A mechanism for the enhanced attachment and proliferation of fibroblasts on anodized 316L stainless steel with nano-pit arrays Siyu Ni,1,2 Linlin Sun,2 Batur Ercan,2 Luting Liu,2 Katherine Ziemer,2 Thomas J. Webster2,3 1

College of Chemistry, Chemical Engineering and Biotechnology, Donghua University, Shanghai 201620, China Department of Chemical Engineering, College of Engineering, Northeastern University, Boston, Massachusetts 3 Center of Excellence for Advanced Materials Research, King Abdulaziz University, Jeddah 21589, Saudi Arabia 2

Received 4 December 2013; revised 24 January 2014; accepted 10 February 2014 Published online 8 March 2014 in Wiley Online Library (wileyonlinelibrary.com). DOI: 10.1002/jbm.b.33127 Abstract: In this study, 316L stainless steel with tunable nanometer pit sizes (0, 25, 50, and 60 nm) were fabricated by an anodization procedure in an ethylene glycol electrolyte solution containing 5 vol % perchloric acid. The surface morphology and elemental composition of the 316L stainless steel were characterized by scanning electron microscopy (SEM), atomic force microscopy (AFM), and X-ray photoelectron spectroscopy (XPS). The nano-pit arrays on all of the 316L stainless steel samples were in a regular arrangement. The surface properties of the 316L stainless steel nano-pit surface showed improved wettability properties as compared with the untreated 316L stainless steel, as demonstrated by the lower contact angles which dropped from 83.0 to 28.6 to 45.4 . The anodized 316L stainless steel surfaces with 50 nm and 60 nm diameter pits were also more rough at the nanoscale. According to MTT assays, compared with unanodized

(that is, nano-smooth) surfaces, the 50 and 60 nm diameter nano-pit surfaces dramatically enhanced initial human dermal fibroblast attachment and growth for up to 3 days in culture. Mechanistically, this study also provided the first evidence of greater select protein adsorption (specifically, vitronectin and fibronectin which have been shown to enhance fibroblast adhesion) on the anodized 316L stainless steel compared with unanodized stainless steel. Such nano-pit surfaces can be designed to support fibroblast growth and, thus, improve the use of 316L stainless steel for various implant applications (such as for enhanced skin healing for amputee devices C 2014 Wiley Periodicals, Inc. J and for percutaneous implants). V Biomed Mater Res Part B: Appl Biomater, 102B: 1297–1303, 2014.

Key Words: nano-porous, stainless steel, adhesion, proliferation, fibroblast, nanotechnology

How to cite this article: Ni S, Sun L, Ercan B, Liu L, Ziemer K, Webster TJ. 2014. A mechanism for the enhanced attachment and proliferation of fibroblasts on anodized 316L stainless steel with nano-pit arrays. J Biomed Mater Res Part B 2014:102B:1297–1303.

INTRODUCTION

Over the past few decades, the field of biomaterials has shifted in emphasis from achieving a bioinert tissue response to stimulating specific cellular responses at the molecular level.1 Designing biomaterial surfaces to direct specific cellular responses in a predictable manner has drawn enormous attention, yet little work has been done for one of our oldest biomaterials, stainless steel;2 316 L stainless steel is a widely used metallic biomaterial in cardiovascular stents, orthopedic implants, and spinal fixation devices because of its high strength, durability, and acceptable biocompatibility.2 However, 316L stainless steel has always been viewed as bioinert and sometimes causes longterm clinical problems (such as loosening from juxtaposed bone for orthopedic applications over the long term). Moreover, the long-term implantation of 316L stainless steel in the body results in the slow release of a small amount of metal ions, such as Ni, Cr, which are toxic and allergenic.3,4 To solve these problems, surface modification seems to be a

more economical and efficient way to promote immediate and long-term implant fixation (than using pharmaceutical agents which may have side effects), thus, avoiding longterm implant problems.1,2 Various surface modification techniques, such as chemical etching, electrochemical treatment, ion implantation, electron beam irradiation, and the application of a variety of coatings, have been used to improve the bioactivity of metallic implants.1,5 Among them, the application of electrochemical oxidation has attracted increasing interest because of its simplicity, low cost, and controllability at the nanoscale.6 Electrochemical anodic oxidation could be used to grow a thick and uniform oxide layer on metals and several alloys, such as titanium, aluminum, tantalum, and their alloys, and has been reported to form nano-structural oxide layers on their surfaces by this method.7,8 At the same time, the formation of a stable oxide layer on metallic implant materials may also act as a barrier to reduce the release of metal ions into the human body.9 In fact, many studies have

Correspondence to: T. J. Webster (e-mail: [email protected]) Contract grant sponsor: Northeastern University

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reported the use of anodization to enhance their corrosion resistance.4 In particular, through changes in surface energy, it has been recognized that surface nanostructures on biomaterials greatly influence cellular behavior.5,7,8 For example, many studies have indicated that nano-tubular and nano-textured titania (created through various anodization conditions) could modulate the functions of many types of cells and might be used as an excellent bioactive interface for implantable devices.2,10–12 Specifically, it has been reported that fine tuning the dimensions of such titania nanofeatures in the 10 to 200 nm range can enhance functions of osteoblasts, keratinocytes and urothelial cells.13–16 Anodization of titanium to form titania nanotubular structures has also been shown to inhibit bacteria growth and macrophage (i.e., inflammatory cell) functions.17 Therefore, similar to titanium, surface modification of 316L stainless steel at the nanoscale may generate functional nano-structured surfaces to also control such cell behavior. In fact, in a previous study, Dıaz et al. used two different anodization methods to grow chromium oxide on 316L stainless steel and evaluate its in vitro cytotoxicity. However, the obtained surface features in their study were not at the nanometer range, and cellular responses to anodized surfaces on stainless steel were not determined.9 Recently, Pan et al. prepared different sized nano-porous surfaces on 316L stainless steel by electrochemical treatment and investigated the effects of these nanopores on the growth and function of fibroblasts. Their results showed that these nano-patterned surfaces can modulate integrin expression, and enhance the adhesion and migration of fibroblasts.18 However, they used a mouse fibroblast cell line (NIH-3T3), which has been shown to not accurately reflect the human fibroblast response. In addition, to the best of our knowledge, no one has elucidated the mechanism of why fibroblast density may be altered on anodized nanostructured stainless steel, which is critical information for the further optimization of anodized stainless steel for numerous applications including its use for amputee and percutaneous applications. Therefore, in the present study, nano-pit arrays with diameters ranging from 0 to 60 nm were created on 316L stainless steel by an anodization procedure. Then, the effects of these anodized nano-structured surfaces on the attachment and proliferation of human dermal fibroblasts were mechanistically evaluated and compared, which might provide an improved understanding of the effects of these nano-pit environments on basic cell behaviors. MATERIALS AND METHODS

Sample preparation Commercial AISI 316L stainless steel (SS) foils (0.5 mm) (polished on both sides) (Goodfellow, Cambridge Ltd. England) were cut into squares 2.5 cm 3 2.5 cm. Before anodization, the specimens were cleaned and degreased in acetone (Sigma-Aldrich), 70% ethanol (Decon), and deionized water (Milli-Q water), separately, each for 15 min. The electrolyte was an ethylene glycol (EG, 99.8%, anhydrous,

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Sigma-Aldrich) solution containing 5 vol % perchloric acid (HClO4, 70%, Sigma-Aldrich). The anodization was performed in a stirred two-electrode configuration by a programmable direct current power supply (Sorensen XHR150-7, San Diego, CA) with a platinum mesh (Alfa-Aesar) serving as the cathode. During anodization, the anode and platinum cathode were kept at a distance of 30 mm from each other. In order to create nano-pits on 316L SS with different diameters, the applied anodization voltages were altered from 20, 30, 40 to 50 V. For the cell experiments, the specimens were cut into identical size pieces (10 mm 3 10 mm) after anodization. Untreated 316L SS was used as a control surface. All samples were sterilized using 70% ethanol and UV light exposure for 30 min and 4 h, respectively. Surface characterization Scanning electron microscopy (SEM). After anodization, the samples were thoroughly rinsed with DI water and then dried at room temperature. The surface morphology of 316L SS before and after anodization was characterized by scanning electron microscopy (SEM, Hitachi S-4800, Tokyo, Japan). Atomic force microscopy (AFM). For surface roughness measurements, an Asylum Research atomic force microscope (AFM) (Santa Barbara, CA, USA) was used to scan the untreated and anodized 316L SS substrates. Each sample was analyzed in ambient air under non-contact mode using a silicone ultrasharp cantilever (probe tip radius of 10 mm; MikroMasch, Wilsonville, OR); 1.5 3 1.5 mm AFM fields were analyzed and the scan rate was chosen as 1 Hz. Image analysis software (XEI) was used to generate micrographs and to quantitatively compare the root-mean-square roughness (RMS) of the untreated and anodized 316L SS substrates. At least five different spots on each sample were measured for statistical purposes (n 5 5). Chemical analysis. For chemical analysis of the top surface layer of the 316L SS before and after anodization, X-ray photoelectron spectroscopy (XPS) and peak curve fitting using software (Escaaab250, Thermo Electron, American) were employed. Contact angle analysis. Water contact angles were determined using a drop shape analysis system (SEO Phoenix 300, Korea). The contact angle of 1 mL sessile droplets of double distilled water (5 s after being placed on the surface) was measured for all samples at room temperature. At least five measurements were carried out for every sample (n 5 5). Cells Human fibroblasts (ATCC, CCL-110) at population numbers less than seven were used for all cell experiments. The cells were cultured in Eagle’s Minimum Essential Medium (EMEM) (ATCC) supplemented with 10% fetal bovine serum

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(FBS; Sigma) and 1% penicillin/streptomycin (P/S; Sigma) at 37 C and 5% CO2 in a humidified atmosphere. Cell attachment and proliferation assays Fibroblasts were seeded onto the substrates at a density of 5000 cells/cm2 and were allowed to adhere for 4 h in a 37 C, humidified 5% CO2 atmosphere. The medium was removed from the wells four hours later and the cells were washed twice with PBS. All samples were then transferred to fresh 24-well tissue culture plates, and 1 mL of fresh growth medium was added to each well. Next, 150 mL of a dye solution from the cell proliferation assay kit (3-(4, 5dimethylthiazol-2-yl)-2, 5-diphenyltetrazolium (MTT); Promega) was added to each well and the plates were incubated at 37 C in a humidified 5% CO2 atmosphere. After four R (Promhours, 1 mL of the Solubilization solution/Stop MixV ega) was added to each well and the plates were allowed to stand overnight in a humidified atmosphere to solubilize the formazan crystals completely. The contents of the wells were then mixed to obtain a uniformly colored solution, and then transferred to 96-well plates (200 mL per well). The absorbance at 570 nm was recorded using a microplate reader (Molecular Devices, SpectraMax M3). The value of the background absorbance at 570 nm without cells run in parallel with the experimental samples was subtracted to yield a corrected absorbance. The viable cell number was determined from a standard curve of absorbance versus known concentrations of fibroblasts in solution run in parallel with the MTT assay. Five specimens of each material were tested and each test was performed in triplicate. A similar process to that described above was used for cell proliferation. The cells were seeded at 5000 cells/cm2 per substrate and allowed to proliferate for 24 and 72 in a 37 C, humidified 5% CO2 environment. The medium in the wells were remove and replaced with fresh EMEM every 24 h for all substrates. At the prescribed time point, the samples were rinsed with PBS and transferred to fresh medium in a new 24-well plate. The MTT assay was conducted as described above to determine cell viability. Five specimens of each material were tested and each test was performed in triplicate. ELISA assays For fibronectin adsorption assays, stainless steel substrates were soaked in the EMEM cell culture media mentioned above supplemented with 5 mg/500 mL of fibronectin in PBS (Sigma) for 1 h under standard incubator conditions. For vitronectin adsorption assays, stainless steel substrates were soaked in the EMEM cell culture media mentioned above supplemented with 0.5 mg/500 mL of vitronectin in PBS (Sigma) for 1 h under standard incubator conditions. The substrates were then rinsed with PBS, blocked with 2% bovine serum albumin (BSA; Sigma) for 1 h, and incubated with anti-bovine vitronectin (1:100; Accurate Chemical) or anti-bovine fibronectin (1:100; Chemicon, Temecula, CA) for 1 h. Immediately thereafter, the substrates were rinsed with Tris buffered saline-0.1% Triton X-100 (Sigma) and incu-

bated with a horse radish peroxidase conjugated anti-rabbit secondary antibody (1:100; Bio-Rad). An ABTS (2,20 -azinobis(3-ethylbenzthiazoline-6-sulfonic acid)) soluble substrate kit (Vector Labs, Burlingame, CA) was used to detect secondary antibodies spectrophotometrically (SpectroMAX 190, 488 nm; Molecular Devices, Palo Alto, CA) per the manufacturer’s instructions. Statistical analysis Data were collected and the significant differences were assessed with the probability associated with a one-tailed Student’s t-test. Statistical analysis was performed using R 2010 (Microsoft Corporation, Redmond, WA). All ExcelV results were expressed as the mean 6 standard deviation from the quintuple samples in each experiment. Differences were considered statistically significant at p < 0.05. RESULTS

Characterization of the prepared samples Figure 1 shows the SEM micrographs of the unanodized (control) and anodized 0 (flat), 25, 50, and 60 nm nano-pits formed on the 316L stainless steel (SS). In order to present the data clearly, 316L stainless steel with tunable pit sizes (0, 25, 50, and 60 nm) were denoted as SS0, SS25, SS50, and SS60, respectively, according to the pit sizes on the SS surfaces. The images reflect changes in the nano-structure because of the differences in the pit size. As can be seen, SS25 through 60 showed an ordered and uniform nanoporous honeycomb morphology with long-range order. Figure 2 shows AFM images of SS before and after anodization, which is in agreement with the SEM results. It is clear from Figure 2(A) that the surface was more smooth at the nanoscale before the electrochemical treatment. The formation of shallow pits can be observed as shown in Figure 2(C). Then, ordered pit arrays were observed on the entire surfaces for Figure 2(D,E). The Rrms values of the different surfaces are shown in Table I. As expected, anodization of the SS altered their roughness. The Rrms value for the untreated SS was 0.24 6 0.04 nm. With the increase of the pit size on the anodized SS surfaces, the roughness increased from 0.15 6 0.01 nm to 3.83 6 0.02 nm (Rrms[SS0] < Rrms[SS25] < Rrms[SS50] < Rrms[SS60]). Specifically, a noticeable change in surface roughness of SS50 and SS60 was seen. At the same time, the Rrms results confirmed that there were significant variations in SS surface nano-topography. Water contact angles are presented in Table I. Surface wettability (i.e., hydrophilicity or hydrophobicity) has been traditionally determined by water contact angles according to the definition of hydrophilic surfaces having a water contact angle less than 65 . Thus, all the anodized SS surfaces in the presents study were hydrophilic according to that definition whereas SS was hydrophobic. With the increase of the pit size on the SS surfaces, the water contact angles varied from 28.6 to 45.4 . SS0, SS25, and SS50 had similar hydrophilic surfaces with water contact angles of 31.2 6 2.1 , 28.6 6 4.9 , and 32.0 6 3.5 , respectively. SS60 showed a higher contact angle (45.4 6 3.2 ). Untreated SS

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FIGURE 1. SEM micrographs of the 316L stainless steel before (A or SS) and after anodization (B: 20V or SS0; C: 30V or SS25; D: 40V or SS50; and E: 50V or SS60) Images show highly ordered nano-porous structures with four different pore sizes (A and B: 0 nm, C: 25 nm, D: 50 nm, and E: 60 nm).

FIGURE 2. Atomic force microscope images showing the topography the 316L stainless steel before (A: SS) and after anodization (B: SS0; C: SS25; D: SS50; and E: SS60).

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TABLE I. Surface Roughness (Rrms) and Static Water Contact Angles on 316 L Stainless Steel (SS), 316L Stainless Steel Anodized at 20 V (SS0), 30 V (SS25), 40 V (SS50), and 50 V (SS60) Sample SS SS0 SS25 SS50 SS60

Rrms(nm)

Contact Angle (u)

0.24 6 0.04 0.15 6 0.01 0.22 6 0.01 0.71 6 0.05 3.83 6 0.02

83.0 6 2.3 31.2 6 2.1 28.6 6 4.9 32.0 6 3.5 45.4 6 3.2

Data are represented as Mean 6SD, n 5 5. All values are statistically (p

A mechanism for the enhanced attachment and proliferation of fibroblasts on anodized 316L stainless steel with nano-pit arrays.

In this study, 316L stainless steel with tunable nanometer pit sizes (0, 25, 50, and 60 nm) were fabricated by an anodization procedure in an ethylene...
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