Article pubs.acs.org/Biomac
Triggering Protein Adsorption on Tailored Cationic Cellulose Surfaces Tamilselvan Mohan,†,‡ Katrin Niegelhell,‡,§ Cíntia Salomaõ Pinto Zarth,∥ Rupert Kargl,⊥ Stefan Köstler,# Volker Ribitsch,†,‡ Thomas Heinze,∥ Stefan Spirk,*,‡,§,⊥ and Karin Stana-Kleinschek⊥ †
Institute for Chemistry, University of Graz, Heinrichstrasse 28, 8010 Graz, Austria Institute for Organic Chemistry and Macromolecular Chemistry, Centre of Excellence for Polysaccharide Research, Friedrich Schiller University of Jena, Humboldtstraße 10, 07743 Jena, Germany ⊥ Institute for the Engineering and Design of Materials, University of Maribor, Smetanova 17, 2000 Maribor, Slovenia § Institute for Chemistry and Technology of Materials, Graz University of Technology, Stremayrgasse 9, 8010 Graz, Austria # Joanneum Research Materials, Institute for Surface Technologies and Photonics, Franz-Pichlerstrasse 30, 8160 Weiz, Austria ∥
ABSTRACT: The equipment of cellulose ultrathin films with BSA (bovine serum albumin) via cationization of the surface by tailor-made cationic celluloses is described. In this way, matrices for controlled protein deposition are created, whereas the extent of protein affinity to these surfaces is controlled by the charge density and solubility of the tailored cationic cellulose derivative. In order to understand the impact of the cationic cellulose derivatives on the protein affinity, their interaction capacity with fluorescently labeled BSA is investigated at different concentrations and pH values. The amount of deposited material is quantified using QCM-D (quartz crystal microbalance with dissipation monitoring, wet mass) and MP-SPR (multi-parameter surface plasmon resonance, dry mass), and the mass of coupled water is evaluated by combination of QCM-D and SPR data. It turns out that adsorption can be tuned over a wide range (0.6−3.9 mg dry mass m−2) depending on the used conditions for adsorption and the type of employed cationic cellulose. After evaluation of protein adsorption, patterned cellulose thin films have been prepared and the cationic celluloses were adsorbed in a similar fashion as in the QCM-D and SPR experiments. Onto these cationic surfaces, fluorescently labeled BSA in different concentrations is deposited by an automatized spotting apparatus and a correlation between the amount of the deposited protein and the fluorescence intensity is established.
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INTRODUCTION Control over protein adsorption on surfaces is a key parameter in the design of advanced materials in a variety of technological fields. While in many areas protein adsorption is undesired (e.g., antifouling surfaces) concomitant with performance losses of the materials, in medical implants and tissue engineering a controlled deposition of proteins often allows for a faster assimilation and healing process, and a recovery with minor complications.1,2 In this context, cationic polysaccharides such as chitins (partially deacetylated) and chitosans have shown to give promising results in terms of nerve regeneration, bone regeneration, and wound healing.3,4 In all these applications, it is assumed that the ability of chitins and chitosans to interact with membrane proteins such as integrins, and laminins is responsible for the excellent performance of the materials.5−7 Interestingly, the interaction of proteins with cationic polysaccharide surfaces is not fully explored up to date. Although there are some studies available that determine the total amount of adsorbed proteins on these implants,8−10 detailed studies on real time adsorption kinetics and unspecific protein interaction are investigated to a minor extent due to the lack of suitable methods for online monitoring. In addition, all © XXXX American Chemical Society
of these studies use only chitins and chitosans as model system and other cationic polysaccharides such as cationic starch11 and cellulose have not attracted significant interest so far. Particularly, cationic cellulose derivatives are an interesting class of compounds due to the availability of the raw material cellulose and the easy modification procedures in order to achieve cationization, which allows to tailor the material properties in order to realize the envisaged application.12−16 In contrast to bulk samples, (protein) adsorption on thin films can be investigated with a variety of methods such as QCM-D and SPR, to mention the two most important ones.17−19 Both methods have in common that the adsorption process can be monitored in situ, that is, a solution containing the adsorbate is pumped via a microfluidic system over the surface of interest and the response is shown in real time. In terms of protein adsorption on cellulosic surfaces, one of the benchmark studies was published by the Rojas group in 2011.20 This paper focuses on the adsorption of anionic (carboxymethyl cellulose) and Received: July 9, 2014 Revised: September 16, 2014
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at least controlled. Therefore, the main motivation of this work is to investigate how and to which extent cationic cellulose derivatives can be used in order to achieve such an enhanced protein adsorption onto cellulosic surfaces.
cationic (chitosan) polysaccharides on cellulose and subsequent adsorption of proteins having different charge densities. The authors nicely demonstrate that unspecific protein adsorption on chitosan for instance is much higher than on neat cellulose, which is known for a rather low unspecific protein adsorption. Very recently, we adapted this concept and showed that medical grade N,N,N-trimethyl chitosans can be immobilized on cellulose in a similar fashion and potentially used as matrices in order to produce patterned slides that are able to detect adsorption from 12 pM solutions of BSA on the pads.21 These results inspired us to extend the gained knowledge to cationic cellulose derivatives adsorbed on cellulose since the charge density of the cationic layer (which is a decisive factor in protein adsorption) can be tuned very easily by altering the degree of substitution (DS). One might expect that the adsorption of a positively charged polymer (e.g., cationic celluloses) onto a negatively charged surface (e.g., cellulose surfaces due to the presence of hemicelluloses and oxidized groups within the chain for instance) is more pronounced if the charge densities of the oppositely charged polymers are very high. However, the charge density is not the only decisive factor for the adsorption of a charged polymer onto an oppositely charged surface.22,23 A key parameter is the solubility of the polymer as well. A polymer that is well soluble will adsorb to a lower extent than a similar polymer that features sparing solubility. Moreover, the conformation of the polymer is influenced by factors such as pH value and electrolyte concentration. While at low ionic strengths charged polymers usually adopt a flat and strained conformation, minimizing electrostatic interactions between charged polymer segments, the presence of electrolytes compensates for the charges between the charged segments of the polymer leading to a coiled-like conformation. Other effects include interactions by conformational stabilization of the adsorbate by the surface. In case of polysaccharides, such specific interactions have been postulated for xyloglucans,24 carboxymethyl cellulose,25,26 and a cationic cellulose derivative.27 This stabilization potentially plays an important role when the polymer enters the hydration sphere of the cellulose film in the initial phase of adsorption. Those compounds having a higher degree of substitution impose a steric pressure onto the system, decreasing conformational stabilization, which results finally in a lower adsorption onto the surface. In all cases, the adsorbed layer consists of adsorbed polymer, electrolyte, and solvent, which are embedded inside the layer.22 In this study, the adsorption behavior of two cationic cellulose derivatives on ultrathin cellulose supports is reported. Onto these cationized surfaces, unspecific protein adsorption is studied by QCM-D and MP-SPR using fluorescently labeled BSA. In the last part of this paper, the applicability of the chosen approach is demonstrated by preparing patterned cationic cellulose films, which are loaded with different amounts of fluorescently labeled BSA. It can be clearly seen that the fluorescence intensity is proportional to the loading with proteins, and adsorption from solutions in the pM range is detectable. This approach should be the basis for future developments, where the unspecific adsorption of BSA on cationic polysaccharide surfaces is exploited in the design of biosensor arrays as well as for cell growth scaffolds where BSA influences cell adhesion. In these arrays, the BSA can act as blocking agent, which can be replaced by larger proteins (e.g., antibodies). In order to use BSA as blocking agent, for instance, unspecific protein surface interactions must be maximized and
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EXPERIMENTAL SECTION
Materials. Trimethylsilyl cellulose (TMSC, DSTMS: 2.8, derived from Avicel PH-101) was purchased from Thüringisches Institut für Textil- and Kunststoff-Forschung e.V. (TITK), Germany and used as starting material for cellulose film preparation. Toluene (99.9%), sodium acetate (anhydrous), disodium phosphate heptahydrate (Na2HPO4·7 H2O), sodium dihydrogen phosphate monohydrate (NaH2PO4·H2O), fluorescein isothiocyanate-bovine serum albumin (FITC-BSA), and tetramethylrhodamine isothiocyanate-bovine serum albumin (TRITC-BSA) were purchased from Sigma-Aldrich, Austria, and used as received. Cellulose-4-[N,N,N-trimethylammonium]butyrate chlorides with two different degrees of substitutions (CMABCL/DS = 0.21; CMABCH/DS = 0.675) were synthesized according to the protocol published elsewhere.28 Cyclic olefin polymer slides (COP, Zeonor1060R slides) were generously provided by SonyDADC Austria and used as received. QCM-D sensors (QSX301) were purchased from LOT-Oriel (Germany). Milli-Q water from a Millipore water purification system (Millipore, U.S.A.; resistivity = 18.2 Ω−1 cm−1) was used for contact angle and QCM-D investigations. Quartz Crystal Microbalance with Dissipation (QCM-D). A QCM-D instrument (model E4) from Q-Sense, Gothenburg, Sweden was used. The instrument simultaneously measures changes in the resonance frequency (Δf) and energy dissipation (ΔD) when the mass of an oscillating piezoelectric crystal changes upon increase/decrease in the mass of the crystal surface due to the added/deduced mass. Dissipation refers to the frictional losses that lead to damping of the oscillation depending on the viscoelastic properties of the material. For a rigid adsorbed layer that is fully coupled to the oscillation of the crystal, Δf n is given by the Sauerbrey equation29 1 Δm = C
Δfn n
(1)
where Δf n is the observed frequency shift, C is the Sauerbrey constant (−0.177 mg Hz−1 m−2 for a 5 MHz crystal), n is the overtone number (n = 1, 3, 5, etc.), and Δm is the change in mass of the crystal due to the adsorbed layer. The mass of a soft (i.e., viscoelastic) film is not fully coupled to the oscillation and the Sauerbrey relation is not valid since energy is dissipated in the film during the oscillation. The damping (or dissipation; D) is defined as D=
Ediss 2πEstor
(2)
where Ediss is the energy dissipated and Estor is the total energy stored in the oscillator during one oscillation cycle. Cellulose Film Preparation. Prior to spin coating of cellulose film, the QCM-D sensors were soaked into a mixture of H2O/H2O2 (30 wt %)/NH4OH (25 wt %; 5:1:1; v/v/v) for 10 min at 70 °C, then immersed in a “piranha” solution containing H2O2 (30 wt %)/H2SO4 (98 wt %; 1:3; v/v) for 60 s, and then rinsed with MQ-water and finally blow dried with N2 gas. For spin coating of TMSC, 50 μL of TMSC solution (1% (w/v), dissolved in toluene by heating to 60 °C, followed by cooling down to room temperature, filtered using 5 μm PTFE syringe filter) was deposited onto the static substrate, rotated for 60 s at a spinning speed of 4000 rpm and an acceleration of 2500 rpm s−1. For converting TMSC into pure cellulose, the TMSC coated sensors were placed into a polystyrene Petri dish (5 cm in diameter) containing 3 mL of 10 wt % hydrochloric acid (HCl). The dish was covered with its cap and the TMSC films were exposed to the vapors of HCl for 15 min. This procedure is designated as “regeneration” in this study. The regeneration of cellulose from TMSC-coated films was verified by water contact angle, XPS, and ATR-IR measurements as reported elsewhere.30,31 Static water contact angles of TMSC and regenerated cellulose films were determined to be B
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97 ± 1° and 33 ± 1°, respectively. The film thickness of regenerated cellulose was 21.0 ± 0.3 nm. Working Solution Preparation and Accomplishment of QCM-D Measurements. Preparation of CMABC Samples. CMABCL and CMABCH samples (c = 1 mg mL−1) were dissolved in pure MQ-water. The ionic strength of the sample was adjusted to 150 mM with sodium chloride (NaCl) electrolyte at pH 7. The pH of the solution was adjusted by using either 0.1 M NaOH or 0.1 M HCl. All solutions were stirred overnight at room temperature and filtered using 5 μm PTFE syringe filters. Preparation of Protein Samples. FITC-BSA was dissolved (1 mg mL−1) in a 10 mM buffer at pH 5 (sodium acetate/acetic acid buffer), 6, and 7 (phosphate buffer). The pH of the buffer solution was adjusted using either glacial acetic acid (pH 5) or 0.1 M NaOH (pH 6 and 7). The ionic strength of the three buffer solutions was adjusted to 100 mM with NaCl electrolyte. Immobilization of Protein on CMABC-Coated Cellulose Surface. Cellulose-coated QCM sensors were mounted in the QCM flow cell and equilibrated first with water for 30 min and then to a 150 mM NaCl solution for 30 min. After equilibration of the films, aqueous solutions of CMABC (c = 1 mg mL−1) at an ionic strength of 150 mM NaCl were pumped over the sensors at a flow rate of 0.1 mL min−1 for 60 min. After this, the CMABC solution was exchanged with NaCl (30 min.) and with water (45 min). After equilibration of the CMABCcoated cellulose films in water and buffer solution at the corresponding pH value, FITC-BSA was introduced into the QCM flow cell. The different FITC-BSA solutions were pumped through the QCM-D cell for 30 min followed by corresponding buffer solution for 30 min. The flow rate was kept at 0.1 mL min−1 throughout all experiments. The temperature was kept 21 ± 0.1 °C for all experiments. All adsorption experiments have been performed in three parallel and a mean value and standard deviation of third overtone dissipation and frequency was calculated. Viscoelastic Modeling. The viscoelastic Voigt model was applied for calculating the adsorbed mass (ΓQCM), film thickness (hf), viscosity (ηf), and elastic shear modulus (μf) of the CMABC- and FITC-BSAcoated layers. In this model, the adsorbed layer was treated as a viscoelastic layer between the quartz crystal and a semi-infinite Newtonian liquid layer. More details on the Voigt modeling can be found elsewhere.17,32 For data evaluation or fitting the different overtones (n = 3, 5, 7, 9 and 11) of frequency and dissipation were used. All calculations were carried out using the software package QTools 3.0.12 (Q-Sense). The fitting parameters used for the modeling are viscosity, from 1 × 10−4 to 0.01 N·s·m−2; elastic shear modulus, from 1 × 104 to 1 × 108 N·m−2; and thickness, from 1 × 10−10 to 1 × 10−6 m. It is worth noting that the values of hf and ρf were not independent variables. In order to calculate the effective thickness and adsorbed mass (eq 3), the density ρf values were varied between 1000 and 1180 kg m−3. It turned out that no mass change for CMABC-coated layer occurred by changing the density value and therefore the density (ρf) of 1000 kg m−3 was used for all calculation (eq 3).
ΓQCM = hf ρf
HCl. The same spin coating and regeneration procedure as for the QCM-D experiments were used. Prior to adsorption of the cationic cellulose derivatives on these cellulose films, the cellulose films were allowed to equilibrate in Milli-Q water, and then in acetate buffer (10 mM, 0.1 M NaCl) at the corresponding pH value intended for protein adsorption. The adsorption procedure of the cationic cellulose derivatives onto the cellulose thin films was accomplished in the same manner as for the QCM-D experiments. The adsorption of FITC-BSA was performed in acetate buffer at pH 5, 6, and 7 at a BSA concentration of 1.0 mg mL−1 under flow conditions (flow rate 0.1 mL min−1). After 30 min adsorption was stopped and the films were rinsed with buffer in order to remove loosely bound material. Protein adsorption onto the cationic cellulose surfaces was quantified according to eq 4, which considers the dependence of the angular response of the surface plasmon resonance in dependence of the refractive index increment (dn/dc) of the adsorbing layer. Γ=
ΔΘ × k × d p (4)
dn/dc
For thin layers ( pH 6 > pH 7, CMABCH > CMABCL at the same pH value). Fluorescence Microscopy. In order to evaluate whether the FITC-BSA is homogeneously distributed over the surfaces,
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CONCLUSION In this paper, a simple approach to tune protein adsorption on cellulosic materials is established. It was demonstrated that the incorporation of charged species on cellulose thin films, namely, tailored cationic cellulose derivative allow for controlling protein adsorption significantly over a range from 0.6 to 3.9 mg m−2 (dry mass). Particularly at pH 5, this adsorption was accompanied by partial denaturation of the protein at the interface, as demonstrated by unusually high contact angles of the protein layer. After evaluation of the interaction mechanism, which mainly takes place via electrostatic and hydrophobic interactions, a simple approach was demonstrated as to how to prepare patterned slides that could be used as substrates in the detection of proteins. In a proof of principle, we showed that fluorescent labeled proteins can be easily detected at loadings ranging from the pM to μM regime. Although this approach is not suitable to act as (bio)sensor, since it lacks selectivity, for instance, we aim at using the adsorbed BSA as blocking agent in order to achieve selective functionalization of cellulosic surfaces by functional proteins (e.g., antibodies), and studies dealing with this topic are underway.
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AUTHOR INFORMATION
Corresponding Author
*E-mail:
[email protected]. Tel.: +43 316 873 32284. Notes ‡
These authors are members of NAWI Graz Members of the European Polysaccharide Network of Excellence (EPNOE). The authors declare no competing financial interest. I
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