Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds for long-term stem cell growth Shelby A. Skoog, Alexander K. Nguyen, Girish Kumar, Jiwen Zheng, Peter L. Goering, Anastasia Koroleva, Boris N. Chichkov, and Roger J. Narayan Citation: Biointerphases 9, 029014 (2014); doi: 10.1116/1.4873688 View online: http://dx.doi.org/10.1116/1.4873688 View Table of Contents: http://scitation.aip.org/content/avs/journal/bip/9/2?ver=pdfcov Published by the AVS: Science & Technology of Materials, Interfaces, and Processing Articles you may be interested in Materials and technologies for fabrication of three-dimensional microstructures with sub-100nm feature sizes by two-photon polymerization J. Laser Appl. 24, 042014 (2012); 10.2351/1.4730807 Synthesis of hybrid microgels by coupling of laser ablation and polymerization in aqueous medium J. Laser Appl. 24, 042012 (2012); 10.2351/1.4730803 PCL loaded with sol-gel synthesized organic-inorganic hybrid fillers: From the analysis of 2D substrates to the design of 3D rapid prototyped composite scaffolds for tissue engineering AIP Conf. Proc. 1459, 26 (2012); 10.1063/1.4738387 Manipulating interfaces in a hybrid solar cell by in situ photosensitizer polymerization and sequential hydrophilicity/hydrophobicity control for enhanced conversion efficiency Appl. Phys. Lett. 92, 193307 (2008); 10.1063/1.2929368 Use of two-photon polymerization for continuous gray-level encoding of diffractive optical elements Appl. Phys. Lett. 90, 073503 (2007); 10.1063/1.2426923
Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds for long-term stem cell growth Shelby A. Skoog Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965 and Office of Science and Engineering Laboratories, Center for Devices and Radiological Health, U.S. Food and Drug Administration, Silver Spring, Maryland 20993
Alexander K. Nguyen Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965 and Department of Nanotechnology, Laser Zentrum Hannover e. V., Hannover D-30419, Germany
Girish Kumar, Jiwen Zheng, and Peter L. Goering Office of Science and Engineering Laboratories, Center for Devices and Radiological Health, U.S. Food and Drug Administration, Silver Spring, Maryland 20993
Anastasia Koroleva and Boris N. Chichkov Department of Nanotechnology, Laser Zentrum Hannover e. V., Hannover D-30419, Germany
Roger J. Narayana) Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965
(Received 4 March 2014; accepted 17 April 2014; published 28 April 2014) Two-photon polymerization is a technique that involves simultaneous absorption of two photons from a femtosecond laser for selective polymerization of a photosensitive material. In this study, two-photon polymerization was used for layer-by-layer fabrication of 3-D scaffolds composed of an inorganic–organic zirconium oxide hybrid material. Four types of scaffold microarchitectures were created, which exhibit layers of parallel line features at various orientations as well as pores between the line features. Long-term cell culture studies involving human bone marrow stromal cells were conducted using these 3-D scaffolds. Cellular adhesion and proliferation were demonstrated on all of the scaffold types; tissuelike structure was shown to span the pores. This study indicates that two-photon polymerization may be used to create microstructured scaffolds out of an inorganic–organic zirconium oxide hybrid material for use in 3-D tissue culture systems. C 2014 American Vacuum Society. [http://dx.doi.org/10.1116/1.4873688] V I. INTRODUCTION Cellular response is strongly influenced by the surrounding biological environment through complex interactions with biochemical stimuli, mechanical forces, and structural components of the extracellular matrix (ECM).1–3 The threedimensional architecture of the ECM consists of interwoven fibrillar proteins embedded within a network of proteoglycans; this structure provides mechanical and physical cues that regulate cellular function. Since it is well known that cells are highly influenced by the local microenvironment, it is important to understand the underlying mechanisms of the cell–structure relationship.4–7 To evaluate the effect of ECM structure on cellular response, researchers employ micro- and nanoscale topographies as well as complex 3-D architectures that mimic the native cell environment. Utilizing 3-D structures for advanced cell culture and tissue engineering provides a more physiologically relevant model of the in vivo environment than 2-D structures. 2-D surfaces may confine cells to a single plane of growth; in addition, they may limit cells from a)
Author to whom correspondence should be addressed; electronic mail: [email protected]
029014-1 Biointerphases 9(2), June 2014
exhibiting their natural morphologies and cell–cell interactions. In contrast, 3-D scaffolds may provide porous microarchitectures similar to those of the natural ECM and may impart physical as well as chemical cues.8,9 Several studies have utilized both textured surfaces and 3-D scaffolds with physiologically relevant features to evaluate cell adhesion,10–12 morphology,13 alignment,13 migration,14–16 differentiation,16–19 proliferation,20 protein expression,21 and gene expression.21 Further, studies have demonstrated increased cell attachment, proliferation, and differentiation of bone-derived cells on 3-D scaffolds compared to 2-D surfaces.22,23 Understanding how cells respond to structural cues in 3-D will contribute to development of advanced biomaterials for improved implants, enhanced in vitro tissue models, and optimized scaffolds for tissue regeneration. For a systematic approach to assess cellular responses to surface topographies and 3-D architectures, materials with micro- and nanoscale features must be fabricated using precise, reproducible methods. Advances in microfabrication technology have enabled production of complex structures at high resolution for a variety of biomaterials. Two-photon polymerization (2PP) is a rapid prototyping technique that enables fabrication of 3-D structures with high spatial resolution
1934-8630/2014/9(2)/029014/7/$30.00
C 2014 American Vacuum Society V
029014-1
029014-2 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
down to 100 nm.24 The process uses the two-photon effect, which involves almost simultaneous absorption of two photons from a femtosecond laser to polymerize a photosensitive polymer resin in a layer-by-layer fashion. In 2PP, an ultrashort-pulsed laser beam is highly focused on a small, three-dimensional unit volume (voxel) of liquid resin. Since the two-photon effect is confined to this small volume, the chemical reactions between the photoinitiator molecules and the monomers (and subsequent polymerization of the material) are localized.24 2PP enables fabrication of structures with complex geometries and precise dimensions. A wide variety of photosensitive polymers, fillers, and photoinitiators have been utilized with 2PP; as such, fabrication of materials with a wide range of surface chemistries and mechanical properties is possible using this approach. This versatility in material properties and high fidelity between computer design and 2PP-fabricated structures may facilitate processing of more accurate models of the biophysical environment. Several researchers have utilized 2PP for fabrication of 3-D microarchitectures and textured surfaces to examine cellular activity. Studies have incorporated 2PP structures with various geometries and dimensions to evaluate cell adhesion,1,25 morphology,4,26,27 orientation,4,26,27 viability,28 proliferation,28 and cell functionality.29 Recently, cell response has been evaluated using 2PP-fabricated structures composed of photosensitive inorganic-organic hybrid copolymers, such as the commercially available ORMOCERV material (Microresist Technology GmbH, Berlin, Germany).4,26,30 ORMOCER material is synthesized using a sol–gel process; the final material contains cross-linked networks of inorganic and organic components. This hybrid sol–gel material exhibits high thermal stability, high chemical stability, good mechanical properties, and biocompatibility.31 In this study, we demonstrate reproducible fabrication of 3-D scaffolds composed of an inorganic–organic zirconium oxide hybrid material using two-photon polymerization. Four different scaffold microarchitectures were produced with different layers of parallel lines at various orientations. Long-term tissue culture studies were conducted using human bone marrow stromal cells (hBMSCs) to evaluate cellular attachment, alignment, and growth on the 3-D microstructures and to assess the potential of two-photon polymerization technology for long term 3-D stem cell culture for enhanced tissue engineering applications. R
029014-2
zirconium n-propoxide (ZPO). The chemical structures for the inorganic–organic hybrid copolymer components are shown in Fig. 1. Both organic monomers, MAPTMS and methacrylic acid, contain methacrylate moieties that are photopolymerizable. The inorganic network components include alkoxysilane groups of MAPTMS and zirconium npropoxide. During synthesis, the resin precursors first underwent catalytic hydrolysis and condensation. MAPTMS and 0.1M hydrochloric acid were combined at a molar ratio of 385:1, and the resulting solution was mixed for 40 min at room temperature. Simultaneously, methacrylic acid and 70% zirconium n-propoxide were mixed at a molar ratio of 10:7 (MAA/ZPO) for 40 min at room temperature. The two solutions were then combined and mixed for an additional 40 min. Water was then added to achieve a final MAPTMS/H2O molar ratio of approximately 1:2. The resulting solution was mixed an additional 40 min. The final MAPTMS:ZPO molar ratio of the resin was approximately 6:4. The photoinitiator 4,40 -Bis(diethylamino) benzophenone (Sigma-Aldrich, St. Louis, MO) also known as ethyl Michler’s ketone (Fig. 1) was added to the solution at 1% w/w concentration and was mixed for approximately 20–30 min. The mixture was filtered with a 0.22 lm syringe filter to remove any polymerized material and undissolved particulates. The final step of resin synthesis involved further condensation using heat treatment. During this step, solvents were removed and gelation took place. The zirconium oxide hybrid material was drop-cast onto glass substrates. Approximately 50 ll of resin was deposited on 10 mm round glass coverslips, resulting in complete surface coverage. The resulting films were baked using a 1 h ramp time to 100 C and cured for 2 h at this temperature to form a hard gel for subsequent photopolymerization by 2PP. B. Two-photon polymerization and postprocessing of scaffolds
A frequency doubled (Yb:KGW) femtosecond laser (FLINT oscillator) (Light Conversion Ltd., Vilnius, Lithuania) with a center emission wavelength at 515 nm was applied for two-photon polymerization-based fabrication of tissue
II. EXPERIMENT A. Zirconium oxide hybrid material synthesis and film preparation
The zirconium oxide hybrid material is a photosensitive sol–gel material that was prepared using a procedure similar to that previously described by Ovsianikov et al.30,32,33 The zirconium oxide hybrid material was synthesized from methacryloxypropyl trimethoxysilane (MAPTMS) (SigmaAldrich, St. Louis, MO), methacrylic acid (MAA), and Biointerphases, Vol. 9, No. 2, June 2014
FIG. 1. Chemical structures of the inorganic–organic zirconium oxide hybrid material components and the photoinitiator used for two-photon polymerization.
029014-3 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
engineering scaffolds composed of the zirconium oxide hybrid material. The laser system delivered pulses of 60 fs duration at a repetition rate of 75.6 MHz. The beam was passed through an ND4 neutral density filter, and average power was further modulated using a quarter wave plate-polarizing beam splitter pair. An acousto-optical modulator was applied to trigger exposure of the sample. The beam is then passed through an EC Plan-Neofluar 20 objective (Zeiss, NA ¼ 0.5) and focused into the sample. The sample was translated perpendicular (xand y-directions) to the beam path by a set of linear stages (Aerotech, Inc., Pittsburgh, PA), while the laser focus height was controlled by moving the microscope objective in the zdirection using a third identical stage. The sample was mounted on an aluminum platform which was leveled relative to the stages using a VM1 kinematic mount (Thorlabs, Newton, NJ). Process observation was achieved by mounting a CCD camera behind a dichroic mirror and by attaching a red light-emitting diode (LED) to the mount for illumination. Eight samples of four different scaffold designs were produced of parallel lines to form scaffolds exhibiting pore sizes in the range of 10–20 lm. The scaffold design was chosen to provide a simplistic model exhibiting pore sizes within the ideal pore size distribution for cell attachment and alignment of cell growth for bone tissue engineering (1–20 lm).34 Each line of the scaffolds were fabricated using a single line scan using 2PP. Scaffold #1 contains one set of parallel lines spaced 20 lm apart. Scaffold #2 contains a set of perpendicular lines to make a square pattern, which covers the surface. Scaffold #3 contains the features associated with scaffold #2 as well as one set of diagonal lines that is located at angle of 45 with respect to the set of perpendicular lines. Scaffold #4 includes a second diagonal set that is perpendicular to the first diagonal set (i.e., the set of diagonal lines associated with scaffold #3). The additional lines incorporated into scaffolds #3 and #4 are located 14.14 lm apart and are aligned diagonally to the structures of scaffold #2; intersections between the diagonal lines and the perpendicular lines are incorporated within the design. The different scaffold designs exhibited various interconnecting pore sizes and three-dimensionality. The laser focus was aligned with the glass–polymer interface by searching for the point where an approximately 100 ms flash would produce an observable polymerized dot. The laser focus was moved 10 lm upward into the glass at this point; the voxel dimensions for the planned processing parameters would ideally result in 20 lm tall lines at this depth. 2PP processing proceeded in a raster pattern across the resin where each line is drawn across the surface. Each line was produced with a single scan produced by moving the stages at the chosen angle (i.e., 0 , 90 , 45 , 135 ) and at a linear speed of 5 mm/s; the sample was exposed to 1 mW of laser power measured before the objective. At the end of the line, the stages would move either 20 or 14.1 lm in a perpendicular manner with respect to the previously drawn line before starting the next line. One set of parallel lines was entirely produced before proceeding to the next set of parallel lines Biointerphases, Vol. 9, No. 2, June 2014
029014-3
(i.e., a full set of parallel lines oriented at 0 was finished before a full set of parallel lines at 90 was started). After 2PP was complete, the structures were developed by immersion in 1-propanol. Nonpatterned zirconium oxide hybrid samples were prepared on the 10 mm glass coverslips. These structures were used as control materials for the cell studies. Fabrication of these samples included drop-casting 500 ll of the zirconium oxide hybrid resin onto the glass. The samples were then baked with a 1 h ramp time to 100 C; this temperature was held overnight. Photopolymerization was performed using UV illumination at a wavelength of 255 nm and 8 W total power for a duration of 24 h. The 2PP-fabricated scaffolds as well as the drop-cast controls were then soaked in 100% ethanol for 1 week. The solvent was replaced twice during this duration to ensure complete removal of unpolymerized material and excess photoinitiator. C. Physicochemical characterization: SEM, AFM, and EDS
Scanning electron microscopy (SEM) was used to evaluate the zirconium oxide hybrid material scaffold architecture. Secondary electron images (SEI) of the zirconium oxide hybrid material 2PP scaffolds were obtained using a JEOL JSM-6390LV scanning electron microscope (JEOL, Tokyo, Japan). An acceleration voltage of 0.5–1.0 keV and a working distance of 10 mm were used for image acquisition. Atomic force microscopy (AFM) analysis of zirconium oxide hybrid material scaffolds was obtained using a MFP3-D microscope (Asylum Research, Santa Barbara, CA). A NCH type silicon noncontact high resonance frequency cantilever (spring constant 42 N/m) (NanoWorld, Neuch^atel, Switzerland) was used for tapping mode AFM imaging in air. The scan rate was set to 0.3–0.5 Hz at various scan size, and the images were collected using a resolution of 512 pixels 512 lines. First-order flattening of AFM height profiles was applied to remove Z offset of scan lines. The areas with higher Z profile were masked from flattening to minimize the influence of image processing artifacts. Elemental analysis of the zirconium oxide hybrid material scaffolds was obtained using energy dispersive x-ray spectroscopy (EDS). A 6733 A-1NUS-SN EDS spectrometer (Thermo Fisher, Waltham, MA) was used in this study. An acceleration voltage of 15 kV was utilized for EDS analysis. The spot size was optimized to keep the dead time around 25%–30% for point shoot EDS analysis. Spectral data were acquired for 30 s. D. Cell culture and cell seeding of scaffolds
Primary human bone marrow stromal cells (Lonza, Basel, Switzerland) were cultured under cell culture conditions (37 C, 5% CO2) in alpha-minimum essential medium (a-MEM) (Life TechnologiesTM, Carlsbad, CA) supplemented with 16% fetal bovine serum (FBS) (Atlanta Biologicals, Flowery Branch, GA), 1% penicillin, streptomycin, neomycin antibiotic mixture (PSN) (Gibco, Carlsbad,
029014-4 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
CA), and 1%–2% L-glutamine (Gibco, Carlsbad, CA). Once the hBMSCs reached approximately 80% confluency, cells were dissociated with 0.25% mass fraction trypsin and resuspended in medium prior to cell counting. Passage 3 cells were used for all experiments. All four scaffold types, zirconium oxide hybrid material drop-cast samples, glass coverslides, and wells without scaffolds (tissue culture polystyrene controls) were used for cell culture experiments. Prior to cell seeding, all of the samples were sterilized in 100% ethanol for 30 min and rinsed three times in sterile 1 PBS (phosphate-buffered saline, 30 min/ rinse). Samples were then incubated for 30 min at room temperature in 1 GibcoV PSN antibiotic mixture, rinsed in sterile PBS, and air-dried overnight. Scaffolds and control samples were placed in 24-well tissue culture polystyrene plates for cell culture experiments. hBMSCs were carefully seeded on the scaffolds at a concentration of 400 000 cells/ml in a 0.05 ml droplet; seeding 20 000 cells per scaffold was achieved using this approach. Cells were allowed to adhere to the scaffold for 1 h incubation before carefully adding 1 ml of fresh medium to each well. Three days following initial cell seeding, osteogenic supplements (Sigma-Aldrich, St. Louis, MO) were added to the culture medium such as 10 nM of dexamethasone, 0.05 mM of ascorbic acid, and 20 mM of b-glycerophosphate. The medium with osteogenic supplements was changed two to three times per week with fresh media that was prepared weekly. hBMSCs were cultured for 45 days on the scaffolds and on the control substrates. R
E. Preparation and SEM imaging of cells on scaffolds
After 45 day cell culture, the cell–scaffold and cell–control samples were rinsed with sterile PBS and fixed with 1% volume fraction of glutaraldehyde for 1 h at room temperature. Samples were then subjected to a graded alcohol dehydration (10%, 25%, 50%, 70%, 95%, 100% ethanol) in 10 min intervals and immersed in hexamethyldisilazane (HMDS) for 10 min. After removal from HMDS, samples were air-dried overnight. Samples were then sputter coated with gold prior to SEM imaging.
029014-4
oxide-polymer hybrid materials, including photopolymerizable materials, have been previously evaluated for use as dental composites.38,39 Scaffolds with microarchitectures were successfully fabricated from zirconium oxide hybrid material using two-photon polymerization. Using 2PP, four gridlike structure designs were fabricated based on lines with height of 14–20 lm and width of 1.7–2 lm. The scaffolds incorporated different layers of parallel lines at various orientations to create microporous structures. Scanning electron micrographs of the scaffolds (Fig. 2) demonstrate the different geometries of the four scaffold types and the uniform nature of the 2PP-fabricated structures. Scaffold #1 [Fig. 2(a)] exhibits parallel ridges with 20 lm spacing. Fragments of material are visible between the ridges; it is hypothesized that these fragments resulted from fracture of the scaffold ridge walls. Improvements to the scaffold design may serve to minimize this phenomenon. Scaffold #2 [Fig. 2(b)] incorporates horizontal and parallel ridges that are oriented perpendicular to one another. A square, gridlike structure with 20 lm spacing is formed by the intersection of the ridges. The perpendicular ridges in scaffold #2 provide a much more stable structure. Scaffold #3 [Fig. 2(c)] has a square gridlike structure with additional diagonal lines that are oriented at 45 and exhibit 14 lm spacing. The diagonal ridges are well aligned and intersect with the cross sections of the lower grid throughout the scaffold. Scaffold #4 [Fig. 2(d)] exhibits the same features as scaffold #3 and also exhibits an additional layer of diagonal lines that are perpendicular to the layer seen in scaffold #3. The orientation of the set of upper diagonal ridges exhibits some minor variations from the computer design; for example, the overlapping portions of the layers do not align perfectly throughout the structure.
III. RESULTS AND DISCUSSION A. Two-photon polymerization of zirconium oxide hybrid material scaffolds
Zirconium oxide is a biocompatible ceramic that provides high strength, stiffness, and hardness, making it an ideal material for hard tissue applications.35 Due to these material properties, zirconium oxide is a common component of orthopedic35,36 and dental implants.36,37 In the present study, we fabricated scaffolds composed of zirconium oxide hybrid materials that combine the unique properties of the organic polymer components and the zirconium oxide inorganic ceramic material. The organic polymers enable photopolymerization of the material by two-photon polymerization, while the zirconium oxide contributes to the biocompatibility and the mechanical stability of the scaffold. Similar zirconium Biointerphases, Vol. 9, No. 2, June 2014
FIG. 2. Scanning electron microscopy images (400 magnification) of the 3-D zirconium oxide hybrid material scaffolds fabricated using two-photon polymerization.
029014-5 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
Atomic force microscopy was further used to evaluate the scaffold geometries and to obtain quantitative information on the scaffold dimensions. 3-D representative images of the scaffolds obtained by AFM are shown in Fig. 3(a). These micrographs further demonstrate the uniform nature of the biograting patterns and indicate the presence of peaklike nodules at the overlapping regions of the ridges on scaffold #2, scaffold #3, and scaffold #4. It should be noted that the 3-D images do not accurately portray the scaffold architecture since the height of the scaffold structures and the overlapping nature of the designs limited the image acquisition process; this phenomenon is particularly relevant for scaffold #3 and scaffold #4. AFM was also used to measure the ridge height of scaffold #1 as well as the ridge spacing of scaffold #2; a ridge height of 14.2 6 0.5 lm was noted for scaffold #1 and a ridge spacing of 20.1 6 0.3 lm was noted for scaffold #2 [Fig. 3(b)]. The structure heights were notably lower than the 20 lm target since fidelity between the design and the features in the fabricated structure was traded for a reduction in the 2PP fabrication time. Objects of this height are usually fabricated
FIG. 3. (Color online) Representative AFM 3-D images of two-photon polymerization-fabricated scaffolds (a). Scaffold structural measurements obtained using atomic force microscopy, showing a ridge height of 14.2 lm for scaffold #1 and a pitch of 20.1 lm for scaffold #2 (b). Biointerphases, Vol. 9, No. 2, June 2014
029014-5
using at least three separate layers and are processed using a lower average laser power. The peaks observed at the intersections and the overlapping lines in scaffold #3 and scaffold #4 are indicative of lensing effects that occur whenever the laser passes over a previously polymerized spot. It is anticipated that these artifacts may be minimized by creating structures using multiple layers. The persistence of the structures after development in 1-propanol and soaking in ethanol demonstrates the stability of the zirconium oxide hybrid material after photopolymerization and postprocessing. The shrinkage properties of 2PP-based structures formed from this zirconium hybrid resin has been previously demonstrated by Ovsianikov et al.33 By optimizing the laser parameters, zirconium oxide hybrid material structures may be fabricated by 2PP without significant shrinkage, facilitating good fidelity between the 2PP-fabricated structure features and the computer design on which they were based. The elemental analysis of the zirconium oxide hybrid material scaffolds by EDS (Fig. 4) showed the representative peaks for the inorganic and organic constituents, including carbon, oxygen, silicon, and zirconium. The characteristic Zr La peak is visible at 2.042 keV on the EDS spectrum [Fig. 4(a)]. Prior to cell culture, samples were postprocessed using an alcohol soaking procedure for 1 week to remove excess monomers and photoinitiator; these components of 2PPfabricated samples may reduce cell viability.33 The lack of a nitrogen peak in the EDS spectrum suggests the removal of photoinitiator 4,40 -Bis(diethylamino) benzophenone. The EDS scan [Fig. 4(b)] mapping of Zr, O, and C on scaffold #2 [Fig. 4(c)] illustrates the presence of elements associated with the zirconium oxide hybrid material in the structure patterned by 2PP.
FIG. 4. (Color online) EDS spectrum (a), line scan (b), and mapping of a scaffold fabricated out of the inorganic–organic zirconium oxide hybrid material by 2PP.
029014-6 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
029014-6
B. Cellular response to 2PP-fabricated zirconium oxide hybrid material microstructured scaffolds
Since zirconium oxide is commonly utilized in orthopedic implants and dental implants, cellular response to the scaffolds was evaluated using hBMSCs. These nonhematopoietic progenitor cells can differentiate into musculoskeletal tissues that come in contact with a variety of implantable medical devices. Long-term tissue culture studies were conducted to evaluate the cellular attachment, orientation, and growth on the 3-D microstructures and to determine the potential of these scaffolds for long-term 3-D cell culture of stem cells for tissue engineering applications. For evaluation of cell interactions with the 2PP fabricated zirconium-based scaffolds, hBMSCs were seeded on the scaffolds at a density of 20 000 cells/scaffold and cultured for 45 days in the presence of osteogenic supplements. The scaffolds maintained their structure with minimal distortion or degradation, demonstrating their stability in a physiologically relevant environment. The cells adhered to all of the scaffold types and remained viable for up to 45 days, forming a dense, tissuelike matrix that bridged across the ridges of the scaffolds. SEM images of this tissue matrix on the scaffolds show that cell attachment conforms to the topographical features of the scaffolds (Fig. 5). Comparison of
FIG. 6. SEM images of hBMSCs on scaffold #2, demonstrating cellular attachment (a) and ingrowth within the scaffold microarchitecture (b).
the scaffolds with cells and without cells illustrates that tissue matrix was formed by cells; the peaklike nodules at the ridge joints and the ridge walls served as the initial cell attachment sites. Evenly distributed cell seeding and/or cell growth is evident in the formation of the tissuelike matrix throughout the entire surface of the scaffolds. High magnification SEM images of the tissue matrix reveal the spatial alignment of cells on the scaffolds; the relationship between cell alignment and ridge orientation varied for each scaffold design. For scaffolds #1, scaffold #2, and scaffold #3, the cells align diagonally to the uppermost scaffold ridges. For scaffold #4, the cells align parallel to the top ridge layer. Cells on the 2-D drop-cast zirconium oxide hybrid material control films and cells on the glass surfaces formed a dense cell layer, though no specific orientation was observed. These results demonstrate cell alignment that is dependent on the 3-D scaffold geometry. Cells were less densely grown in some regions toward the edge of scaffold #2 (Fig. 6). Individual cells can be seen spanning across and adhering to the walls of the grid. Cellular projections of the hBMSCs are visible, extending across the gaps between the scaffold ridges. Figure 6(b) shows a high magnification image of scaffold #2. In this image, cells have infiltrated the scaffold, stretching and forming adhesions on the scaffold in a three-dimensional fashion. Significant cell attachment and proliferation is evidenced by infiltration of hBMSCs into the interior of the scaffold and formation of a tissuelike matrix on the 2PPfabricated 3-D scaffolds.
IV. SUMMARY AND CONCLUSIONS
FIG. 5. SEM images (500 magnification) of hBMSCs grown on glass control substrate (a), drop-cast film of zirconium oxide hybrid material (b), scaffold #1 (c), scaffold #2 (d), scaffold #3 (e), and scaffold #4 (f) after 45 day cell culture. Biointerphases, Vol. 9, No. 2, June 2014
Four types of scaffold microarchitectures were fabricated out of a zirconium oxide hybrid material using two-photon polymerization. Scanning electron microscopy and atomic force microscopy demonstrated that the scaffolds exhibit uniform dimensions that correspond closely with the computer designs used in scaffolding processing. Energy dispersive x-ray spectroscopy was used to confirm presence of elements that are consistent with the organic and inorganic moieties in the fabricated structure and the absence of residual photoinitiator; no unanticipated impurities were noted in the 2PP-fabricated structures. Multiple identical samples of
029014-7 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
each scaffold design were fabricated, demonstrating the reproducibility of the two-photon polymerization process. Long-term cell culture studies using hBMSCs showed cellular adhesion and cellular alignment on all scaffold microarchitectures; a dense, tissuelike structure was noted to span the gaps of the porous scaffolds. The cell orientation was shown to be dependent on the microarchitectures of the scaffolds. This study demonstrates the potential of zirconium oxide hybrid material-based 3-D microarchitectures for long-term growth of hBMSCs. These microstructured materials may be used as a model for development of a 3-D tissue culture system and as a valuable tool for bone tissue engineering. ACKNOWLEDGMENTS The authors would like to acknowledge the FDA White Oak Nanotechnology Core Facility for instrument use and scientific and technical assistance. In addition, the authors would like to acknowledge financial support from U.S. National Science Foundation Award #1041375. The authors would also like to thank the Laser Zentrum Hannover (LZH) in Hannover, Germany for use of their two-photon polymerization system for fabrication procedures. A.K. and B.N.C. acknowledge support by the German Research Foundation Cluster of Excellence Rebirth “From Regenerative Biology to Reconstructive Therapy” and the Lower Saxony project “Biofabrication for NIFE.” The mention of commercial products, their sources, or their use in connection with material reported herein is not to be construed as either an actual or implied endorsement of such products by the Department of Health and Human Services. 1
V. Aprile, S. M. Eaton, M. Lagana, G. Cerullo, M. T. Raimondi, and R. Osellame, Proc. SPIE 8247, 824708 (2012). 2 O. Z. Fisher, A. Khademhosseini, R. Langer, and N. A. Peppas, Acc. Chem. Res. 43, 419 (2010). 3 A. M. Greiner, B. Richter, and M. Bastmeyer, Macromol. Biosci. 12, 1301 (2012). 4 H. Jeon, H. Hidai, D. J. Hwang, and C. P. Grigoropoulos, J. Biomed. Mater. Res., Part A 93A, 56 (2009). 5 P. J. Rodrigo, L. Kelemen, D. Palima, C. A. Alonzo, P. Ormos, and J. Gluckstad, Opt. Express 17, 6578 (2009). 6 H. Jeon, H. Hidai, D. J. Hwang, K. E. Healy, and C. P. Grigoropoulos, Biomaterials 31, 4286 (2010). 7 M. Licht, A. Uchugonova, K. Konig, and M. Straub, J. Opt. 14, 065601 (2012). 8 E. Cukierman, R. Pankov, and K. M. Yamada, Curr. Opin. Cell Biol. 14, 633 (2002). 9 D. J. Maltman and S. A. Przyborski, Biochem. Soc. Trans. 38, 1072 (2010). 10 E. Kaivosoja, P. Suvanto, G. Barreto, S. Aura, A. Soininen, S. Franssila, and Y. T. Konttinen, J. Biomed. Mater. Res., Part A 101A, 842 (2013).
Biointerphases, Vol. 9, No. 2, June 2014
11
029014-7
W. A. Loesberg, J. te Riet, F. C. M. J. M. van Delft, P. Schon, C. G. Figdor, S. Speller, J. J. W. A. van Loon, X. F. Walboomers, and J. A. Jansen, Biomaterials 28, 3944 (2007). 12 A. L. Teixeria, G. A. McKie, J. D. Foley, P. J. Bertics, P. F. Nealey, and C. J. Murphy, Biomaterials 27, 3945 (2006). 13 B. J. Papenburg, L. Vogelaar, L. A. M. Bolhuis-Versteeg, R. G. H. Lammertink, D. Stamatialis, and M. Wessling, Biomaterials 28, 1998 (2007). 14 J. P. Kaiser, A. Reinmann, and A. Bruinink, Biomaterials 27, 5230 (2006). 15 P. Tayalia, E. Mazur, and D. J. Mooney, Biomaterials 32, 2634 (2011). 16 Y. Lu, G. Mapili, G. Suhali, S. Chen, and K. Roy, J. Biomed. Mater. Res. A 77A, 396 (2006). 17 O. Zinger, G. Zhao, Z. Schwartz, J. Simpson, M. Wieland, D. Landolt, and B. Boyan, Biomaterials 26, 1837 (2005). 18 S. A. Ruiz and C. S. Chen, Stem Cells 26, 2921 (2008). 19 G. Kumar, C. K. Tison, K. Chatterjee, P. S. Pine, J. H. McDaniel, M. L. Salit, M. F. Young, and C. G. Simon, Jr., Biomaterials 32, 9188 (2011). 20 Y. Seol, D. Y. Park, J. Y. Park, S. W. Kim, S. J. Park, and D. Cho, Biotechnol. Bioeng. 110, 1444 (2013). 21 C. H. Thomas, J. H. Collier, C. S. Sfeir, and K. E. Healy, Proc. Natl. Acad. Sci. U.S.A. 99, 1972 (2002). 22 H. Hosseinkhani, M. Hosseinkhani, and H. Kobayashi, J. Bioact. Compat. Polym. 21, 277 (2006). 23 C. S. Ki, S. Y. Park, H. J. Kim, H. M. Jung, K. M. Woo, J. W. Lee, and Y. H. Park, Biotechnol. Lett. 30, 405 (2008). 24 R. J. Narayan, A. Doraiswamy, D. B. Chrisey, and B. N. Chichkov, Mater. Today 13, 42 (2010). 25 R. Schade, T. Weiss, A. Berg, M. Schnabelrauch, and K. Liefeith, Int. J. Artif. Organs 33, 219 (2010). 26 T. Weiss, G. Hildebrand, R. Schade, and K. Liefeith, Eng. Life Sci. 9, 384 (2009). 27 S. Englehardt, E. Hoch, K. Borchers, W. Meyer, H. Kruger, G. E. M. Tovar, and A. Gillner, Biofabrication 3, 025003 (2011). 28 S. Psycharakis, A. Tosca, V. Melissinaki, A. Giakoumaki, and A. Ranella, Biomed. Mater. 6, 045008 (2011). 29 T. M. Hsieh, C. W. B. Ng, K. Naryanan, A. C. A. Wan, and J. Y. Ying, Biomaterials 31, 7648 (2010). 30 A. Ovsianikov et al., Laser Chem. 2008, 1. 31 A. Ovsianikov et al., ACS Nano 2, 2257 (2008). 32 A. Ovsianikov, X. Shizhou, M. Farsari, M. Vamvakaki, C. Fotakis, and B. N. Chichkov, Opt. Express 17, 4 (2009). 33 A. Ovsianikov et al., Acta Biomater. 7, 967 (2011). 34 M. Ramalingam, E. Jabbarl, S. Ramakrishna, and A. Khademhosseini, Micro and Nanotechnologies in Engineering Stem Cells and Tissues (IEEE, Piscataway, NJ, 2013), pp. 85–86. 35 C. Piconi and G. Maccauro, Biomaterials 20, 1 (1999). 36 G. Maccauro, P. R. Iommetti, L. Raffaelli, and P. F. Manicone, “Alumina and zirconia ceramics for orthopaedic and dental devices,” in Biomaterials Applications for Nanomedicine, edited by R. Pignatello (InTech, Rijeka, Croatia, 2011), Chap. 5, pp. 300–307. 37 C. A. M. Volpato, L. G. D. Garbelotto, M. C. Fredel, and F. Bandioli, “Application of zirconia in dentistry: Biological, mechanical, and optical considerations,” in Advances in Ceramics—Electric and Magnetic Ceramics, Bioceramics, Ceramics and Environment, edited by C. Sikalidis (InTech, Rijeka, Croatia, 2011), Chap. 17, pp. 397–404. 38 S. Bayou, M. Mouzali, F. Aloui, L. Lecamp, and P. Lebaudy, Polym. J. 45, 863 (2013). 39 K. S. Chan, D. P. Nicolella, B. R. Furman, S. T. Wellinghoff, H. R. Rawls, and S. E. Pratsinis, J. Mater. Sci. 44, 22 (2009).
Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds for long-term stem cell growth Shelby A. Skoog Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965 and Office of Science and Engineering Laboratories, Center for Devices and Radiological Health, U.S. Food and Drug Administration, Silver Spring, Maryland 20993
Alexander K. Nguyen Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965 and Department of Nanotechnology, Laser Zentrum Hannover e. V., Hannover D-30419, Germany
Girish Kumar, Jiwen Zheng, and Peter L. Goering Office of Science and Engineering Laboratories, Center for Devices and Radiological Health, U.S. Food and Drug Administration, Silver Spring, Maryland 20993
Anastasia Koroleva and Boris N. Chichkov Department of Nanotechnology, Laser Zentrum Hannover e. V., Hannover D-30419, Germany
Roger J. Narayana) Joint Department of Biomedical Engineering, University of North Carolina at Chapel Hill and North Carolina State University, Raleigh, North Carolina 27965
(Received 4 March 2014; accepted 17 April 2014; published 28 April 2014) Two-photon polymerization is a technique that involves simultaneous absorption of two photons from a femtosecond laser for selective polymerization of a photosensitive material. In this study, two-photon polymerization was used for layer-by-layer fabrication of 3-D scaffolds composed of an inorganic–organic zirconium oxide hybrid material. Four types of scaffold microarchitectures were created, which exhibit layers of parallel line features at various orientations as well as pores between the line features. Long-term cell culture studies involving human bone marrow stromal cells were conducted using these 3-D scaffolds. Cellular adhesion and proliferation were demonstrated on all of the scaffold types; tissuelike structure was shown to span the pores. This study indicates that two-photon polymerization may be used to create microstructured scaffolds out of an inorganic–organic zirconium oxide hybrid material for use in 3-D tissue culture systems. C 2014 American Vacuum Society. [http://dx.doi.org/10.1116/1.4873688] V I. INTRODUCTION Cellular response is strongly influenced by the surrounding biological environment through complex interactions with biochemical stimuli, mechanical forces, and structural components of the extracellular matrix (ECM).1–3 The threedimensional architecture of the ECM consists of interwoven fibrillar proteins embedded within a network of proteoglycans; this structure provides mechanical and physical cues that regulate cellular function. Since it is well known that cells are highly influenced by the local microenvironment, it is important to understand the underlying mechanisms of the cell–structure relationship.4–7 To evaluate the effect of ECM structure on cellular response, researchers employ micro- and nanoscale topographies as well as complex 3-D architectures that mimic the native cell environment. Utilizing 3-D structures for advanced cell culture and tissue engineering provides a more physiologically relevant model of the in vivo environment than 2-D structures. 2-D surfaces may confine cells to a single plane of growth; in addition, they may limit cells from a)
Author to whom correspondence should be addressed; electronic mail: [email protected]
029014-1 Biointerphases 9(2), June 2014
exhibiting their natural morphologies and cell–cell interactions. In contrast, 3-D scaffolds may provide porous microarchitectures similar to those of the natural ECM and may impart physical as well as chemical cues.8,9 Several studies have utilized both textured surfaces and 3-D scaffolds with physiologically relevant features to evaluate cell adhesion,10–12 morphology,13 alignment,13 migration,14–16 differentiation,16–19 proliferation,20 protein expression,21 and gene expression.21 Further, studies have demonstrated increased cell attachment, proliferation, and differentiation of bone-derived cells on 3-D scaffolds compared to 2-D surfaces.22,23 Understanding how cells respond to structural cues in 3-D will contribute to development of advanced biomaterials for improved implants, enhanced in vitro tissue models, and optimized scaffolds for tissue regeneration. For a systematic approach to assess cellular responses to surface topographies and 3-D architectures, materials with micro- and nanoscale features must be fabricated using precise, reproducible methods. Advances in microfabrication technology have enabled production of complex structures at high resolution for a variety of biomaterials. Two-photon polymerization (2PP) is a rapid prototyping technique that enables fabrication of 3-D structures with high spatial resolution
1934-8630/2014/9(2)/029014/7/$30.00
C 2014 American Vacuum Society V
029014-1
029014-2 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
down to 100 nm.24 The process uses the two-photon effect, which involves almost simultaneous absorption of two photons from a femtosecond laser to polymerize a photosensitive polymer resin in a layer-by-layer fashion. In 2PP, an ultrashort-pulsed laser beam is highly focused on a small, three-dimensional unit volume (voxel) of liquid resin. Since the two-photon effect is confined to this small volume, the chemical reactions between the photoinitiator molecules and the monomers (and subsequent polymerization of the material) are localized.24 2PP enables fabrication of structures with complex geometries and precise dimensions. A wide variety of photosensitive polymers, fillers, and photoinitiators have been utilized with 2PP; as such, fabrication of materials with a wide range of surface chemistries and mechanical properties is possible using this approach. This versatility in material properties and high fidelity between computer design and 2PP-fabricated structures may facilitate processing of more accurate models of the biophysical environment. Several researchers have utilized 2PP for fabrication of 3-D microarchitectures and textured surfaces to examine cellular activity. Studies have incorporated 2PP structures with various geometries and dimensions to evaluate cell adhesion,1,25 morphology,4,26,27 orientation,4,26,27 viability,28 proliferation,28 and cell functionality.29 Recently, cell response has been evaluated using 2PP-fabricated structures composed of photosensitive inorganic-organic hybrid copolymers, such as the commercially available ORMOCERV material (Microresist Technology GmbH, Berlin, Germany).4,26,30 ORMOCER material is synthesized using a sol–gel process; the final material contains cross-linked networks of inorganic and organic components. This hybrid sol–gel material exhibits high thermal stability, high chemical stability, good mechanical properties, and biocompatibility.31 In this study, we demonstrate reproducible fabrication of 3-D scaffolds composed of an inorganic–organic zirconium oxide hybrid material using two-photon polymerization. Four different scaffold microarchitectures were produced with different layers of parallel lines at various orientations. Long-term tissue culture studies were conducted using human bone marrow stromal cells (hBMSCs) to evaluate cellular attachment, alignment, and growth on the 3-D microstructures and to assess the potential of two-photon polymerization technology for long term 3-D stem cell culture for enhanced tissue engineering applications. R
029014-2
zirconium n-propoxide (ZPO). The chemical structures for the inorganic–organic hybrid copolymer components are shown in Fig. 1. Both organic monomers, MAPTMS and methacrylic acid, contain methacrylate moieties that are photopolymerizable. The inorganic network components include alkoxysilane groups of MAPTMS and zirconium npropoxide. During synthesis, the resin precursors first underwent catalytic hydrolysis and condensation. MAPTMS and 0.1M hydrochloric acid were combined at a molar ratio of 385:1, and the resulting solution was mixed for 40 min at room temperature. Simultaneously, methacrylic acid and 70% zirconium n-propoxide were mixed at a molar ratio of 10:7 (MAA/ZPO) for 40 min at room temperature. The two solutions were then combined and mixed for an additional 40 min. Water was then added to achieve a final MAPTMS/H2O molar ratio of approximately 1:2. The resulting solution was mixed an additional 40 min. The final MAPTMS:ZPO molar ratio of the resin was approximately 6:4. The photoinitiator 4,40 -Bis(diethylamino) benzophenone (Sigma-Aldrich, St. Louis, MO) also known as ethyl Michler’s ketone (Fig. 1) was added to the solution at 1% w/w concentration and was mixed for approximately 20–30 min. The mixture was filtered with a 0.22 lm syringe filter to remove any polymerized material and undissolved particulates. The final step of resin synthesis involved further condensation using heat treatment. During this step, solvents were removed and gelation took place. The zirconium oxide hybrid material was drop-cast onto glass substrates. Approximately 50 ll of resin was deposited on 10 mm round glass coverslips, resulting in complete surface coverage. The resulting films were baked using a 1 h ramp time to 100 C and cured for 2 h at this temperature to form a hard gel for subsequent photopolymerization by 2PP. B. Two-photon polymerization and postprocessing of scaffolds
A frequency doubled (Yb:KGW) femtosecond laser (FLINT oscillator) (Light Conversion Ltd., Vilnius, Lithuania) with a center emission wavelength at 515 nm was applied for two-photon polymerization-based fabrication of tissue
II. EXPERIMENT A. Zirconium oxide hybrid material synthesis and film preparation
The zirconium oxide hybrid material is a photosensitive sol–gel material that was prepared using a procedure similar to that previously described by Ovsianikov et al.30,32,33 The zirconium oxide hybrid material was synthesized from methacryloxypropyl trimethoxysilane (MAPTMS) (SigmaAldrich, St. Louis, MO), methacrylic acid (MAA), and Biointerphases, Vol. 9, No. 2, June 2014
FIG. 1. Chemical structures of the inorganic–organic zirconium oxide hybrid material components and the photoinitiator used for two-photon polymerization.
029014-3 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
engineering scaffolds composed of the zirconium oxide hybrid material. The laser system delivered pulses of 60 fs duration at a repetition rate of 75.6 MHz. The beam was passed through an ND4 neutral density filter, and average power was further modulated using a quarter wave plate-polarizing beam splitter pair. An acousto-optical modulator was applied to trigger exposure of the sample. The beam is then passed through an EC Plan-Neofluar 20 objective (Zeiss, NA ¼ 0.5) and focused into the sample. The sample was translated perpendicular (xand y-directions) to the beam path by a set of linear stages (Aerotech, Inc., Pittsburgh, PA), while the laser focus height was controlled by moving the microscope objective in the zdirection using a third identical stage. The sample was mounted on an aluminum platform which was leveled relative to the stages using a VM1 kinematic mount (Thorlabs, Newton, NJ). Process observation was achieved by mounting a CCD camera behind a dichroic mirror and by attaching a red light-emitting diode (LED) to the mount for illumination. Eight samples of four different scaffold designs were produced of parallel lines to form scaffolds exhibiting pore sizes in the range of 10–20 lm. The scaffold design was chosen to provide a simplistic model exhibiting pore sizes within the ideal pore size distribution for cell attachment and alignment of cell growth for bone tissue engineering (1–20 lm).34 Each line of the scaffolds were fabricated using a single line scan using 2PP. Scaffold #1 contains one set of parallel lines spaced 20 lm apart. Scaffold #2 contains a set of perpendicular lines to make a square pattern, which covers the surface. Scaffold #3 contains the features associated with scaffold #2 as well as one set of diagonal lines that is located at angle of 45 with respect to the set of perpendicular lines. Scaffold #4 includes a second diagonal set that is perpendicular to the first diagonal set (i.e., the set of diagonal lines associated with scaffold #3). The additional lines incorporated into scaffolds #3 and #4 are located 14.14 lm apart and are aligned diagonally to the structures of scaffold #2; intersections between the diagonal lines and the perpendicular lines are incorporated within the design. The different scaffold designs exhibited various interconnecting pore sizes and three-dimensionality. The laser focus was aligned with the glass–polymer interface by searching for the point where an approximately 100 ms flash would produce an observable polymerized dot. The laser focus was moved 10 lm upward into the glass at this point; the voxel dimensions for the planned processing parameters would ideally result in 20 lm tall lines at this depth. 2PP processing proceeded in a raster pattern across the resin where each line is drawn across the surface. Each line was produced with a single scan produced by moving the stages at the chosen angle (i.e., 0 , 90 , 45 , 135 ) and at a linear speed of 5 mm/s; the sample was exposed to 1 mW of laser power measured before the objective. At the end of the line, the stages would move either 20 or 14.1 lm in a perpendicular manner with respect to the previously drawn line before starting the next line. One set of parallel lines was entirely produced before proceeding to the next set of parallel lines Biointerphases, Vol. 9, No. 2, June 2014
029014-3
(i.e., a full set of parallel lines oriented at 0 was finished before a full set of parallel lines at 90 was started). After 2PP was complete, the structures were developed by immersion in 1-propanol. Nonpatterned zirconium oxide hybrid samples were prepared on the 10 mm glass coverslips. These structures were used as control materials for the cell studies. Fabrication of these samples included drop-casting 500 ll of the zirconium oxide hybrid resin onto the glass. The samples were then baked with a 1 h ramp time to 100 C; this temperature was held overnight. Photopolymerization was performed using UV illumination at a wavelength of 255 nm and 8 W total power for a duration of 24 h. The 2PP-fabricated scaffolds as well as the drop-cast controls were then soaked in 100% ethanol for 1 week. The solvent was replaced twice during this duration to ensure complete removal of unpolymerized material and excess photoinitiator. C. Physicochemical characterization: SEM, AFM, and EDS
Scanning electron microscopy (SEM) was used to evaluate the zirconium oxide hybrid material scaffold architecture. Secondary electron images (SEI) of the zirconium oxide hybrid material 2PP scaffolds were obtained using a JEOL JSM-6390LV scanning electron microscope (JEOL, Tokyo, Japan). An acceleration voltage of 0.5–1.0 keV and a working distance of 10 mm were used for image acquisition. Atomic force microscopy (AFM) analysis of zirconium oxide hybrid material scaffolds was obtained using a MFP3-D microscope (Asylum Research, Santa Barbara, CA). A NCH type silicon noncontact high resonance frequency cantilever (spring constant 42 N/m) (NanoWorld, Neuch^atel, Switzerland) was used for tapping mode AFM imaging in air. The scan rate was set to 0.3–0.5 Hz at various scan size, and the images were collected using a resolution of 512 pixels 512 lines. First-order flattening of AFM height profiles was applied to remove Z offset of scan lines. The areas with higher Z profile were masked from flattening to minimize the influence of image processing artifacts. Elemental analysis of the zirconium oxide hybrid material scaffolds was obtained using energy dispersive x-ray spectroscopy (EDS). A 6733 A-1NUS-SN EDS spectrometer (Thermo Fisher, Waltham, MA) was used in this study. An acceleration voltage of 15 kV was utilized for EDS analysis. The spot size was optimized to keep the dead time around 25%–30% for point shoot EDS analysis. Spectral data were acquired for 30 s. D. Cell culture and cell seeding of scaffolds
Primary human bone marrow stromal cells (Lonza, Basel, Switzerland) were cultured under cell culture conditions (37 C, 5% CO2) in alpha-minimum essential medium (a-MEM) (Life TechnologiesTM, Carlsbad, CA) supplemented with 16% fetal bovine serum (FBS) (Atlanta Biologicals, Flowery Branch, GA), 1% penicillin, streptomycin, neomycin antibiotic mixture (PSN) (Gibco, Carlsbad,
029014-4 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
CA), and 1%–2% L-glutamine (Gibco, Carlsbad, CA). Once the hBMSCs reached approximately 80% confluency, cells were dissociated with 0.25% mass fraction trypsin and resuspended in medium prior to cell counting. Passage 3 cells were used for all experiments. All four scaffold types, zirconium oxide hybrid material drop-cast samples, glass coverslides, and wells without scaffolds (tissue culture polystyrene controls) were used for cell culture experiments. Prior to cell seeding, all of the samples were sterilized in 100% ethanol for 30 min and rinsed three times in sterile 1 PBS (phosphate-buffered saline, 30 min/ rinse). Samples were then incubated for 30 min at room temperature in 1 GibcoV PSN antibiotic mixture, rinsed in sterile PBS, and air-dried overnight. Scaffolds and control samples were placed in 24-well tissue culture polystyrene plates for cell culture experiments. hBMSCs were carefully seeded on the scaffolds at a concentration of 400 000 cells/ml in a 0.05 ml droplet; seeding 20 000 cells per scaffold was achieved using this approach. Cells were allowed to adhere to the scaffold for 1 h incubation before carefully adding 1 ml of fresh medium to each well. Three days following initial cell seeding, osteogenic supplements (Sigma-Aldrich, St. Louis, MO) were added to the culture medium such as 10 nM of dexamethasone, 0.05 mM of ascorbic acid, and 20 mM of b-glycerophosphate. The medium with osteogenic supplements was changed two to three times per week with fresh media that was prepared weekly. hBMSCs were cultured for 45 days on the scaffolds and on the control substrates. R
E. Preparation and SEM imaging of cells on scaffolds
After 45 day cell culture, the cell–scaffold and cell–control samples were rinsed with sterile PBS and fixed with 1% volume fraction of glutaraldehyde for 1 h at room temperature. Samples were then subjected to a graded alcohol dehydration (10%, 25%, 50%, 70%, 95%, 100% ethanol) in 10 min intervals and immersed in hexamethyldisilazane (HMDS) for 10 min. After removal from HMDS, samples were air-dried overnight. Samples were then sputter coated with gold prior to SEM imaging.
029014-4
oxide-polymer hybrid materials, including photopolymerizable materials, have been previously evaluated for use as dental composites.38,39 Scaffolds with microarchitectures were successfully fabricated from zirconium oxide hybrid material using two-photon polymerization. Using 2PP, four gridlike structure designs were fabricated based on lines with height of 14–20 lm and width of 1.7–2 lm. The scaffolds incorporated different layers of parallel lines at various orientations to create microporous structures. Scanning electron micrographs of the scaffolds (Fig. 2) demonstrate the different geometries of the four scaffold types and the uniform nature of the 2PP-fabricated structures. Scaffold #1 [Fig. 2(a)] exhibits parallel ridges with 20 lm spacing. Fragments of material are visible between the ridges; it is hypothesized that these fragments resulted from fracture of the scaffold ridge walls. Improvements to the scaffold design may serve to minimize this phenomenon. Scaffold #2 [Fig. 2(b)] incorporates horizontal and parallel ridges that are oriented perpendicular to one another. A square, gridlike structure with 20 lm spacing is formed by the intersection of the ridges. The perpendicular ridges in scaffold #2 provide a much more stable structure. Scaffold #3 [Fig. 2(c)] has a square gridlike structure with additional diagonal lines that are oriented at 45 and exhibit 14 lm spacing. The diagonal ridges are well aligned and intersect with the cross sections of the lower grid throughout the scaffold. Scaffold #4 [Fig. 2(d)] exhibits the same features as scaffold #3 and also exhibits an additional layer of diagonal lines that are perpendicular to the layer seen in scaffold #3. The orientation of the set of upper diagonal ridges exhibits some minor variations from the computer design; for example, the overlapping portions of the layers do not align perfectly throughout the structure.
III. RESULTS AND DISCUSSION A. Two-photon polymerization of zirconium oxide hybrid material scaffolds
Zirconium oxide is a biocompatible ceramic that provides high strength, stiffness, and hardness, making it an ideal material for hard tissue applications.35 Due to these material properties, zirconium oxide is a common component of orthopedic35,36 and dental implants.36,37 In the present study, we fabricated scaffolds composed of zirconium oxide hybrid materials that combine the unique properties of the organic polymer components and the zirconium oxide inorganic ceramic material. The organic polymers enable photopolymerization of the material by two-photon polymerization, while the zirconium oxide contributes to the biocompatibility and the mechanical stability of the scaffold. Similar zirconium Biointerphases, Vol. 9, No. 2, June 2014
FIG. 2. Scanning electron microscopy images (400 magnification) of the 3-D zirconium oxide hybrid material scaffolds fabricated using two-photon polymerization.
029014-5 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
Atomic force microscopy was further used to evaluate the scaffold geometries and to obtain quantitative information on the scaffold dimensions. 3-D representative images of the scaffolds obtained by AFM are shown in Fig. 3(a). These micrographs further demonstrate the uniform nature of the biograting patterns and indicate the presence of peaklike nodules at the overlapping regions of the ridges on scaffold #2, scaffold #3, and scaffold #4. It should be noted that the 3-D images do not accurately portray the scaffold architecture since the height of the scaffold structures and the overlapping nature of the designs limited the image acquisition process; this phenomenon is particularly relevant for scaffold #3 and scaffold #4. AFM was also used to measure the ridge height of scaffold #1 as well as the ridge spacing of scaffold #2; a ridge height of 14.2 6 0.5 lm was noted for scaffold #1 and a ridge spacing of 20.1 6 0.3 lm was noted for scaffold #2 [Fig. 3(b)]. The structure heights were notably lower than the 20 lm target since fidelity between the design and the features in the fabricated structure was traded for a reduction in the 2PP fabrication time. Objects of this height are usually fabricated
FIG. 3. (Color online) Representative AFM 3-D images of two-photon polymerization-fabricated scaffolds (a). Scaffold structural measurements obtained using atomic force microscopy, showing a ridge height of 14.2 lm for scaffold #1 and a pitch of 20.1 lm for scaffold #2 (b). Biointerphases, Vol. 9, No. 2, June 2014
029014-5
using at least three separate layers and are processed using a lower average laser power. The peaks observed at the intersections and the overlapping lines in scaffold #3 and scaffold #4 are indicative of lensing effects that occur whenever the laser passes over a previously polymerized spot. It is anticipated that these artifacts may be minimized by creating structures using multiple layers. The persistence of the structures after development in 1-propanol and soaking in ethanol demonstrates the stability of the zirconium oxide hybrid material after photopolymerization and postprocessing. The shrinkage properties of 2PP-based structures formed from this zirconium hybrid resin has been previously demonstrated by Ovsianikov et al.33 By optimizing the laser parameters, zirconium oxide hybrid material structures may be fabricated by 2PP without significant shrinkage, facilitating good fidelity between the 2PP-fabricated structure features and the computer design on which they were based. The elemental analysis of the zirconium oxide hybrid material scaffolds by EDS (Fig. 4) showed the representative peaks for the inorganic and organic constituents, including carbon, oxygen, silicon, and zirconium. The characteristic Zr La peak is visible at 2.042 keV on the EDS spectrum [Fig. 4(a)]. Prior to cell culture, samples were postprocessed using an alcohol soaking procedure for 1 week to remove excess monomers and photoinitiator; these components of 2PPfabricated samples may reduce cell viability.33 The lack of a nitrogen peak in the EDS spectrum suggests the removal of photoinitiator 4,40 -Bis(diethylamino) benzophenone. The EDS scan [Fig. 4(b)] mapping of Zr, O, and C on scaffold #2 [Fig. 4(c)] illustrates the presence of elements associated with the zirconium oxide hybrid material in the structure patterned by 2PP.
FIG. 4. (Color online) EDS spectrum (a), line scan (b), and mapping of a scaffold fabricated out of the inorganic–organic zirconium oxide hybrid material by 2PP.
029014-6 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
029014-6
B. Cellular response to 2PP-fabricated zirconium oxide hybrid material microstructured scaffolds
Since zirconium oxide is commonly utilized in orthopedic implants and dental implants, cellular response to the scaffolds was evaluated using hBMSCs. These nonhematopoietic progenitor cells can differentiate into musculoskeletal tissues that come in contact with a variety of implantable medical devices. Long-term tissue culture studies were conducted to evaluate the cellular attachment, orientation, and growth on the 3-D microstructures and to determine the potential of these scaffolds for long-term 3-D cell culture of stem cells for tissue engineering applications. For evaluation of cell interactions with the 2PP fabricated zirconium-based scaffolds, hBMSCs were seeded on the scaffolds at a density of 20 000 cells/scaffold and cultured for 45 days in the presence of osteogenic supplements. The scaffolds maintained their structure with minimal distortion or degradation, demonstrating their stability in a physiologically relevant environment. The cells adhered to all of the scaffold types and remained viable for up to 45 days, forming a dense, tissuelike matrix that bridged across the ridges of the scaffolds. SEM images of this tissue matrix on the scaffolds show that cell attachment conforms to the topographical features of the scaffolds (Fig. 5). Comparison of
FIG. 6. SEM images of hBMSCs on scaffold #2, demonstrating cellular attachment (a) and ingrowth within the scaffold microarchitecture (b).
the scaffolds with cells and without cells illustrates that tissue matrix was formed by cells; the peaklike nodules at the ridge joints and the ridge walls served as the initial cell attachment sites. Evenly distributed cell seeding and/or cell growth is evident in the formation of the tissuelike matrix throughout the entire surface of the scaffolds. High magnification SEM images of the tissue matrix reveal the spatial alignment of cells on the scaffolds; the relationship between cell alignment and ridge orientation varied for each scaffold design. For scaffolds #1, scaffold #2, and scaffold #3, the cells align diagonally to the uppermost scaffold ridges. For scaffold #4, the cells align parallel to the top ridge layer. Cells on the 2-D drop-cast zirconium oxide hybrid material control films and cells on the glass surfaces formed a dense cell layer, though no specific orientation was observed. These results demonstrate cell alignment that is dependent on the 3-D scaffold geometry. Cells were less densely grown in some regions toward the edge of scaffold #2 (Fig. 6). Individual cells can be seen spanning across and adhering to the walls of the grid. Cellular projections of the hBMSCs are visible, extending across the gaps between the scaffold ridges. Figure 6(b) shows a high magnification image of scaffold #2. In this image, cells have infiltrated the scaffold, stretching and forming adhesions on the scaffold in a three-dimensional fashion. Significant cell attachment and proliferation is evidenced by infiltration of hBMSCs into the interior of the scaffold and formation of a tissuelike matrix on the 2PPfabricated 3-D scaffolds.
IV. SUMMARY AND CONCLUSIONS
FIG. 5. SEM images (500 magnification) of hBMSCs grown on glass control substrate (a), drop-cast film of zirconium oxide hybrid material (b), scaffold #1 (c), scaffold #2 (d), scaffold #3 (e), and scaffold #4 (f) after 45 day cell culture. Biointerphases, Vol. 9, No. 2, June 2014
Four types of scaffold microarchitectures were fabricated out of a zirconium oxide hybrid material using two-photon polymerization. Scanning electron microscopy and atomic force microscopy demonstrated that the scaffolds exhibit uniform dimensions that correspond closely with the computer designs used in scaffolding processing. Energy dispersive x-ray spectroscopy was used to confirm presence of elements that are consistent with the organic and inorganic moieties in the fabricated structure and the absence of residual photoinitiator; no unanticipated impurities were noted in the 2PP-fabricated structures. Multiple identical samples of
029014-7 Skoog et al.: Two-photon polymerization of 3-D zirconium oxide hybrid scaffolds
each scaffold design were fabricated, demonstrating the reproducibility of the two-photon polymerization process. Long-term cell culture studies using hBMSCs showed cellular adhesion and cellular alignment on all scaffold microarchitectures; a dense, tissuelike structure was noted to span the gaps of the porous scaffolds. The cell orientation was shown to be dependent on the microarchitectures of the scaffolds. This study demonstrates the potential of zirconium oxide hybrid material-based 3-D microarchitectures for long-term growth of hBMSCs. These microstructured materials may be used as a model for development of a 3-D tissue culture system and as a valuable tool for bone tissue engineering. ACKNOWLEDGMENTS The authors would like to acknowledge the FDA White Oak Nanotechnology Core Facility for instrument use and scientific and technical assistance. In addition, the authors would like to acknowledge financial support from U.S. National Science Foundation Award #1041375. The authors would also like to thank the Laser Zentrum Hannover (LZH) in Hannover, Germany for use of their two-photon polymerization system for fabrication procedures. A.K. and B.N.C. acknowledge support by the German Research Foundation Cluster of Excellence Rebirth “From Regenerative Biology to Reconstructive Therapy” and the Lower Saxony project “Biofabrication for NIFE.” The mention of commercial products, their sources, or their use in connection with material reported herein is not to be construed as either an actual or implied endorsement of such products by the Department of Health and Human Services. 1
V. Aprile, S. M. Eaton, M. Lagana, G. Cerullo, M. T. Raimondi, and R. Osellame, Proc. SPIE 8247, 824708 (2012). 2 O. Z. Fisher, A. Khademhosseini, R. Langer, and N. A. Peppas, Acc. Chem. Res. 43, 419 (2010). 3 A. M. Greiner, B. Richter, and M. Bastmeyer, Macromol. Biosci. 12, 1301 (2012). 4 H. Jeon, H. Hidai, D. J. Hwang, and C. P. Grigoropoulos, J. Biomed. Mater. Res., Part A 93A, 56 (2009). 5 P. J. Rodrigo, L. Kelemen, D. Palima, C. A. Alonzo, P. Ormos, and J. Gluckstad, Opt. Express 17, 6578 (2009). 6 H. Jeon, H. Hidai, D. J. Hwang, K. E. Healy, and C. P. Grigoropoulos, Biomaterials 31, 4286 (2010). 7 M. Licht, A. Uchugonova, K. Konig, and M. Straub, J. Opt. 14, 065601 (2012). 8 E. Cukierman, R. Pankov, and K. M. Yamada, Curr. Opin. Cell Biol. 14, 633 (2002). 9 D. J. Maltman and S. A. Przyborski, Biochem. Soc. Trans. 38, 1072 (2010). 10 E. Kaivosoja, P. Suvanto, G. Barreto, S. Aura, A. Soininen, S. Franssila, and Y. T. Konttinen, J. Biomed. Mater. Res., Part A 101A, 842 (2013).
Biointerphases, Vol. 9, No. 2, June 2014
11
029014-7
W. A. Loesberg, J. te Riet, F. C. M. J. M. van Delft, P. Schon, C. G. Figdor, S. Speller, J. J. W. A. van Loon, X. F. Walboomers, and J. A. Jansen, Biomaterials 28, 3944 (2007). 12 A. L. Teixeria, G. A. McKie, J. D. Foley, P. J. Bertics, P. F. Nealey, and C. J. Murphy, Biomaterials 27, 3945 (2006). 13 B. J. Papenburg, L. Vogelaar, L. A. M. Bolhuis-Versteeg, R. G. H. Lammertink, D. Stamatialis, and M. Wessling, Biomaterials 28, 1998 (2007). 14 J. P. Kaiser, A. Reinmann, and A. Bruinink, Biomaterials 27, 5230 (2006). 15 P. Tayalia, E. Mazur, and D. J. Mooney, Biomaterials 32, 2634 (2011). 16 Y. Lu, G. Mapili, G. Suhali, S. Chen, and K. Roy, J. Biomed. Mater. Res. A 77A, 396 (2006). 17 O. Zinger, G. Zhao, Z. Schwartz, J. Simpson, M. Wieland, D. Landolt, and B. Boyan, Biomaterials 26, 1837 (2005). 18 S. A. Ruiz and C. S. Chen, Stem Cells 26, 2921 (2008). 19 G. Kumar, C. K. Tison, K. Chatterjee, P. S. Pine, J. H. McDaniel, M. L. Salit, M. F. Young, and C. G. Simon, Jr., Biomaterials 32, 9188 (2011). 20 Y. Seol, D. Y. Park, J. Y. Park, S. W. Kim, S. J. Park, and D. Cho, Biotechnol. Bioeng. 110, 1444 (2013). 21 C. H. Thomas, J. H. Collier, C. S. Sfeir, and K. E. Healy, Proc. Natl. Acad. Sci. U.S.A. 99, 1972 (2002). 22 H. Hosseinkhani, M. Hosseinkhani, and H. Kobayashi, J. Bioact. Compat. Polym. 21, 277 (2006). 23 C. S. Ki, S. Y. Park, H. J. Kim, H. M. Jung, K. M. Woo, J. W. Lee, and Y. H. Park, Biotechnol. Lett. 30, 405 (2008). 24 R. J. Narayan, A. Doraiswamy, D. B. Chrisey, and B. N. Chichkov, Mater. Today 13, 42 (2010). 25 R. Schade, T. Weiss, A. Berg, M. Schnabelrauch, and K. Liefeith, Int. J. Artif. Organs 33, 219 (2010). 26 T. Weiss, G. Hildebrand, R. Schade, and K. Liefeith, Eng. Life Sci. 9, 384 (2009). 27 S. Englehardt, E. Hoch, K. Borchers, W. Meyer, H. Kruger, G. E. M. Tovar, and A. Gillner, Biofabrication 3, 025003 (2011). 28 S. Psycharakis, A. Tosca, V. Melissinaki, A. Giakoumaki, and A. Ranella, Biomed. Mater. 6, 045008 (2011). 29 T. M. Hsieh, C. W. B. Ng, K. Naryanan, A. C. A. Wan, and J. Y. Ying, Biomaterials 31, 7648 (2010). 30 A. Ovsianikov et al., Laser Chem. 2008, 1. 31 A. Ovsianikov et al., ACS Nano 2, 2257 (2008). 32 A. Ovsianikov, X. Shizhou, M. Farsari, M. Vamvakaki, C. Fotakis, and B. N. Chichkov, Opt. Express 17, 4 (2009). 33 A. Ovsianikov et al., Acta Biomater. 7, 967 (2011). 34 M. Ramalingam, E. Jabbarl, S. Ramakrishna, and A. Khademhosseini, Micro and Nanotechnologies in Engineering Stem Cells and Tissues (IEEE, Piscataway, NJ, 2013), pp. 85–86. 35 C. Piconi and G. Maccauro, Biomaterials 20, 1 (1999). 36 G. Maccauro, P. R. Iommetti, L. Raffaelli, and P. F. Manicone, “Alumina and zirconia ceramics for orthopaedic and dental devices,” in Biomaterials Applications for Nanomedicine, edited by R. Pignatello (InTech, Rijeka, Croatia, 2011), Chap. 5, pp. 300–307. 37 C. A. M. Volpato, L. G. D. Garbelotto, M. C. Fredel, and F. Bandioli, “Application of zirconia in dentistry: Biological, mechanical, and optical considerations,” in Advances in Ceramics—Electric and Magnetic Ceramics, Bioceramics, Ceramics and Environment, edited by C. Sikalidis (InTech, Rijeka, Croatia, 2011), Chap. 17, pp. 397–404. 38 S. Bayou, M. Mouzali, F. Aloui, L. Lecamp, and P. Lebaudy, Polym. J. 45, 863 (2013). 39 K. S. Chan, D. P. Nicolella, B. R. Furman, S. T. Wellinghoff, H. R. Rawls, and S. E. Pratsinis, J. Mater. Sci. 44, 22 (2009).
