Bioinspired porous membranes containing polymer nanoparticles for wound healing Ana M. Ferreira,1 Clara Mattu,1 Elia Ranzato,2 Gianluca Ciardelli1,3 1

Department of Mechanical and Aerospace Engineering, Politecnico di Torino, Corso Duca degli Abruzzi 24 10129 Turin, Italy 2 Dipartimento di Scienze e Innovazione Tecnologica, University of Piemonte Orientale, Viale Teresa Michel, 11 15121 Alessandria, Italy 3 CNR - Consiglio Nazionale delle Ricerche, Istituto per i Processi Chimico Fisici (IPCF-CNR), UOS di Pisa, Area della Ricerca, Via G. Moruzzi 1, 56124 Pisa, Italy Received 5 November 2013; revised 23 January 2014; accepted 11 February 2014 Published online in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/jbm.a.35121 Abstract: Skin damages covering a surface larger than 4 cm2 require a regenerative strategy based on the use of appropriate wound dressing supports to facilitate the rapid tissue replacement and efficient self-healing of the lost or damaged tissue. In the present work, a novel biomimetic approach is proposed for the design of a therapeutic porous construct made of poly(L-lactic acid) (PLLA) fabricated by thermally induced phase separation (TIPS). Biomimicry of ECM was achieved by immobilization of type I collagen through a twostep plasma treatment for wound healing. Anti-inflammatory (indomethacin)-containing polymeric nanoparticles (nps) were loaded within the porous membranes in order to mini-

mize undesired cell response caused by post-operative inflammation. The biological response to the scaffold was analyzed by using human keratinocytes cell cultures. In this work, a promising biomimetic construct for wound healing and soft tissue regeneration with drug-release properties was fabricated since it shows (i) proper porosity, pore size, and mechanical properties, (ii) biomimicry of ECM, and (iii) theraC 2014 Wiley Periodicals, Inc. J Biomed Mater Res peutic potential. V Part A: 00A:000–000, 2014.

Key Words: collagen, nanoparticles, plasma, porous membrane, wound healing

How to cite this article: Ferreira AM, Mattu C, Ranzato E, Ciardelli G. 2014. Bioinspired porous membranes containing polymer nanoparticles for wound healing. J Biomed Mater Res Part A 2014:00A:000–000.

INTRODUCTION

Skin is a complex tissue and an efficient barrier against external factors, such as UV radiation and pathogenic microbial agents. Moreover, it prevents the substantial loss of body fluids and plays a significant role in thermoregulation and immune defense. Skin loss can occur for many reasons, including disorders, acute trauma, chronic wounds, and thermal trauma. Chronic wounds represent a growing health and economic problem due to the high morbidity, risk of amputations, and a heavy socioeconomic burden.1 Thermal traumas involving substantial areas of the body, often cannot be properly or completely regenerated resulting in the formation of disabling and disfiguring hypertrophic scars.2 In addition, wounds that for any reason fail to heal may impact negatively on the patient’s quality of life and result in higher cost to health services. The “clinical gold-standard” when treating skin damages is the transplantation of autologous tissue from uninjured sites. Unfortunately, this strategy cannot be applied for large, extended damages, due to the lack of donor sites. Larger tissue defects or hard-to heal wounds require appropriate advanced wound therapies in order to promote a faster and

more complete healing of tissue without scar formation.3 Engineered skin substitutes represent an alternative to autologous transplantation, as well as a prospective of advanced therapy in treating acute and chronic skin wounds.4 Among them, scaffolds based on different biomaterials represent a successful approach to regenerate cutaneous tissue since they provide structural support to cell functions, including adhesion, proliferation and differentiation, and subsequent development of new tissue without scar formation.5–8 The design of scaffolds with the capability to mimic important features of the extracellular matrix (ECM) architecture and to create an optimized cellular microenvironment, influences the functionality and regenerative potential of the construct for tissue repair.9 Three key features influence the in vivo functionality of the scaffold: (a) the nature of the material, (b) the surface topography, and (c) the architecture.10 The scaffolding material must be biocompatible to avoid rejection, and promote cell growth and colonization. The chosen biomaterial should not cause inflammatory reactions or the formation of a fibrous capsule11 and be biodegradable with kinetics that are compatible with the time required for the tissue to regenerate. Moreover, the degradation products must not be

Correspondence to: A. M. Ferreira; e-mail: [email protected] Contract grant sponsors: “Manunet—BIODRESS” Project; Lagrange CRT

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toxic.12 The scaffold must interact with the cellular components of the host tissue by providing specific biochemical signals in order to support cellular ingrowth and favor specific cellular responses to inhibit scar formation and accelerate growth of new normal tissue. These characteristics can be obtained by means of surface functionalization and insertion of appropriate biomolecules present in the ECM of the targeted tissue.13 These topographical and chemical modifications allow proteins to carry out their normal activities without transposing danger signals, while the insertion of specific biomolecules may induce the desired cellular response.14 The optimal scaffold for cutaneous tissue regeneration should possess a porous architecture, with a pore size in the order of dozen to a few hundred mm and a well interconnected structure to facilitate the migration of cells, make the transport of nutrients and metabolites more efficient and provide the necessary space for the formation of new tissue.15,16 Moreover, the mechanical properties of the scaffold should be similar to those of the host tissue and must allow good handle-ability during the surgical insertion. The scope of this work is to design and characterize a biomimetic, porous construct for wound healing able to fulfill the current clinical need of multifunctional scaffolds to repair large tissue defects in skin tissue engineering. The combination of topographical and morphological cues was achieved in an all-in-one construct, based on a poly L-lactid acid (PLLA) scaffold, obtained through the TIPS technique, surface-modified by immobilization of biochemical signals such as type I collagen via the plasma technique. In addition, anti-inflammatory drug (Indomethacin, IND)-loaded nps based on biodegradable polymers were introduced within the bio-functionalized membrane with the aim to enhance cutaneous tissue healing by preventing postoperative infections and inflammation. Nanocarriers are known to improve the control over the release kinetics of encapsulated drugs, whilst protecting it from degradation and prolonging its activity in vivo. IND is a nonsteroidal anti-inflammatory drug, which inhibits the efflux of prostaglandins following injury, thus reducing dermal ischemia and edema formation.17 The biomimetic, multifunctional system was then characterized in terms of mechanical and chemical properties, morphology, and by assessing keratinocytes viability and growth in vitro. EXPERIMENTAL SECTION

Materials Soluble collagen type I was isolated and purified following the methodology described by Ferreira et al.18 Sodium heparin was purchased from Polysciences, Inc. (Milan, average Mn: 100M USP Units). Poly(L-lactide) (PLLA) was purchased from Purac Biochem (Milan, average Mn 100,000 Da). Bidistilled water was used in all experiments. Indomethacin, n-boc-serinol, 1,6 hexamethylene diisocyanate (HDI), PCL diol (Mn 2000 Da) were purchased from Sigma–Aldrich (Milan, Italy). All solvents were of analytical grade. Preparation of PLLA porous membranes and thin films PLLA films were prepared by solvent casting technique on a glass mold. PLLA was dissolved homogenously in chloro-

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form (0.03 g mL21) and the solvent was removed by evaporation at room temperature for 24 h. Films were cut into rectangular samples of 1 cm2. Porous PLLA membranes were prepared via thermally induced phase separation technique (TIPS) by dissolving PLLA in a mixture of 1,4-dioxane (Sigma–Aldrich) (solvent) and bi-distilled water (nonsolvent) at 70 C. In detail, PLLA (5 wt %) was dissolved in a mixture of nonsolvent (water, 5 vol %) in 1,4 dioxane and poured into metallic containers in order to favor heat transfer and cooled for 2 h at room temperature. The containers were frozen at 220 C overnight. The solvent was removed by immersing the frozen solution in a cooled bath of 90 vol % of ethanol (Sigma–Aldrich) at 220 C for 3 days, frequently changing the bath solution during the day, in order to obtain a porous structure. The scaffold obtained was dried at room temperature to evaporate all remaining ethanol. Control membranes were prepared by dissolving PLLA (5 wt %) in pure 1,4 dioxane (in absence of nonsolvent). Subsequently, polymer solution was transferred into metallic containers, cooled for 2 h at room temperature and frozen at 220 C overnight. The solvent was removed following the steps previously described. Functionalization with type I collagen by a two-step plasma treatment Plasma parameters were firstly optimized on PLLA thin films using Ar plasma at different power outputs (50, 100, and 150 W) for 5 min, followed by 15-min treatment with acrylic acid (AAc) monomer vapors, maintaining the pressure of the plasma reactor chamber at 0.06 mBar. The Ar plasma was applied using a Diemer Electronic, Pico Low Pressure Plasma System in order to activate the surface by free radical formation for further grafting of AAc molecules. Finally, the residual monomers and homopolymers were removed by washing with distilled water. For PLLA porous membranes functionalization with AAc, samples of 2-mm thick and 1 cm2 of surface area were washed with EtOH/ H2O (50 vol %) and dried at room temperature, prior to Ar plasma treatment at 50 W. The carboxyl groups concentration after AAc grafting/polymerization was determined by the colorimetric titration method using toluidine blue O (TBO).19 Samples were immersed into 0.05 mM TBO aqueous solution at pH 10 during 12 h at room temperature in order to allow the formation of ionic complexes between the COOH groups and the cationic dye. Sample surfaces were rinsed with 0.1 mM NaOH solution at pH 10 to remove excess and unbound TBO molecules. TBO bound to COOH groups present on PLLA-AAc surfaces was released in 1 mL 50 vol % of acetic acid solution for 10 min and the absorbance was measured by UV–vis spectrophotometer (PerkinElmer, Lambda 25) at 632 nm. The amount of COOH was calculated using a calibration curved built-up at different concentrations of TBO in 50% of acetic acid in the same conditions, assuming that 1 mol TBO corresponds exactly to 1 mol carboxyl groups.20,21 The colorimetric protocol was performed by triplicate. The chemical functionalization with type I collagen was carried out after plasma treatment by immersing the

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samples in 0.2 mM aqueous solution (pH 5) of 1-ethyl-3-(3dimethylaminopropyl) carbodiimide (EDC), in the presence of 0.12 mM of N-hydroxysuccinimide (NHS) at 4 C. The NHS facilitates the amidation on the acrylic acid-grafted surface, by the formation of intermediate succinimidyl ester groups. The activated samples were washed with distilled water and then immersed into a type I collagen solution (1 mg mL21) in acetic acid (0.5M, pH 5) at 4 C, thus allowing the reaction between the amine groups of collagen and the active succinimidyl ester groups, forming stable amide linkages on the material surface. Process parameters for the COOH groups activation though carbodiimide and N-hydroxysuccinimide-mediated chemistry and the further functionalization with biomolecules on substrates surfaces, including collagen, may change according to the kind of substrate and intrinsic polymer properties, such as molecular weight, crystallinity, and so forth. Thus, in order to obtain the most suitable process conditions for the activation step with EDC/ NHS and subsequent superficial grafting with collagen type I, two different combinations were evaluated on PLLA thin films: (i) 1 h of surface activation with EDC/NHS followed by 1-h immersion in collagen solution; (ii) 1 h of surface activation with EDC/NHS followed by 20-h coupling with collagen and (iii) vice versa, 20-h activation in EDC/ NHS and 1 h coupling with collagen. The amount of immobilized collagen was determined by BiocolorTM Sirius Redbased colorimetric micro assay.22 The colorimetric protocol was performed in triplicates. For porous constructs, optical NIKON microscope and stereo zoom microscope smz1500 with DS CAMERA HEAD DS-Fi1 was used to visualize the stained ACOOH groups and collagen molecules as well as the homogeneity of the functionalization. Carboxylic groups and collagen were stained blue and red on a white background on the PLLA porous membranes, respectively. Mechanical tests on porous membranes Porous membranes were mechanically tested using uniaxial tensile testing (MTS Qtest/10 Elite Controller) in order to assess the Young modulus, the maximum load before failure and the maximum elongation at break. Rectangular membranes with a cross-section of 3 mm were tested in triplicates, by using a load cell of 500 N at a set load speed of 1 mm min21. The Young’s modulus E [MPa], the maximum elongation at break (%) emax and the maximum load r max (MPa) values were determined from the stress–strain curves. Hooke’s law was applied to the linear portion of each curve in order to calculate the elastic modulus. PLLA control membrane was also mechanically tested. Contact angle measurements The wettability of PLLA samples before and after plasma treatment and collagen grafting was evaluated by contact angle measurements using CAM 200 KSV Instrument, equipped with Tetha software. Static contact angle measurements were carried-out by depositing a 3 mL drop of bidistilled water at room temperature on the sample’s surface. Three different points were tested for each sample. All measurements were performed in triplicates.

Scanning electron microscopy (SEM) The surface morphology of treated and non-treated samples was observed by LEO 1430VP SEM Equipment, LeoCo. LTD. Prior to SEM examination, sample surfaces were coated with a conductive gold layer. Nanoparticles preparation and characterization Nps were prepared with a commercial polymer, PCL (average Mn 43,000–50,000 Da, Polysciences) and a novel polyesterurethane NS-HC2000, in the following referred as PU. The acronym represents the components of the material: NS is the chain extender (n-BOC-serinol), H corresponds to the diisocyanate (HDI) and C2000 indicates the macrodiol (PCL-diol with a molecular weight of 2000 g mol21). The polymer was synthesized according to a two-step polymerization reaction, as reported by Mattu et al.23 IND-loaded nps (5 wt %) were prepared by a modified single emulsion solvent evaporation method to obtain nps of small size ranging between 150 and 250 nm, as already described.24 Empty nps were also prepared as controls. The particle size and polydispersity index (PDI) analysis were performed using dynamic laser light scattering (Nanoseries, Nano-ZS; Malvern Instruments, UK). The dispersion of nps was diluted with ultrapure water and completely dispersed before measurement. The measured values were averaged by five runs. Surface charge was determined by electrophoretic light scattering (Nanoseries, Nano-ZS; Malvern Instruments, UK) at room temperature in ultrapure water. The nps morphology was assessed by SEM microscopy (LEO 1430VP SEM Equipment, LeoCo. LTD). The sample was prepared by placing the particles onto aluminum substrates followed by a coating with a thin layer of gold. A working distance of 10 mm and a voltage of 20 kV were applied. Indomethacin encapsulation efficiency (EE) and release profiles were determined by UV spectroscopy (Perkin Elmer Lambda 25) at 320 nm. Briefly, 10 mg of lyophilized nps fabricated with the two polymers (PCL and PU) were dissolved in ethyl acetate. The solvent was then completely evaporated, the drug was recovered with a solution of ethanol and water (3:1) and analyzed by UV. EE was calculated using the following formula [Eq. (1)]: EEð%Þ5

DM  100 DT

(1)

where DM is the amount of drug contained in 10 mg of nps, measured by UV, and DT is the theoretical amount of drug that is expected in 10 mg of nps (5 wt %). The drug release profile was determined by dispersing nps into 1 mL of distilled water, and by incubating the suspension at 37 C under gentle and continuous stirring in order to simulate the physiological environment in an IKA KS 4000 incubator. The total volume of incubated nps was centrifuged and the supernatant analyzed by UV. The release volume was renewed by adding 1 mL of fresh deionized water. The drug release profile was measured after 1, 2, and 3 h of incubation, followed by daily measures for up to 5 days. All measurements were performed in triplicates and the results were expressed as mean 6 standard deviation. After collagen immobilization onto the surface of the porous membranes, nps were deposited by immersing the

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membranes in aqueous solution in which the nps were dispersed at a theoretical loading of 30 mg of IND. The negatively charged nps are expected to electrostatically interact with the positively charged collagen deposited on the surface, thus remaining entrapped inside the construct. After particles loading the construct were gently rinsed with distilled water to remove excess of nanoparticles and drug release profiles were analyzed. Preliminary cellular assay HaCaT cells are immortalized human skin keratinocytes, which can differentiate under appropriate experimental conditions.25 Cells were maintained at 37 C, 5% CO2, in DMEM supplemented with 10 vol % fetal bovine serum (FBS) and 1 vol % antibiotic mixture. Cell viability was assessed by staining cells with crystal violet dye. Briefly, cells were seeded on porous membranes and grown for 1, 3, and 7 days. Thereafter, the medium was removed; cells were gently washed with PBS, stained for 10 min with 0.5 wt % crystal violet in 145 mmol L21 NaCl, 0.5 vol % formal saline, 50 vol % ethanol, and washed thrice with water. Crystal violet was eluted from cells with 33% acetic acid and the absorption of the supernatants was measured at 540 nm in a plate reader (Infinite 200 Pro, Tecan, Wien, Austria). After 14 days, porous membranes were fixed in 10% buffered neutral formalin, processed with paraffin and sectioned at 2 mm. Filtered, aqueous solutions of toluidine blue were used for staining. Initial experiments performed by varying dye concentrations and staining times were used to establish the concentrations and times that gave satisfactory results in terms of cell distribution. Paraffin-embedded tissue sections were stained in 0.01 wt % toluidine blue for 2 min. Digital pictures were taken of toluidine blue-stained samples at 4003 magnification. Statistical analysis The statistical analysis in terms of significant difference of collagen biomolecules quantification and cell viability was performed for n 5 3 samples using the ANOVA one-way analysis of variance of the Origin-7 software. The post hoc multiple comparisons between the independent groups were evaluated using the Turkey’s test.

ence of AAc groups on the surface.22 The peak at 1665 cm21 was not detected in the spectrum of the samples functionalized at 150 W, confirming the lower COOH grafting detected by TBO assay on the surface of these samples [Fig. 1(C)]. At a macroscopic examination, films assumed a darker coloration as the power of plasma treatment increased, indicating that a certain degree of degradation occurred during the functionalization process. SEM micrographs [Fig. 1(B)] confirm this observation, since the surface of the sample treated at higher plasma power (150 W) showed signs of degradation, bubbles and surface brittleness. Based on these results, the treatment at 50 W was selected for further biofunctionalization with type I collagen. Contact angle measurements of plasma-treated PLLA films and the density of grafted carboxyl groups quantified by TBO-colorimetric method are summarized in Figure 1(C). As expected, the wettability of PLLA films after AAc functionalization increased compared with the untreated controls, due to the presence of grafted carboxyl groups.26 Increasing the treatment power resulted in an increase of the surface wettability, while the surface density of COOH groups decreased with the power increasing from 50 to 150 W. The amount of carboxyl groups on the surface of samples treated at 150 W was found to be 1.8 6 0.3 mg cm22, significantly lower than COOH density for of samples treated at 100 and 50 W which, found to be at 3.4 6 0.3 and 3.8 6 0.9 mg cm22, respectively (differences not statistically significant). Surface functionalization with type I collagen The effect of the immersion time in the EDC-NHS solution to activate the surface-exposed carboxyl groups,27 and the effect of the duration of the incubation with collagen for surface functionalization, were evaluated on PLLA films by Sirius-Red colorimetric assay. The amount of grafted collagen, quantified by Sirius-Red colorimetric assay, increased with increasing immersion time in the collagen solution, as shown in Figure 2. Thus, shorter lapses of treatment time for COO2 activation with EDC/NHS, combined with longer lapses of immersion in collagen solution resulted in an enhanced biomolecule coupling. Based on these results, 1-h activation with EDC-NHS followed by 20-h immersion in collagen solution was selected for further functionalization of PLLA porous membranes.

RESULTS AND DISCUSSION

Optimizing plasma experimental setup Three different plasma power outputs were tested for the grafting of AAc on PLLA thin films: 50, 100, and 150 W. Differences in hydrophilicity, carboxylic groups density and topography associated with the surface treatment were evaluated. Figure 1 summarizes the properties of plasma-treated PLLA films. IR-ATR spectra of PLLA films after plasma treatment [Fig. 1(A)] evidenced the presence of two new peaks for samples functionalized at 50 and 100 W. The peak at 2923 cm21 was attributed to the stretching of OH bonds of the carboxylic acids, characterized by a very broad band between 3300 and 2500 cm21, centered at about 3000 cm21. The second peak, at about 1665 cm21, was attributed to the stretching vibrations of C@O of carboxylic acids, detectable at around 1700 cm21. These two new peaks are evidence of the pres-

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Plasma functionalization with AAc and collagen grafting on porous constructs AAc-functionalized membrane treated by plasma at 50 W and untreated control membranes were immersed into a TBO solution for 4 h to allow the TBO molecules to bind to carboxyl groups exposed on the surface. The functionalized samples were unifomly stained in blue after the incubation in the TBO solution, evidencing the homogeneous distribution of carboxyl groups on the surface of the construct [Fig. 3(A)]. No blue staining was observed on the untreated sample (control), confirming the successful AAc functionalization. After EDC-mediated collagen grafting, samples were analyzed by Sirius Red assay to verify the homogeneity of the biofunctionalization. The bio-functionalized sample turned red, indicating that the molecules of Sirius Red were successfully

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FIGURE 1. (A) Comparison between the spectra of films treated at different plasma power outputs: PLLA (black), 50 W (red), 100 W (blue) and 150 W (green). Peaks associated to the presence of COOH groups are highlighted in red squares. (B) SEM micrographs of PLLA films treated with different plasma power outputs: (i) Control sample, (ii) 50 W, (iii) 100 W, and (iv) 150 W. Scale bar represents 100 mm. (C) Density of carboxylic groups on PLLA films as function of plasma power output. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

coupled to the collagen attached to surface. Optical microscopy also showed uniformly distributed red coloring, which demonstrates the homogeneity of the grafting and confirms the successful bio-functionalization of the construct [Fig. 3(C)]. A significant decrease in the water contact angle of functionalized samples also confirms this statement. Nonfunctionalized membranes were highly hydrophobic, with a contact angle of 106 6 0.1 , as shown in Figure 3(B). After collagen coupling membranes turned highly hydrophilic, as demonstrated by the complete absorption of the water drop within just 34 ms [Fig. 3(B)]. The average contact angle on PLLA membranes after collagen functionalization was significantly lower, at 48.9 6 1.1 . IR-ATR spectra of PLLA porous membranes before and after plasma treatment at 50 W and after collagen coupling are shown in Figure 4. Spectrum of collagen-functionalized membranes evidences the presence of two characteristic peaks of collagen molecules, which are not observed for the untreated controls. The first peak, at about 1664 cm21, corresponds to the amide I signal (stretching of C@O bonds), while the second peak at 1574 cm21 is due to the amide II signal (bending of NH bond).18 These two peaks suggest the presence of collagen on the membranes surface, confirming successful grafting.

Morphological and mechanical characterization of PLLA porous constructs SEM images, mechanical, and morphological properties of PLLA porous membranes fabricated by TIPS are presented in Figure 5.

FIGURE 2. Quantification of collagen mass after different times of incubation in EDC/NHS and collagen solution. Data is presented as mean 6 standard deviation of n 5 3 samples. (*) Corresponds to a significant difference compared to EDC/NHS treatment for 1 h and collagen coupling (COL) for 1 h (p < 0.05).

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FIGURE 3. (A) Optical micrographs of porous membranes after the immersion in TBO solution, before (control) and after AAc grafting: the blue color evidences the presence of homogeneously distributed carboxyl groups linked to the TBO molecules. (B) Water contact angle measurements on PLLA films (upper figures refer to PLLA membranes before collagen functionalization, while lower row represents PLLA functionalized with collagen, after 34 ms of water-drop deposition). (C) Optical micrographs of porous membranes after the immersion in Sirius red solution, before and after collagen grafting. The red color observed evidences the presence of collagen molecules linked to the Sirius red molecules. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

Porous membranes with open and interconnected pores in the range of about 25 and 125 mm, main dimensional distribution of pores at 59.55 6 24.9 mm and porosity of 92% 6 2% were obtained. The construct showed an elas-

tic behavior with an initial linear trend, followed by a plastic-plateau, which brought to the failure of the membrane after an elongation of 4.6 and 8.0%, respectively. The elastic modulus, calculated from the linear part of the

FIGURE 4. Comparison between the FTIR-ATR spectra of the membranes before and after plasma treatment and collagen grafting: PLLA (black), membrane treated with plasma at 50 W (red), and membrane after reaction with collagen (green). Peaks associated to collagen (amide I and amide II) are highlighted in the red squares. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

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FIGURE 5. SEM micrographs, mechanical properties and porosity of PLLA porous membranes prepared by TIPS after dissolving the polymer (5% w/v) in dioxane. (A) Control membrane (prepared in absence of nonsolvent) and (B) porous constructs (water, 5 vol %, as nonsolvent). Scale bar represents 100 mm.

stress-strain curve, is 18 6 1 MPa and the elongation reaches about 10% 6 1%. Nanoparticles characterization Nano-sized particles with negative surface charge were successfully obtained with both polymers, PU and PCL, as summarized in Figure 6(A). Mean size of plain nps varies between 196 and 218 nm for PU and PCL, respectively. Drug encapsulation did not lead to any significant variation in nps size or polydispersity index (PDI), indicating that IND did not affect the size distribution of the carriers.28 Particles made of PCL showed larger dimensions compared to those obtained from the synthesized polyester urethane, regardless of the presence of indomethacin, as confirmed by SEM micrographs [Fig. 6(B)]. PU nps showed a high encapsulation efficiency of 89% 6 2.3% while EE of PCL nps was even higher at 99% 6 0.8%. This observation is in agreement with the results reported by other authors for PCL nps. For instance, Bernardi et al.29 obtained small PCL nps of 230 nm with an IND EE close to 100%. Drug release profiles are shown in Figure 6(C). Release profile from PU carriers show an initial linear pattern, with about 20% of the drug released during the initial 2 h of incubation [see inset in Fig. 6(C)], followed by a sustained release over the following 48 h, when 91% of the encapsulated drug was released. On the other hand, nps prepared with PCL showed a higher burst effect compared to PU nps, releasing 60% of the encapsulated drug within the first 2 h. Moreover, the drug release occurs much faster, achieving almost entire release of the loaded drug (92%) at 24 h of incubation. Zavisova et al.30 showed a complete IND release from PLLA nps during the first day of incubation (30 h), while Suksiriworapong et al.31 reported a similar release profile from PCL-PEG nps. PU nps showed a better ability to control the release of IND during the initial hours of incubation compared to PCL nps, reducing the burst effect. This is particularly advantageous when dealing with potentially toxic drugs that can cause cell death or tissue damage. These results confirm

our earlier studies on PU nps, which have shown to be promising drug carriers, due to their ability to sustain the release of drugs for longer periods of time compared to polyesters.23 To our knowledge, no studies on the encapsulation of IND inside PU nps have been so far reported in the literature. IND release from nanoparticles-containing scaffolds can be sustained for 24–48 h, depending on the nps-forming polymer (Fig. 7). Similarly to free nps, scaffolds with PCL nps displayed a faster release. About 50% of the theoretically loaded drug was released at 24 h, followed by no additional release. This indicates that a part of the loaded nanoparticles were not adhered to the scaffolds surface and were removed during the washing step. A more sustained IND release was observed from scaffolds loaded with PU nps, for which 60% of the loaded-drug was released between 48 and 72 h, followed by no additional release. To our knowledge, only few authors have analyzed the release on IND from porous scaffolds.32,33 In all cases the drug was dispersed in the scaffold polymer matrix, rather than encapsulated into nanocarriers. For instance Thakur et al.33 have studied Indomethacin release from cross-linked gelatin sponges. In spite of the high initial content of indomethacin, scaffolds were only able to release 30% of the loaded drug, indicating that a substantial loss of the active principle occurred during processing. The release was rapid and completed within the first 7 h of incubation, followed by no additional release. Our approach, based on embedding IND-loaded nps on porous scaffolds showed promising results in terms of higher IND loading and sustained drug release, thus warranting further investigation. Characterization of porous membranes functionalized with collagen and nanoparticles The presence of the nps in the porous membranes was verified by SEM microscopy [Fig. 6(D)]. SEM micrographs demonstrated that nps were well distributed on the surface of the porous membranes without altering the porous

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FIGURE 6. (A) SEM micrographs of nanoparticles made of PCL and NS-HC2000 loaded with Indomethacin. Scale bar represents 250 nm. (B) Drug release profile of indomethacin encapsulated within nanoparticles prepared with PCL and PU (The inset provides a detail of the release profile for first 3 h of incubation). (C) Mean size, PDI zeta potential and EE of nps prepared with PCL and with the synthesized polyesterurethane NS-HC2000.

structure or closing the pores. It is possible that the negatively charged nps surface led to their adsorption on the collagen-modified scaffold. Cell viability of keratinocytes Cellular viability and ingrowth were analyzed using keratinocytes seeded on the functionalized porous membranes with and without IND-loaded nps after 1, 3, and 7 days culture. Figure 8(A) shows the cellular viability determined by crystal violet assay on porous membranes before (PLLA) and after (PLLA-COL) collagen grafting, and on porous

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membranes containing IND-loaded PCL and PUR nps. In general, an increase of cell viability along the cell incubation period is observed. In particular, cell adhesion shows no significant differences among samples at 24-h incubation. The presence of collagen molecules on the polymeric membranes surface enhanced significantly the cell viability compared to their un-functionalized counterparts, as observed after 3 days of incubation. Moreover, a more evident increase of cell growth was observed at day 3 for collagenfunctionalized porous membranes and in presence of INDloaded PCL nps while the presence of IND-loaded PU nps resulted in significantly lower cell viability compared to

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FIGURE 7. (A) SEM micrographs of nanoparticles fabricated with PCL and NS-HC2000 within the porous membranes. Scale bar represents 2 lm. (B) IND release profiles from PCL and PU nanoparticles loaded on the porous membrane (S-PCL-nps and S-PU-nps respectively).

functionalized membranes with only collagen. Thus, in contrast to the results obtained for collagen-modified membranes loaded with PU-nps, the presence of PCL-nps does not seem to have a negatively effect on cell adhesion and growth, since the results for these samples are comparable with controls. Figure 8(B) shows the cellular colonization, adhesion and distribution within the different porous membranes. Functionalization of PLLA porous membranes with type I collagen evidenced an enhanced cell migration and colonization of the construct compared with the untreated counterpart. In addition, the presence of the polymeric loaded-nps does not interfere with the processes of migration, colonization, adhesion and proliferation within the functionalized membranes.

DISCUSSION AND CONCLUSIONS

The complex hierarchical organization of the dermal tissue requires biomimetic strategies to induce skin regeneration. These are based on the use of porous constructs with good mechanical properties, biocompatibility and proper architecture, able to enhance and support new tissue ingrowth through specific biological cues. Reduction of the risk of inflammation and post-operative infections should be considered as an additional requirement for cutaneous tissue regeneration. Three fundamental features for the design and fabrication of a biomimetic skin substitute are: (i) the nature of the material, (ii) the surface topography and (iii) the architecture. In this work, polymers used for the scaffold and for nps manufacturing are well-known biodegradable and

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FIGURE 8. (A) Keratinocytes cell viability on fabricated PLLA porous membranes before and after collagen grafting and, containing Indomethacin-loaded PCL and PU nanoparticles after 1, 3, and 7 days of culture. (*) Corresponds to a significant difference compared to PLLA unfunctionalized membranes, (**) significant difference compared to collagen-functionalized membranes and (o) significant difference between collagen-functionalized membranes containing both Ind-loaded PCL Nps (PLLA-COL PCLIND) and PU Nps (PLLA-COL PUIND) (p < 0.05) (B). Cellular histology of: (i) PLLA membranes, (ii) Collagen-functionalized membranes (PLLA-COL), (iii) Collagen-functionalized membranes containing Indloaded PCL Nps (PLLA-COL PCLIND), and (iv) Collagen-functionalized membranes containing Ind-loaded PU Nps (PLLA-COL PUIND), after 14 days of culture. The scale bar corresponds to 100 mm. [Color figure can be viewed in the online issue, which is available at wileyonlinelibrary.com.]

biocompatible synthetic polymers. Furthermore, an ECM-like coating was assembled using collagen, the major structural protein in skin, which is known to promote cell adhesion and colonization. Surface topography is fundamental to impart specific biological cues that may enhance cellular response, guiding the formation of a well-organized new tissue. For this purpose, an Ar plasma-driven chemical surface modification was performed for covalent collagen grafting on the PLLA scaffold surface, in order to bio-activate the material and to stimulate a desired cellular response, since interactions between the material and the biological environment always take place at the surface. Bio-functionalization was obtained by AAc immobilization on PLLA through Ar-plasma treatment, followed by covalent coupling of collagen through EDC-NHS mediated chemistry. Three different plasma power outputs, at 50, 100, and 150 W, were tested in order to obtain the optimal conditions in terms of higher number of superficial carboxylic groups for further collagen coupling and reduced surface damages. Higher treatment power (150 W) resulted in lower -COOH groups exposure, despite the higher wettability indicated by contact angle measurements. The increase in hydrophilicity of sample functionalized at 150 W could

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be attributed to a possible increase of surface roughness26 caused by plasma etching of amorphous regions34 and/or formation of diverse polar groups on the surface, such as C@O and OH groups.35 Polymer degradation caused by higher power promotes the weakening of the structure and changes the topography of the films. Similarly, treatment at 100 W also caused certain degradation when compared to the lowest plasma potency (50 W), as evidenced by SEM. Because no significant differences in carboxyl groups exposure was evidenced between samples treated at 50 and 100 W, the lowest power was selected for AAc grafting on PLLA scaffolds. Collagen was successfully coupled to the superficial carboxylic acids using the long-term processing condition of 1h of activation in EDC/NHS followed by 20 h of coupling reaction in collagen solution, and the uniformity and quantity of the biomolecule was demonstrated by optical microscopy and by Sirus-Red colorimetric assay. The presence of type I collagen enhanced cell migration and colonization within the construct, since higher cell viability, indicative of cell growth, was observed at day 3 collagen-functionalized porous membranes compared to their un-functionalized counterparts. The presence of the biomolecule favors the cellular interaction due to the

POLYMER NANOPARTICLES FOR WOUND HEALING

ORIGINAL ARTICLE

improved surface properties36 such as hydrophilicity, and the presence of cell recognition sites in collagen molecules (GFOGER).37 Our bio-functionalization approach resulted in a homogenous and continuous coating with collagen, mimicking the natural environment in which cells grow, without altering the porous architecture of the constructs. Topography is another key feature for successful dermal regeneration. The optimal pore size for cutaneous tissue substitutes is found to be between 20 and 125 mm, which is known to favor cell penetration and migration as well as to enhance gas and nutrients diffusion.15 PLLA porous membranes showed a promising morphology for wound dressing applications, with adequate pore size in the range of 25–125 mm, good pore interconnection and a fairly uniformly distributed porosity of 92% 6 2%. The construct showed an elastic behavior with an elongation at break 8% and an E modulus of 18 6 1 MPa. These values are very close to the optimal values required for wound repair (E modulus of ca. 20 MPa).38 The PLLA membranes showed adequate mechanical, morphological and topographical properties to be applied as scaffolds for wound dressing. To reduce the risk of implant failures associated to infections or inflammatory processes, PCL and PU INDloaded nps were introduced inside the bio-functionalized constructs. Our results indicate that both types of nps are suitable candidates for the release of anti-inflammatory drugs from porous scaffolds in wound dressings applications, since they allow the release of the active principle in a rather short time. This phenomenon is compatible with the physiological process of wound healing. Indeed, the first phase of tissue healing is characterized by an inflammatory process of the affected tissue which, begins just a few hours after the injury has occurred and usually finishes after 3 days.39,40 During this inflammation step, the controlled dosage of an anti-inflammatory may facilitate the healing process, by limiting inflammation. Cellular tests demonstrated that the introduction of Indomethacinloaded PCL has a positive effect on cellular ingrowth and distribution inside the wound-dressing construct that can potentially enhance a healing response and tissue regeneration. In summary, our results indicate that this biomimetic device is a promising candidate for wound healing and soft tissue regeneration since it shows (i) proper porosity, pore size and mechanical properties, (ii) biomimicry of ECM, and (iii) therapeutic potential. ACKNOWLEDGMENTS

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