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3D printing of novel osteochondral scaffolds with graded microstructure
This content has been downloaded from IOPscience. Please scroll down to see the full text. 2016 Nanotechnology 27 414001 (http://iopscience.iop.org/0957-4484/27/41/414001) View the table of contents for this issue, or go to the journal homepage for more
Download details: IP Address: 207.162.240.147 This content was downloaded on 09/09/2016 at 07:26
Please note that terms and conditions apply.
You may also be interested in: Directing chondrogenic differentiation of mesenchymal stem cells with a solid-supported chitosan thermogel for cartilage tissue engineering Hongjie Huang, Xin Zhang, Xiaoqing Hu et al. A synergistic approach to the design, fabrication and evaluation of 3D printed micro and nano featured scaffolds for vascularized bone tissue repair Benjamin Holmes, Kartik Bulusu, Michael Plesniak et al. Surface energy and stiffness discrete gradients in additive manufactured scaffolds for osteochondral regeneration Andrea Di Luca, Alessia Longoni, Giuseppe Criscenti et al. Triphasic scaffolds for the regeneration of the bone–ligament interface G Criscenti, A Longoni, A Di Luca et al. Additive manufacturing of collagen scaffolds by three-dimensional plotting of highly viscous dispersions Anja Lode, Michael Meyer, Sophie Brüggemeier et al. Recent progress in stem cell differentiation directed by material and mechanical cues Xunxun Lin, Yuan Shi, Yilin Cao et al.
Nanotechnology Nanotechnology 27 (2016) 414001 (10pp)
doi:10.1088/0957-4484/27/41/414001
3D printing of novel osteochondral scaffolds with graded microstructure Margaret A Nowicki1, Nathan J Castro1, Michael W Plesniak1 and Lijie Grace Zhang1,2,3 1
Department of Mechanical and Aerospace Engineering, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA 2 Department of Biomedical Engineering, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA 3 Department of Medicine, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA E-mail: [email protected] Received 20 January 2016, revised 29 July 2016 Accepted for publication 9 August 2016 Published 8 September 2016 Abstract
Osteochondral tissue has a complex graded structure where biological, physiological, and mechanical properties vary significantly over the full thickness spanning from the subchondral bone region beneath the joint surface to the hyaline cartilage region at the joint surface. This presents a significant challenge for tissue-engineered structures addressing osteochondral defects. Fused deposition modeling (FDM) 3D bioprinters present a unique solution to this problem. The objective of this study is to use FDM-based 3D bioprinting and nanocrystalline hydroxyapatite for improved bone marrow human mesenchymal stem cell (hMSC) adhesion, growth, and osteochondral differentiation. FDM printing parameters can be tuned through computer aided design and computer numerical control software to manipulate scaffold geometries in ways that are beneficial to mechanical performance without hindering cellular behavior. Additionally, the ability to fine-tune 3D printed scaffolds increases further through our investment casting procedure which facilitates the inclusion of nanoparticles with biochemical factors to further elicit desired hMSC differentiation. For this study, FDM was used to print investment-casting molds innovatively designed with varied pore distribution over the full thickness of the scaffold. The mechanical and biological impacts of the varied pore distributions were compared and evaluated to determine the benefits of this physical manipulation. The results indicate that both mechanical properties and cell performance improve in the graded pore structures when compared to homogeneously distributed porous and non-porous structures. Differentiation results indicated successful osteogenic and chondrogenic manipulation in engineered scaffolds. Keywords: 3D bioprinting, osteochondral, gradient, nanomaterials, tissue engineering (Some figures may appear in colour only in the online journal) Introduction
weight may lead to increased joint loading, thus inducing or further exacerbating acute trauma and progressive degeneration. Furthermore, 33% of this population are workforce contributors between 45 and 64 years old [2]. This disease undoubtedly impacts patient quality of life, but the age range of a large portion of its target population also indirectly impacts economic productivity through reduced contribution to the work force and increased disability compensation.
In 2005, 47.8 million Americans were diagnosed with osteoarthritis, a degenerative joint disease marked by symptomatic joint pain and dysfunction [1]. This population is currently predicted to grow to 67 million by 2030; the national obesity epidemic is not considered in this estimate and could further inflate the 2030 prediction. Increased 0957-4484/16/414001+10$33.00
1
© 2016 IOP Publishing Ltd Printed in the UK
Nanotechnology 27 (2016) 414001
M A Nowicki et al
investigated 3D printing biomimetic nanocomposite scaffolds for improved osteochondral tissue regeneration using stereolithography. It was found that human bone marrow-derived mesenchymal stem cell adhesion, proliferation, and osteochondral differentiation were greatly improved in the biomimetic, graded, 3D printed, osteochondral construct in vitro. This work also illustrated the benefit of nano-ink integration in 3D printing technology for efficient and effective fabrication of novel nanocomposite hydrogel scaffolds [10]. The current study will extend these techniques while transitioning to the fused deposition modeling (FDM) 3D printing platform facilitating fabrication of more complex and robust designs. Mechanical performance is another key feature of osteochondral scaffolds due to their load-bearing nature. Integrating pores increases surface area available for cell attachment. It also allows cells and nutrients to infiltrate deep within the scaffolds maximizing the potential for uniform tissue development as the scaffold degrades [13]. Pores also act as stress concentrations, especially in load-bearing cases, that potentially compromise structural integrity and decrease peak material performance. Tang et al investigated stress concentration interactions in porous materials in 2005. The team found that, when loaded in compression, cracks nucleate from circular stress concentrations and grow parallel to the direction of axial compression [12]. It is postulated that disrupting pore continuity in the direction parallel to axial compression improves mechanical performance in load bearing scaffolds. The objective of this study is to use FDM-based 3D bioprinting and nanocrystalline hydroxyapatite (nHA) for improved bone marrow human mesenchymal stem cell (hMSC) adhesion, growth, and osteochondral differentiation using novel investment-casting molds innovatively designed with varied pore distribution over the full thickness of the scaffold. Four different pore distributions were investigated in this study to compare the effects of homogeneous and inhomogeneous distributions. It was hypothesized that an increased mechanical performance could be obtained by varying the pore distribution allowing greater load bearing capacity, favorable for osteogenic differentiation. nHA was incorporated within the graded scaffolds to further assist with osteogenic differentiation. Thus, this single-phase scaffold can promote biphasic differentiation favorable to the osteochondral region.
Joint degeneration is caused by the gradual loss of cartilaginous tissue at the joint surface. Though existing minimally invasive methods of treatment or mitigation of disease progression do exist, most severe cases eventually lead to total joint arthroplasty. Tissue engineering (TE) applies the principles of engineering and life sciences for the development of biological substitutes that restore, maintain, or improve tissue function [3]. The developing field of TE holds great promise for realizing effective methods of repairing damaged tissue, thus eliminating or prolonging the need for total joint replacement. TE for joint repair involves harvesting cells from a patient, seeding and growing them upon a threedimensional (3D) scaffold in vitro, and transplanting them back into the patient after desired cellular differentiation is achieved. The major challenge presented by TE scaffolds at joint surfaces, and especially at the osteochondral interface, is the diverse nature of tissue-specific extracellular matrix (ECM) composition and mechanical properties. ECM composition found in the osteochondral region is extremely diverse and challenging to reproduce as a single unit. While bone TE advances rapidly, with much success due in part to the great regenerative capacities of this type of tissue, cartilage has proven more challenging. Cartilage remains illusive in large part because of its avascular nature which limits its ability to circulate nutrients within the tissue [4]. Additionally, this challenge is further complicated at joint surfaces where cartilage also contains a small number of condrocytes with limited mobility within the dense ECM and high amounts of protease inhibitors, possibly responsible for the low regenerative capacity [5]. Since damaged cartilage is not naturally replaced in an effective manner with functional tissue, severe damage often requires surgical intervention [4]. Addressing this challenge alone is difficult, but coupling this with the disparate nature of the tissue spanning this entire region, from bone to cartilage, further compounds the problem. There are generally two approaches in repairing tissue in this region: scaffold-based and scaffold-free. Scaffoldbased approaches combine cells on biodegradable, biocompatible support structures with growth factors for the promotion of desired cellular responses [6]. Two types of scaffolds exist within the scaffold-based concept: single phase and bi/multi-phase. Single-phase scaffolds are largely composed of one homogeneous material often supplemented with growth factors. Bi- or multiphasic scaffolds are often composed of more than one material and again frequently supplemented with one or more types of growth factors distributed within to promote cellular response [6, 7]. ‘Scaffoldless TE refers to any platform that does not require cell seeding or adherence within an exogenous, 3D material’ [9]. In scaffold-free processes, the material properties of the cells control ECM production. Additionally, large amounts of cells are required for adequate ECM formation [8]. For this reason, cell source identification is critical and often requires the use of fully differentiated, autologous cells to provide adequate guidance and support while minimizing the risk of rejection [11]. Most scaffold-based TE approaches are constructed using various 3D bioprinting techniques. Our lab recently
Materials and methods Scaffold development Nanohydroxyapatite synthesis. nHA was synthesized using
a wet chemistry process, followed by hydrothermal treatment. Briefly, 75 ml of 0.6 M ammonium phosphate (Sigma Aldrich, St. Louis, MO) solution was added to 750 ml of water; the pH was then adjusted to 10 using ammonium hydroxide (Fisher Scientific, Pittsburg, PA). After achieving adequate pH, 1 M calcium nitrate (Signma Aldrich, St. Louis, MO) was added at a rate of 5 ml min−1 while stirring. The 2
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Figure 1. CAD drawings of scaffolds showing intended pore density and alignment. (A) Homogeneous high density, (B) homogeneous low
density, (C) biphasic, and (D) triphasic scaffolds [15].
firm. Irgacure 819 photoinitiator (Ciba Specialty Chemicals Inc., Tarrtown, NY) was used to assist in the photocrosslinking process. Heated sonication in D-limonene and ultrapure water assisted in dissolution of the HIPS from the crosslinked hydrogel leaving a highly interconnected porous structure. Specimens were collected with an 8 mm biopsy punch during the sonication process to expedite HIPS leaching. D-limonene is listed in the Code of Federal Regulations as generaly recognized as safe for human consumption with fairly low toxicity [16]. D-limonene, though safe for humans, must be thoroughly removed to prevent second order effects to ensure cytocompatibility. Additionally, D-limonene is extremely oily, and thus hydrophobic, which can minimize cell attachment and spreading. Therefore it is necessary to thoroughly rinse samples after D-limonene exposure using several rinses followed by soaking in ethanol. Figure 2 illustrates the scaffold fabrication process. For in vitro cell studies, nHA was incorporated in the biphasic and triphasic scaffolds to validate the effects of osteoconductive nanomaterials on scaffold performance. An nHA equivalent of 60 wt% of PEG-DA was uniformly mixed into the hydrogel before casting into the HIPS molds. This nHA conentration was selected based on data from a previous study in our lab [17]. These samples were cross-linked under UV light, leached, and punched as previously described.
mixture was then left at room temperature for 10 min to allow nHA precipitation. The HA mixture was then placed in a Teflon liner within an acid digestion chamber (Parr Instrument Company, Moline, IL) and treated at 200 °C for 20 h. Once complete, the nHA solution was cetrifuged and the precipitate washed three times before being dried in an 80 °C oven for 12 h. Finally, the dried nHA was ground into a fine powder and characterized thoroughly [14].
Scaffold design and fabrication.
Square molds measuring 35 mm×35 mm×3.6 mm (length×width×height) for investment casting of poly(ethylene glycol) (PEG) hydrogels were drawn using computer aided design (CAD) and prepared for 3D printing by computer numerical control (CNC) software. Overall mold thickness of each sample remained constant for all samples groups consisting of homogeneous, biphasic, and triphasic pore density and distribution. Modification of the CNC file with regards to infill density allows for precise control of pore distribution and density. Homogeneous scaffolds were designed and fabricated with one continuous infill density over the entire 3.6 mm thickness; two pore densities were created, one with high density (40% infill) and one with low-density (20% infill) pore distribution. The density of the biphasic scaffold varied over the full thickness. The lower 1.8 mm was fabricated with high pore distribution, while the upper 1.8 mm contained a less dense pore distribution. The triphasic scaffold pore density also varied over entirety of the scaffold thickness in 1.2 mm increments transitioning from high to low density with an intermediate layer of 30% infill density. CAD models of each scaffold are illustrated in figure 1. FDM of high impact polystyrene (HIPS) was used to fabricate the scaffold molds. After printing, a PEG/PEG–diacrylate (DA) (40/60 wt%) hydrogel solution was cast to each mold and cross-linked under ultraviolet (UV) light until
Scaffold characterization Optical and scanning electron microscopy (SEM).
Optical microscopy was used to validate and examine complete leaching of mold material and pore distribution. Samples were left to air dry at room temperature for 2 h and viewed using a CM4210 Optical System with manual zoom (6.4×) and fine focus (656×492 pixel size) (DSA25, Kruss USA,
3
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Figure 2. Scaffold production process moving from FDM mold production, through the leaching process, to scaffold completion. Adapted and modified from Nowicki et al [15].
Grand Island, NY) under standard culture conditions (5% CO2/95% air controlled at 37 °C). All samples were sterilized under UV light for 30 min and soaked in 70% ethanol for 15 min. After sterilization, samples were washed in phosphate-buffered solution (PBS) 3 times and pre-wet in complete media over night before cell seeding.
Matthews, NC). SEM analysis was used to determine average pore size and spacing, as well as to examine nHA distribution. SEM samples were dried at room tmeperature overnight and viewed at various magnifications and 3 kV accelerating voltage (Zeiss NVision 40 FIB, Thornwood, NY). Mechanical testing. Uniaxial unconfined compression testing was conducted on each sample group using an MTS Criterion Model 43 mechanical tester with a 1 kN load cell at a crosshead speed of 2 mm min−1 (n=5, Applied Test Systems, Butler, PA). 8 mm samples were stored in ultrapure water and blotted dry prior to testing. The loads and displacements were continuously monitored during testing and employed to plot stress versus strain after complete. The slope of the elastic portion of the stress versus strain curve identififed the Young’s modulus and the highest recorded load identified the peak stress.
hMSC adhesion and proliferation. For cell adhesion and
proliferation studies, scaffolds were seeded at a density of 30 000 cells/scaffold. For cell adhesion, cell-seeded scaffolds were incubated for four hours and transferred to a new well plate for evaluation using CellTiter 96 AQueous One Solution Cell proliferation Assay (colorimetric MTS assay). For cell proliferation studies, samples were cultured for 1, 3, and 5 d in complete media, respectively. After predetermined time intervals, samples were transferred to new well plates and evaluated via colorimetric assay. Absorbance was measured at 490 nm wavelength. For qualitative cell proliferation confirmation, laser scanning confocal microscopy (LSCM 10, Zeiss) was used to visually assess cell performance on the triphasic PEG-DA hydrogel scaffold. After 1, 3, and 5 d time points cells were rinsed twice with PBS and fixed with 10% Formalin for 10 min at room temperature. After fixation, cells were permeabilized with 0.1% Triton X-100 solution in PBS for 5 min at room temperature and then rinsed twice with PBS. Double-staining with Texas Red (ThermoFisher Scientific) and 4′,6-Diamidine-2′-phenylindole dihydrochloride (DAPI)
In vitro hMSC evaluation Cell culture. Primary hMSCs were obtained from healthy consenting donors and thoroughly characterized at the Texas A&M Health Science Center, Institute for Regenerative Medicine. All cell studies were performed with cell of passages #3–6. Cells were cultured in complete media composed of Alpha minimum essential medium (α-MEM, Gibco, Grand Island, NY), 16% fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA), 1% L-Glutamine (Lonza, Walkersville, MD), and 1% penicillin:streptomycin (Gibco, 4
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Table 1. Scaffold dimensions measured using SEM images and CAD design parameters.
Measured Pore distribution
Pore density
Pore diameter (SEM)
Homogeneous Homogeneous Biphasic
High Low High
200–300 μm 200–300 μm 200–300 μm
Triphasic
High
200–300 μm
Designed
Avg. horizontal pore spacing (SEM)
Pore diameter (CAD)
Horizontal pore spacing (CAD)
750 μm 1450 μm Phase 1: 750 μm Phase 2: 1450 μm Phase 1: 750 μm Phase 2: 950 μm Phase 3: 1450 μm
250 μm 250 μm 250 μm
750 μm 1450 μm Phase 1: 750 μm Phase 2: 1450 μm Phase 1: 750 μm Phase 2: 950 μm Phase 3: 1450 μm
250 μm
24 h 20 μl of supernatant was mixed with 1 ml calcium colorant, o-cresolphthalein complexone. 100 μl of each mixture was then transferred to a 96 well-plate and read using an absorbance spectrophotometer at a wavelength of 570 nm. Human collagen, types I and II, was evaluated by double-antibody sandwich enzyme-linked immunosorbent assay (ELISA) (Fisher Scientific, Pittsburg, PA) per manufacturer’s instructions. Digested sample aliquots were added to 96-well plates pre-coated with purified human collagen type I or type II antibody and incubated for 40 min. Unbound sample was washed from each well and collagen type I or type II specific detection antibody was added and incubated for 20 min then removed and washed. After washing, HRPStreptavidin enzyme conjugate was added to each well, incubated for 10 min, removed and washed. Tetramethylbenzidine (TMB) substrate was subsequently added and samples were incubated for 15 min. Lastly, an acidic stop solution was directly added to the well plates and samples were read using an absorbance spectrophotometer at a wavelength of 450 nm. All differentiation assays were normalized to cell density per scaffold. Cell density was quantified by DNA PicoGreen assay and read by a fluorescent spectrophotometer with excitation and emission at 485 and 520 nm, respectively.
was used to stain the cytoskeleton and cell nucleus, respectively. Scaffolds were incubated at room temperature for 20 min and 5 min with two washing cycles using PBS. Scaffolds were then imaged at 20× magnification.
hMSC differentiation studies. For hMSC differentiation
studies, the following scaffolds were selected: solid bare PEG-DA, triphasic bare PEG-DA, triphasic PEG-DA with 60% nHA, and biphasic PEG-DA with 60% nHA. Scaffolds were seeded at 100 000 cells per scaffold and cultured in complete media supplemented with chondrogenic factors for 1, 2, and 3 weeks, respectively. nHA was added to induce osteogenesis within the scaffold and the chondrogenic media facilitated chondrogenesis in regions of the scaffold void of nHA. After each time point, scaffolds were transferred to a fresh well-plate and frozen at −80 °C for 24 h, lyophilized for 48 h, and digested in papain for 24 h at 60 °C. Total collagen was quantified via Picro Sirius Red staining. Analysis consisted of drying 100 μl of each sample in a well plate overnight. After drying, 150 μl of 0.1% Sirius Red in saturated picric acid was added to each well and incubated for 1 h at room temperature on an orbital shaker. After 1 h, each well was washed three times with 200 μl of 5% acetic acid. Lastly, 150 μl of 0.1 M sodium hydroxide was added to each well, incubated at room temperature for 30 min, and analyzed spectrophotometrically at a wavelength of 550 nm. Glycosaminoglycan (GAG), a key component of cartilage ECM, was quantified using a dimethylmethylene Blue (DMB) assay. After combining 3.04 g glycine, 2.37 g sodium chloride, and 95 ml of 0.1 M hydrochloric acid in 1 l of distilled water 16 mg of DMB was added and mixed well for 20 min. Samples were tested against a GAG standard (Biocolor Life Science Assays, UK) diluted in papain. For both standard and sample analysis, 20 μl of standard or sample was added to 250 μl DMB and read spectrophotometrically at a wavelength of 525 nm. Extracellular calcium deposition was evaluated as a latestage osteogenic marker for in vitro hMSC osteogenic differentiation. Calcium deposition was measured using a calcium reagent kit (Pointe Scientific Inc., Canton, MI). Briefly, lysed scaffolds, as well as uncultured scaffolds were immersed in a 0.6 N hydrochloric acid solution at 37 °C. After
Statistical analysis
All data are represented as the mean value±standard error of the mean and a student’s t-test was used to evaluate differences among the groups. Student’s t-test comparisons yielding p
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My IOPscience
3D printing of novel osteochondral scaffolds with graded microstructure
This content has been downloaded from IOPscience. Please scroll down to see the full text. 2016 Nanotechnology 27 414001 (http://iopscience.iop.org/0957-4484/27/41/414001) View the table of contents for this issue, or go to the journal homepage for more
Download details: IP Address: 207.162.240.147 This content was downloaded on 09/09/2016 at 07:26
Please note that terms and conditions apply.
You may also be interested in: Directing chondrogenic differentiation of mesenchymal stem cells with a solid-supported chitosan thermogel for cartilage tissue engineering Hongjie Huang, Xin Zhang, Xiaoqing Hu et al. A synergistic approach to the design, fabrication and evaluation of 3D printed micro and nano featured scaffolds for vascularized bone tissue repair Benjamin Holmes, Kartik Bulusu, Michael Plesniak et al. Surface energy and stiffness discrete gradients in additive manufactured scaffolds for osteochondral regeneration Andrea Di Luca, Alessia Longoni, Giuseppe Criscenti et al. Triphasic scaffolds for the regeneration of the bone–ligament interface G Criscenti, A Longoni, A Di Luca et al. Additive manufacturing of collagen scaffolds by three-dimensional plotting of highly viscous dispersions Anja Lode, Michael Meyer, Sophie Brüggemeier et al. Recent progress in stem cell differentiation directed by material and mechanical cues Xunxun Lin, Yuan Shi, Yilin Cao et al.
Nanotechnology Nanotechnology 27 (2016) 414001 (10pp)
doi:10.1088/0957-4484/27/41/414001
3D printing of novel osteochondral scaffolds with graded microstructure Margaret A Nowicki1, Nathan J Castro1, Michael W Plesniak1 and Lijie Grace Zhang1,2,3 1
Department of Mechanical and Aerospace Engineering, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA 2 Department of Biomedical Engineering, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA 3 Department of Medicine, The George Washington University, 800 22nd Street, NW, Washington, DC, 20052, USA E-mail: [email protected] Received 20 January 2016, revised 29 July 2016 Accepted for publication 9 August 2016 Published 8 September 2016 Abstract
Osteochondral tissue has a complex graded structure where biological, physiological, and mechanical properties vary significantly over the full thickness spanning from the subchondral bone region beneath the joint surface to the hyaline cartilage region at the joint surface. This presents a significant challenge for tissue-engineered structures addressing osteochondral defects. Fused deposition modeling (FDM) 3D bioprinters present a unique solution to this problem. The objective of this study is to use FDM-based 3D bioprinting and nanocrystalline hydroxyapatite for improved bone marrow human mesenchymal stem cell (hMSC) adhesion, growth, and osteochondral differentiation. FDM printing parameters can be tuned through computer aided design and computer numerical control software to manipulate scaffold geometries in ways that are beneficial to mechanical performance without hindering cellular behavior. Additionally, the ability to fine-tune 3D printed scaffolds increases further through our investment casting procedure which facilitates the inclusion of nanoparticles with biochemical factors to further elicit desired hMSC differentiation. For this study, FDM was used to print investment-casting molds innovatively designed with varied pore distribution over the full thickness of the scaffold. The mechanical and biological impacts of the varied pore distributions were compared and evaluated to determine the benefits of this physical manipulation. The results indicate that both mechanical properties and cell performance improve in the graded pore structures when compared to homogeneously distributed porous and non-porous structures. Differentiation results indicated successful osteogenic and chondrogenic manipulation in engineered scaffolds. Keywords: 3D bioprinting, osteochondral, gradient, nanomaterials, tissue engineering (Some figures may appear in colour only in the online journal) Introduction
weight may lead to increased joint loading, thus inducing or further exacerbating acute trauma and progressive degeneration. Furthermore, 33% of this population are workforce contributors between 45 and 64 years old [2]. This disease undoubtedly impacts patient quality of life, but the age range of a large portion of its target population also indirectly impacts economic productivity through reduced contribution to the work force and increased disability compensation.
In 2005, 47.8 million Americans were diagnosed with osteoarthritis, a degenerative joint disease marked by symptomatic joint pain and dysfunction [1]. This population is currently predicted to grow to 67 million by 2030; the national obesity epidemic is not considered in this estimate and could further inflate the 2030 prediction. Increased 0957-4484/16/414001+10$33.00
1
© 2016 IOP Publishing Ltd Printed in the UK
Nanotechnology 27 (2016) 414001
M A Nowicki et al
investigated 3D printing biomimetic nanocomposite scaffolds for improved osteochondral tissue regeneration using stereolithography. It was found that human bone marrow-derived mesenchymal stem cell adhesion, proliferation, and osteochondral differentiation were greatly improved in the biomimetic, graded, 3D printed, osteochondral construct in vitro. This work also illustrated the benefit of nano-ink integration in 3D printing technology for efficient and effective fabrication of novel nanocomposite hydrogel scaffolds [10]. The current study will extend these techniques while transitioning to the fused deposition modeling (FDM) 3D printing platform facilitating fabrication of more complex and robust designs. Mechanical performance is another key feature of osteochondral scaffolds due to their load-bearing nature. Integrating pores increases surface area available for cell attachment. It also allows cells and nutrients to infiltrate deep within the scaffolds maximizing the potential for uniform tissue development as the scaffold degrades [13]. Pores also act as stress concentrations, especially in load-bearing cases, that potentially compromise structural integrity and decrease peak material performance. Tang et al investigated stress concentration interactions in porous materials in 2005. The team found that, when loaded in compression, cracks nucleate from circular stress concentrations and grow parallel to the direction of axial compression [12]. It is postulated that disrupting pore continuity in the direction parallel to axial compression improves mechanical performance in load bearing scaffolds. The objective of this study is to use FDM-based 3D bioprinting and nanocrystalline hydroxyapatite (nHA) for improved bone marrow human mesenchymal stem cell (hMSC) adhesion, growth, and osteochondral differentiation using novel investment-casting molds innovatively designed with varied pore distribution over the full thickness of the scaffold. Four different pore distributions were investigated in this study to compare the effects of homogeneous and inhomogeneous distributions. It was hypothesized that an increased mechanical performance could be obtained by varying the pore distribution allowing greater load bearing capacity, favorable for osteogenic differentiation. nHA was incorporated within the graded scaffolds to further assist with osteogenic differentiation. Thus, this single-phase scaffold can promote biphasic differentiation favorable to the osteochondral region.
Joint degeneration is caused by the gradual loss of cartilaginous tissue at the joint surface. Though existing minimally invasive methods of treatment or mitigation of disease progression do exist, most severe cases eventually lead to total joint arthroplasty. Tissue engineering (TE) applies the principles of engineering and life sciences for the development of biological substitutes that restore, maintain, or improve tissue function [3]. The developing field of TE holds great promise for realizing effective methods of repairing damaged tissue, thus eliminating or prolonging the need for total joint replacement. TE for joint repair involves harvesting cells from a patient, seeding and growing them upon a threedimensional (3D) scaffold in vitro, and transplanting them back into the patient after desired cellular differentiation is achieved. The major challenge presented by TE scaffolds at joint surfaces, and especially at the osteochondral interface, is the diverse nature of tissue-specific extracellular matrix (ECM) composition and mechanical properties. ECM composition found in the osteochondral region is extremely diverse and challenging to reproduce as a single unit. While bone TE advances rapidly, with much success due in part to the great regenerative capacities of this type of tissue, cartilage has proven more challenging. Cartilage remains illusive in large part because of its avascular nature which limits its ability to circulate nutrients within the tissue [4]. Additionally, this challenge is further complicated at joint surfaces where cartilage also contains a small number of condrocytes with limited mobility within the dense ECM and high amounts of protease inhibitors, possibly responsible for the low regenerative capacity [5]. Since damaged cartilage is not naturally replaced in an effective manner with functional tissue, severe damage often requires surgical intervention [4]. Addressing this challenge alone is difficult, but coupling this with the disparate nature of the tissue spanning this entire region, from bone to cartilage, further compounds the problem. There are generally two approaches in repairing tissue in this region: scaffold-based and scaffold-free. Scaffoldbased approaches combine cells on biodegradable, biocompatible support structures with growth factors for the promotion of desired cellular responses [6]. Two types of scaffolds exist within the scaffold-based concept: single phase and bi/multi-phase. Single-phase scaffolds are largely composed of one homogeneous material often supplemented with growth factors. Bi- or multiphasic scaffolds are often composed of more than one material and again frequently supplemented with one or more types of growth factors distributed within to promote cellular response [6, 7]. ‘Scaffoldless TE refers to any platform that does not require cell seeding or adherence within an exogenous, 3D material’ [9]. In scaffold-free processes, the material properties of the cells control ECM production. Additionally, large amounts of cells are required for adequate ECM formation [8]. For this reason, cell source identification is critical and often requires the use of fully differentiated, autologous cells to provide adequate guidance and support while minimizing the risk of rejection [11]. Most scaffold-based TE approaches are constructed using various 3D bioprinting techniques. Our lab recently
Materials and methods Scaffold development Nanohydroxyapatite synthesis. nHA was synthesized using
a wet chemistry process, followed by hydrothermal treatment. Briefly, 75 ml of 0.6 M ammonium phosphate (Sigma Aldrich, St. Louis, MO) solution was added to 750 ml of water; the pH was then adjusted to 10 using ammonium hydroxide (Fisher Scientific, Pittsburg, PA). After achieving adequate pH, 1 M calcium nitrate (Signma Aldrich, St. Louis, MO) was added at a rate of 5 ml min−1 while stirring. The 2
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Figure 1. CAD drawings of scaffolds showing intended pore density and alignment. (A) Homogeneous high density, (B) homogeneous low
density, (C) biphasic, and (D) triphasic scaffolds [15].
firm. Irgacure 819 photoinitiator (Ciba Specialty Chemicals Inc., Tarrtown, NY) was used to assist in the photocrosslinking process. Heated sonication in D-limonene and ultrapure water assisted in dissolution of the HIPS from the crosslinked hydrogel leaving a highly interconnected porous structure. Specimens were collected with an 8 mm biopsy punch during the sonication process to expedite HIPS leaching. D-limonene is listed in the Code of Federal Regulations as generaly recognized as safe for human consumption with fairly low toxicity [16]. D-limonene, though safe for humans, must be thoroughly removed to prevent second order effects to ensure cytocompatibility. Additionally, D-limonene is extremely oily, and thus hydrophobic, which can minimize cell attachment and spreading. Therefore it is necessary to thoroughly rinse samples after D-limonene exposure using several rinses followed by soaking in ethanol. Figure 2 illustrates the scaffold fabrication process. For in vitro cell studies, nHA was incorporated in the biphasic and triphasic scaffolds to validate the effects of osteoconductive nanomaterials on scaffold performance. An nHA equivalent of 60 wt% of PEG-DA was uniformly mixed into the hydrogel before casting into the HIPS molds. This nHA conentration was selected based on data from a previous study in our lab [17]. These samples were cross-linked under UV light, leached, and punched as previously described.
mixture was then left at room temperature for 10 min to allow nHA precipitation. The HA mixture was then placed in a Teflon liner within an acid digestion chamber (Parr Instrument Company, Moline, IL) and treated at 200 °C for 20 h. Once complete, the nHA solution was cetrifuged and the precipitate washed three times before being dried in an 80 °C oven for 12 h. Finally, the dried nHA was ground into a fine powder and characterized thoroughly [14].
Scaffold design and fabrication.
Square molds measuring 35 mm×35 mm×3.6 mm (length×width×height) for investment casting of poly(ethylene glycol) (PEG) hydrogels were drawn using computer aided design (CAD) and prepared for 3D printing by computer numerical control (CNC) software. Overall mold thickness of each sample remained constant for all samples groups consisting of homogeneous, biphasic, and triphasic pore density and distribution. Modification of the CNC file with regards to infill density allows for precise control of pore distribution and density. Homogeneous scaffolds were designed and fabricated with one continuous infill density over the entire 3.6 mm thickness; two pore densities were created, one with high density (40% infill) and one with low-density (20% infill) pore distribution. The density of the biphasic scaffold varied over the full thickness. The lower 1.8 mm was fabricated with high pore distribution, while the upper 1.8 mm contained a less dense pore distribution. The triphasic scaffold pore density also varied over entirety of the scaffold thickness in 1.2 mm increments transitioning from high to low density with an intermediate layer of 30% infill density. CAD models of each scaffold are illustrated in figure 1. FDM of high impact polystyrene (HIPS) was used to fabricate the scaffold molds. After printing, a PEG/PEG–diacrylate (DA) (40/60 wt%) hydrogel solution was cast to each mold and cross-linked under ultraviolet (UV) light until
Scaffold characterization Optical and scanning electron microscopy (SEM).
Optical microscopy was used to validate and examine complete leaching of mold material and pore distribution. Samples were left to air dry at room temperature for 2 h and viewed using a CM4210 Optical System with manual zoom (6.4×) and fine focus (656×492 pixel size) (DSA25, Kruss USA,
3
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Figure 2. Scaffold production process moving from FDM mold production, through the leaching process, to scaffold completion. Adapted and modified from Nowicki et al [15].
Grand Island, NY) under standard culture conditions (5% CO2/95% air controlled at 37 °C). All samples were sterilized under UV light for 30 min and soaked in 70% ethanol for 15 min. After sterilization, samples were washed in phosphate-buffered solution (PBS) 3 times and pre-wet in complete media over night before cell seeding.
Matthews, NC). SEM analysis was used to determine average pore size and spacing, as well as to examine nHA distribution. SEM samples were dried at room tmeperature overnight and viewed at various magnifications and 3 kV accelerating voltage (Zeiss NVision 40 FIB, Thornwood, NY). Mechanical testing. Uniaxial unconfined compression testing was conducted on each sample group using an MTS Criterion Model 43 mechanical tester with a 1 kN load cell at a crosshead speed of 2 mm min−1 (n=5, Applied Test Systems, Butler, PA). 8 mm samples were stored in ultrapure water and blotted dry prior to testing. The loads and displacements were continuously monitored during testing and employed to plot stress versus strain after complete. The slope of the elastic portion of the stress versus strain curve identififed the Young’s modulus and the highest recorded load identified the peak stress.
hMSC adhesion and proliferation. For cell adhesion and
proliferation studies, scaffolds were seeded at a density of 30 000 cells/scaffold. For cell adhesion, cell-seeded scaffolds were incubated for four hours and transferred to a new well plate for evaluation using CellTiter 96 AQueous One Solution Cell proliferation Assay (colorimetric MTS assay). For cell proliferation studies, samples were cultured for 1, 3, and 5 d in complete media, respectively. After predetermined time intervals, samples were transferred to new well plates and evaluated via colorimetric assay. Absorbance was measured at 490 nm wavelength. For qualitative cell proliferation confirmation, laser scanning confocal microscopy (LSCM 10, Zeiss) was used to visually assess cell performance on the triphasic PEG-DA hydrogel scaffold. After 1, 3, and 5 d time points cells were rinsed twice with PBS and fixed with 10% Formalin for 10 min at room temperature. After fixation, cells were permeabilized with 0.1% Triton X-100 solution in PBS for 5 min at room temperature and then rinsed twice with PBS. Double-staining with Texas Red (ThermoFisher Scientific) and 4′,6-Diamidine-2′-phenylindole dihydrochloride (DAPI)
In vitro hMSC evaluation Cell culture. Primary hMSCs were obtained from healthy consenting donors and thoroughly characterized at the Texas A&M Health Science Center, Institute for Regenerative Medicine. All cell studies were performed with cell of passages #3–6. Cells were cultured in complete media composed of Alpha minimum essential medium (α-MEM, Gibco, Grand Island, NY), 16% fetal bovine serum (Atlanta Biologicals, Flowery Branch, GA), 1% L-Glutamine (Lonza, Walkersville, MD), and 1% penicillin:streptomycin (Gibco, 4
Nanotechnology 27 (2016) 414001
M A Nowicki et al
Table 1. Scaffold dimensions measured using SEM images and CAD design parameters.
Measured Pore distribution
Pore density
Pore diameter (SEM)
Homogeneous Homogeneous Biphasic
High Low High
200–300 μm 200–300 μm 200–300 μm
Triphasic
High
200–300 μm
Designed
Avg. horizontal pore spacing (SEM)
Pore diameter (CAD)
Horizontal pore spacing (CAD)
750 μm 1450 μm Phase 1: 750 μm Phase 2: 1450 μm Phase 1: 750 μm Phase 2: 950 μm Phase 3: 1450 μm
250 μm 250 μm 250 μm
750 μm 1450 μm Phase 1: 750 μm Phase 2: 1450 μm Phase 1: 750 μm Phase 2: 950 μm Phase 3: 1450 μm
250 μm
24 h 20 μl of supernatant was mixed with 1 ml calcium colorant, o-cresolphthalein complexone. 100 μl of each mixture was then transferred to a 96 well-plate and read using an absorbance spectrophotometer at a wavelength of 570 nm. Human collagen, types I and II, was evaluated by double-antibody sandwich enzyme-linked immunosorbent assay (ELISA) (Fisher Scientific, Pittsburg, PA) per manufacturer’s instructions. Digested sample aliquots were added to 96-well plates pre-coated with purified human collagen type I or type II antibody and incubated for 40 min. Unbound sample was washed from each well and collagen type I or type II specific detection antibody was added and incubated for 20 min then removed and washed. After washing, HRPStreptavidin enzyme conjugate was added to each well, incubated for 10 min, removed and washed. Tetramethylbenzidine (TMB) substrate was subsequently added and samples were incubated for 15 min. Lastly, an acidic stop solution was directly added to the well plates and samples were read using an absorbance spectrophotometer at a wavelength of 450 nm. All differentiation assays were normalized to cell density per scaffold. Cell density was quantified by DNA PicoGreen assay and read by a fluorescent spectrophotometer with excitation and emission at 485 and 520 nm, respectively.
was used to stain the cytoskeleton and cell nucleus, respectively. Scaffolds were incubated at room temperature for 20 min and 5 min with two washing cycles using PBS. Scaffolds were then imaged at 20× magnification.
hMSC differentiation studies. For hMSC differentiation
studies, the following scaffolds were selected: solid bare PEG-DA, triphasic bare PEG-DA, triphasic PEG-DA with 60% nHA, and biphasic PEG-DA with 60% nHA. Scaffolds were seeded at 100 000 cells per scaffold and cultured in complete media supplemented with chondrogenic factors for 1, 2, and 3 weeks, respectively. nHA was added to induce osteogenesis within the scaffold and the chondrogenic media facilitated chondrogenesis in regions of the scaffold void of nHA. After each time point, scaffolds were transferred to a fresh well-plate and frozen at −80 °C for 24 h, lyophilized for 48 h, and digested in papain for 24 h at 60 °C. Total collagen was quantified via Picro Sirius Red staining. Analysis consisted of drying 100 μl of each sample in a well plate overnight. After drying, 150 μl of 0.1% Sirius Red in saturated picric acid was added to each well and incubated for 1 h at room temperature on an orbital shaker. After 1 h, each well was washed three times with 200 μl of 5% acetic acid. Lastly, 150 μl of 0.1 M sodium hydroxide was added to each well, incubated at room temperature for 30 min, and analyzed spectrophotometrically at a wavelength of 550 nm. Glycosaminoglycan (GAG), a key component of cartilage ECM, was quantified using a dimethylmethylene Blue (DMB) assay. After combining 3.04 g glycine, 2.37 g sodium chloride, and 95 ml of 0.1 M hydrochloric acid in 1 l of distilled water 16 mg of DMB was added and mixed well for 20 min. Samples were tested against a GAG standard (Biocolor Life Science Assays, UK) diluted in papain. For both standard and sample analysis, 20 μl of standard or sample was added to 250 μl DMB and read spectrophotometrically at a wavelength of 525 nm. Extracellular calcium deposition was evaluated as a latestage osteogenic marker for in vitro hMSC osteogenic differentiation. Calcium deposition was measured using a calcium reagent kit (Pointe Scientific Inc., Canton, MI). Briefly, lysed scaffolds, as well as uncultured scaffolds were immersed in a 0.6 N hydrochloric acid solution at 37 °C. After
Statistical analysis
All data are represented as the mean value±standard error of the mean and a student’s t-test was used to evaluate differences among the groups. Student’s t-test comparisons yielding p
