary, the very thick fibrils observed here are more probably resulting from fibrillation in solution, and could be at the origin of the moderate success of assembly with Fn. If many collagen molecules are involved in large fibrils, less collagen individual molecules are indeed available for adsorption in the LbL construction. Water contact angles were measured after each adsorption step as reported in Figure 5, on Fn/collagen assemblies prepared with n-Col or d-Col, and in absence or presence of a first layer of PEI. In all cases, adsorption of Fn makes the film more hydrophobic and adsorption of collagen makes the film more hydrophilic. This is a direct evidence of modification of the film after each adsorption step, pointing again to the successful deposition of the two proteins, in agreement with QCM results. The difference is more marked for d-Col compared to n-Col, in line with the frequency shifts recorded by QCM. The layers obtained in presence of PEI show a systematically higher hydrophobicity. Since this would not be attributed to PEI itself, which is not expected to impart more hydrophobicity to the films, it may arise from the film structure J. Bionanosci. 8, 407–418, 2014

and in particular from the more pronounced exposure of the underlying polystyrene substrate (polystyrene has a water contact angle of ∼90 ). Further interpretation would require to image the films by AFM at different stages of their build-up, and to compare QCM results (hydrated mass) with ellipsometry results (dry mass). At this stage of the research, there are still some doubts regarding the

Fig. 5. Evolution of the water contact angle after each step of the LbL construction (Plain line: a first layer of PEI was used and n-Col was assembled; dotted line: n-Col was used; broken line: d-Col was used). Films were made a thin polystyrene layer spin-coated on silicon. PEI was adsorbed 30 min at a concentration of 1 mg/ml; n-Col and d-Col were adsorbed 30 min at a concentration of 100 g/ml and Fn during 30 min at a concentration of 25 g/ml. After each step of adsorption, three rinsing with PBS are performed during 1 min. The temperature is fixed at 37 C. At the end of the construction, the films are rinsed 7 times with water and dried under nitrogen flow. The error bars are confidence intervals computed on 5 or 6 measurements.

415

REVIEW

Fig. 4. (a) QCM monitoring of the construction of (Fn/Col)6 films. The assembly is performed in phosphate buffered saline (PBS) on gold-covered quartz crystals spin-coated with polystyrene. CPEI = 1 mg/ml; CCol = 100 g/ml; CF n = 25 g/ml; flow = 30 l/min; T = 25 C. After each adsorption step, rinsing is performed using PBS. Plain line: a first layer of PEI was used and n-Col was assembled; dotted line: n-Col was used; broken line: d-Col was used. (b) and (c) AFM topographic images acquired in tapping mode. Films were prepared on silicon spin-coated with polystyrene. PEI is adsorbed during 30 min at a concentration of 1 mg/ml; n-Col and d-Col are adsorbed during 30 min at a concentration of 100 g/ml and Fn during 30 min at a concentration of 25 g/ml. After each step of adsorption, three rinsing with PBS are performed during 1 min. The temperature is fixed at 37 C. At the end of the construction, the films are rinsed 7 times with water and dried under a nitrogen flow.

Mauquoy et al.


DC (a)

(b)

+

REVIEW

(c)

2 μm

(e)

(d)

50 μm

Fluorescence intensity (a. u.)

(f) 5 4 3 2 1 0 4

24

48

72

96

Time (h) Fig. 6. (a) Scheme of the process used for nanotube synthesis and collection; (b) Evolution of the number of nanotubes (NTs) collected by EPD as a function of applied voltage; (c) SEM image of collagen-based nanotubes; (d) Evolution of the number of nanotubes (NTs) collected by EPD as a function of deposition time; (e) Fluorescence microscopy image of preosteoblast cells grown on a biointerface decorated with nanotubes (blue = nuclei; red = actin; green = nanotubes); (f) Evolution of the fluorescence intensity signal from the Alamar blue assay performed on preosteoblast cells cultured on ITO glass (ITO, •), on ITO glass on which collagen was simply adsorbed (ITO-COL, ), and on ITO glass decorated with collagen-based nanotubes (ITO-NTs, ). Panels (b, d, e): Adapted with permission from Ref. [50], D. M. Kalaskar, et al., Biomacromolecules 12, 4104 (2011). © 2011, American Chemical Society. Panel (f): Reprinted with permission from [51], D. M. Kalaskar, et al., Colloids Surf B 111, 134 (2013). © 2013, Elsevier.

416

J. Bionanosci. 8, 407–418, 2014

Mauquoy et al.

occurrence of a true layer-by-layer construction for collagen and fibronectin, and some questions regarding the nature of the interactions between these two proteins. We may indeed wonder if a simple progressive saturation of the interface is not taking place. The obtained results are however promising in the perspective to produce biomimetic interfaces for the control of cell behavior.

5. SURFACE-IMMOBILIZED COLLAGEN NANOTUBES

J. Bionanosci. 8, 407–418, 2014

necessary, and that collagen could be deposited first, allowing nanotubes with an outer collagen layer to be synthesized. This was required to provide signaling cues to cells. The presence of collagen at the outermost surface of these nanotubes was demonstrated by ToF-SIMS imaging.50 Electrophoretic deposition (EPD) was used to immobilize collagen-based nanotubes at the surface of indiumtin oxide (ITO)-coated glass, as illustrated in Figure 6(a) (see reference 50 for details). Briefly, the nanotube suspension (∼6.106 nanotubes/ml in DMF) was filled in a cell formed by two ITO glass slides, serving as the working and the counter electrodes, separated by a 5 mm-thick silicone spacer. A platinum wire was used as the pseudo-reference electrode. The EPD procedure was carried out for 1000s at different applied voltages, as illustrated in Figure 6(b), as well as for different durations at an applied voltage of 7.5 V, as illustrated in Figure 6(d). A clear correlation between applied voltage or deposition time and density of immobilized nanotubes was found, giving the opportunity to tune the surface coverage by collagen-based nanotubes by adjusting these parameters. The nanotubes could be observed using fluorescent microscopy owing to the introduction of a fluorescently-labeled polyelectrolyte (poly(fluorescein isothiocyanate allylamine hydrochloride); Flu-PAH) in the core of the nanotubes, which were obtained using (ncol/PSS)3 /(Flu-PAH/PSS)3 assembly. A homogeneous distribution of mostly individual nanotubes was obtained using EPD (see Fig. 6(e) and Ref. [50]). The adhesion and growth of preosteoblasts was examined on the created biointerfaces decorated with collagen-based nanotubes.51 Some cells were observed to directly interact with the nanotubes through filopodia, as shown in Figure 6(e). Changes in cell morphology were attributed to the presence of the nanotubes, while cell adhesion and growth were not modified by their presence on ITO glass, as indicated by the result of a metabolic assay (Alamar blue test, see Fig. 6(f)) and by counting labelled cells.51 It indeed appeared that cell adhesion and growth were similar on naked ITO glass, on ITO glass coated with adsorbed collagen, and on ITO glass decorated with collagen-based nanotubes (Fig. 6(f)). The nanotubes thus do not bring any obvious cytotoxicity, and these biointerfaces may then be used to trigger desired cell behaviors by filling them with signaling molecules which will be released locally to the cells.

6. CONCLUSION Collagen can be incorporated in organized biointerfaces using different approaches. Chemical patterns were drawn at the interface to trigger collagen assembly into defined areas. Collagen was included in layer-by-layer assemblies, notably with fibronectin, another important ECM component. Collagen-based nanotubes were synthesized using LbL deposition within the pores of a membrane, and 417

REVIEW

Instead of exploiting the self-assembling properties of collagen to build structured biointerfaces mimicking the ECM, an alternative approach consists in synthesizing collagen-containing nanotubes, which could then be deposited at the interface. This gives the opportunity to precisely control the dimensions of the collagen-based structures to which cells will be submitted. Moreover, the inner void of the nanotubes could be further used to encapsulate active molecules (growth or differentiation factors; drugs) which could be delivered to the cells anchored to the biointerface. The possibility to incorporate collagen in LbL assemblies (see Section 4), combined to the use of nanoporous templates in which LbL is performed, provides a strategy to elaborate collagen-based nanotubes, as depicted in Figure 6(a). Here, the nanotube dimensions are imposed by the geometry of the pores of track-etched polycarbonate membranes. The outer diameter of the nanotubes (i.e., pore diameter) can be varied between 30 nm up to a few m, and their length (i.e., template thickness) can range from 5 to 50 m. The successful assembly of PAH/(PSS/collagen)6 in templates with pores of a diameter of 200 and 500 nm was first demonstrated by gas flow porometry,22 and a multilayer thickness of 30 nm and 80 nm was determined. This is by far thicker than multilayers obtained in the same conditions on a flat substrate, as previously observed for (PAH/PSS) assemblies.49 It is believed that macromolecules can be interconnected across the pores of the template, forming a gel which fills the pores when hydrated. The thinner layer obtained in 200 nm pores may be attributed to the more limited diffusion of 300 nm-long collagen molecules in that case. The polycarbonate template was then dissolved in dichloromethane or dimethylformamide (DMF) to free the nanotubes. To demonstrate the synthesis of the nanotubes, the dissolution was performed on top of a silver membrane which was imaged subsequently by scanning electron microscopy, as shown in Figure 6(c) for nanotubes obtained in a 21 m-thick template with 200 nm-diameter pores. The dimensions of the observed nanotubes were in agreement with the template geometry. Transmission electron microscopy images also revealed the hollow structures of the formed nanotubes.44 It was further shown that the PAH layer deposited to initiate the assembly was not



the obtained nanotubes were immobilized on ITO-glass using electrophoretic deposition. The creation of collagenbased biointerfaces with tailored architectures constitutes a promising approach to better control cell-material interactions. Throughout this research, the potentialities of AFM used in its different modes (imaging and force spectroscopy) were shown to be crucial to better understand the involved mechanisms and to visualize the obtained structures. Acknowledgments: This work was supported by the Belgian Science Policy through the Interuniversity Attraction Pole Program (P7/05). The financial support of the Belgian National Foundation for Scientific Research (FNRS) is acknowledged. Sara Mauquoy thanks the FNRS for her Research Fellow position.

REVIEW

References and Notes 1. H. Lodish, A. Berk, S. L. Zipursky, P. Matsudaira, D. Baltimore, and J. Darnell, Molecular Cell Biology, 4th edn., edited by W. H. Freeman and Company, Basingstoke (1999). 2. R. O. Hynes, Cell 69, 11 (1992). 3. R. I. Freshney, Culture of Animal Cells, 3rd edn., Wiley-Liss, New York (1994). 4. C. Gonzalez-Garcia, S. R. Sousa, D. Moratal, P. Rico, M. SalmeronSanchez, Colloids Surf. B 77, 181 (2010). 5. J. T. Elliott, J. T. Woodward, A. Umarji, Y. Mei, and A. Tona, Biomaterials 28, 576 (2007). 6. Z. Keresztes, P. G. Rouxhet, C. Remacle, C. C. Dupont-Gillain, J. Biomed. Mater. Res. A 76, 223 (2006). 7. K. A. Piez, Collagen, Encyclopedia of Polymer Science and Engineering, edited J. I. Kroschwitz, J. I. Wiley, New York (1985), Vol. 3, pp. 699–727. 8. G. A. Di Lullo, S. M. Sweeney, J. Körkkö, L. Ala-Kokko, and J. D. San Antonio, J. Biol. Chem. 277, 4223 (2002). 9. C. C. Dupont-Gillain, E. Pamula, F. A. Denis, V. M. De Cupere, Y. F. Dufrêne, and P. G. Rouxhet, J. Mater. Sci.: Mater. Med. 15, 347 (2004). 10. C. C. Dupont-Gillain, E. Pamula, F. A. Denis, and P. G. Rouxhet, Prog. Colloid Polym. Sci. 128, 1 (2004). 11. C. C. Dupont-Gillain and P. G. Rouxhet, Langmuir 17, 7261 (2001). 12. I. Jacquemart, E. Pamula, V. M. De Cupere, P. G. Rouxhet, and C. C. Dupont-Gillain, J. Colloid Interface Sci. 278, 63 (2004). 13. D. L. Wilson, R. Martin, S. Hong, M. Cronin-Golomb, C. A. Mirkin, and D. L. Kaplan, Proc. Nat. Acad. Sci. USA 98, 13660 (2001). 14. K. B. Lee, J. H. Lim, and C. A. Mirkin, J. Am. Chem. Soc. 125, 5588 (2003). 15. H. W. Li, B. V. O. Muir, G. Fichet, and W. T. S. Huck, Langmuir 19, 1963 (2003). 16. D. Falconnet, D. Pasqui, S. Park, R. Eckert, H. Schift, J. Gobrecht, R. Barbucci, and M. Textor, Nano Letters 4, 1909 (2004). 17. D. Falconnet, G. Csucs, M. Grandin, and M. Textor, Biomaterials 27, 3044 (2006). 18. G. Decher, J. D. Hong, and J. Schmitt, Thin Solid Films 210–211, 831 (1992). 19. G. Decher, Science 277, 1232 (1997). 20. V. Gribova, C. Gauthier-Rouvière, C. Albigès-Rizo, R. Auzely-Velty, and C. Picart, Acta Biomaterialia 9, 6468 (2013). 21. D. T. Haynie, L. Zhang, J. S. Rudra, W. Zhao, Y. Zhong, and N. Palath, Biomacromolecules 6, 2895 (2005).

Mauquoy et al. 22. J. Landoulsi, S. Demoustier-Champagne, and C. Dupont-Gillain, Soft Matter 7, 3337 (2011). 23. J. Å. Johansson, T. Halthur, M. Herranen, L. Söderberg, U. Elofsson, and J. Hilborn, Biomacromolecules 6, 1353 (2005). 24. J. Zhang, B. Senger, D. Vautier, C. Picart, P. Schaaf, J. C. Voegel, and P. Lavalle, Biomaterials 26, 3353 (2005). 25. X. Li, Q. Luo, Y. Huang, X. Li, F. Zhang, and S. Zhao, Polym. Adv. Technol. 23, 756 (2012). 26. C. C. Chou, H. J. Zeng, and C. H. Yeh, Thin Solid Films 549, 117 (2013). 27. J. Chen, C. Chen, Z. Chen, J. Chen, Q. Li, and N. Huang, J. Biomed. Mater. Res. A 95, 341 (2010). 28. Q. Lin, J. Yan, F. Qiu, X. Song, G. Fu, and J. Ji, J. Biomed. Mater. Res. A 96, 132 (2011). 29. S. Lu, P. Zhang, X. Sun, F. Gong, S. Yang, L. Shen, Z. Huang, and C. Wang, ACS Applied Materials and Interfaces 5, 7360 (2013). 30. R. F. Mhanna, J. Vörös, and M. Zenobi-Wong, Biomacromolecules 12, 609 (2011). 31. C. Chaubaroux, E. Vrana, C. Debry, P. Schaaf, B. Senger, J. C. Voegel, Y. Haikel, C. Ringwald, J. Hemmerlé, P. Lavalle, and F. Boulmedais, Biomacromolecules 13, 2128 (2012). 32. W. Li, P. Zhao, C. Lin, X. Wen, E. Katsanevakis, D. Gero, O. Félix, and Y. Liu, Biomacromolecules 14, 2647 (2013). 33. K. C. Ingham, R. Landwehr, and J. Engel, Eur. J. Biochem. 148, 219 (1985). 34. K. Kadowaki, M. Matsusaki, and M. Akashi, Langmuir 26, 5670 (2010). 35. A. Matsuzawa, M. Matsusaki, and M. Akashi, Langmuir 29, 7362 (2013). 36. Y. Nakahara, M. Matsusaki, and M. Akashi, J. Biomater. Sci., Polym. Ed. 18, 1565 (2007). 37. M. Matsusaki, Bulletin of the Chemical Society of Japan 85, 401 (2012). 38. M. Matsusaki, K. Kadowaki, Y. Nakahara, and M. Akashi, Angewandte Chemie–Int. Ed. 46, 4689 (2007). 39. S. Shinohara, T. Kihara, S. Sakai, M. Matsusaki, M. Akashi, M. Taya, and J. Miyake, J. Biosci. Bioeng. 116, 231 (2013). 40. F. A. Denis, A. Pallandre, B. Nysten, A. M. Jonas, and C. C. DupontGillain, Small 1, 984 (2005). 41. E. M. Zuyderhoff and C. C. Dupont-Gillain, Langmuir 28, 2007 (2012). 42. K. E. Kadler, D. F. Holmes, J. A. Trotter, and J. A. Chapman, Biochem. J. 316, 1 (1996). 43. A. Pallandre, K. Glinel, A. M. Jonas, and B. Nysten, Nano Letters 4, 365 (2004). 44. J. Landoulsi, C. J. Roy, C. Dupont-Gillain, and S. DemoustierChampagne, Biomacromolecules 10, 1021 (2009). 45. M. Elzbieciak-Wodka and P. Warszyński, Electrochimica Acta 104, 348 (2013). 46. M. Kolasińska, R. Krastev, and P. Warszyński, J. Colloid Interface Sci. 305, 46 (2007). 47. Y. Li, A. Asadi, M. R. Monroe, and E. P. Douglas, Mater. Sci. Eng. C 29, 1643 (2009). 48. M. J. Beckman, K. J. Shields, and R. F. Diegelmann, Collagen, Encyclopedia of Biomaterials and Biomedical Engineering, Marcel Dekker, New-York (2004), pp. 324–334. 49. C. J. Roy, C. Dupont-Gillain, S. Demoustier-Champagne, A. M. Jonas, and J. Landoulsi, Langmuir 26, 3350 (2010). 50. D. M. Kalaskar, C. Poleunis, C. C. Dupont-Gillain, and S. Demoustier-Champagne, Biomacromolecules 12, 4104 (2011). 51. D. M. Kalaskar, S. Demoustier-Champagne, and C. C. DupontGillain, Colloids Surf B 111, 134 (2013).

Received: 20 December 2013. Revised/Accepted: 20 November 2014. 418

J. Bionanosci. 8, 407–418, 2014