3D-Bioprinting of Polylactic Acid (PLA) NanofiberâAlginate Hydrogel

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3D-Bioprinting of Polylactic Acid (PLA) Nanofibers-Alginate Hydrogel Bioink Containing Human Adipose-Derived Stem Cells Lokesh Karthik Narayanan, Pedro Huebner, Matthew B. Fisher, Jeffrey T Spang, Binil Starly, and Rohan A Shirwaiker ACS Biomater. Sci. Eng., Just Accepted Manuscript • DOI: 10.1021/acsbiomaterials.6b00196 • Publication Date (Web): 06 Jul 2016 Downloaded from http://pubs.acs.org on July 9, 2016

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ACS Biomaterials Science & Engineering

3D-Bioprinting of Polylactic Acid (PLA) Nanofibers-Alginate Hydrogel Bioink Containing Human Adipose-Derived Stem Cells

Lokesh Karthik Narayanan1,2,3, Pedro Huebner1,2,3, Matthew B. Fisher3,4,5, Jeffrey T. Spang5, Binil Starly1,2,3,4, Rohan A. Shirwaiker1,2,3,4*

1

Edward P. Fitts Department of Industrial and Systems Engineering, North Carolina State University,

400 Daniels Hall, Raleigh, NC 27695, USA 2

Center for Additive Manufacturing and Logistics, North Carolina State University, Raleigh, NC 27695,

USA 3

4

Comparative Medicine Institute, North Carolina State University, Raleigh, NC 27695, USA Department of Biomedical Engineering, North Carolina State University, Engineering Building III,

Raleigh, NC 27695, USA, and University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA 5

Department of Orthopaedics, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599, USA

Abstract Bioinks play a central role in 3D-bioprinting by providing the supporting environment within which encapsulated cells can endure the stresses encountered during the digitally-driven fabrication process, and continue to mature, proliferate, and eventually form extracellular matrix (ECM). In order to be most effective, it is important that bioprinted constructs recapitulate the native tissue milieu as closely as possible. As such, musculoskeletal soft tissue constructs can benefit from bioinks that mimic their nanofibrous matrix constitution, which is also critical to their function. This study focuses on the development and proof-of-concept assessment of a fibrous bioink composed of alginate hydrogel, polylactic acid nanofibers and human adipose-derived stem cells (hASC) for bioprinting such tissue

* Corresponding Author: [email protected]

ACS Paragon Plus Environment


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constructs. First, hASC proliferation and viability were assessed in 3D-bioplotted strands over 16 days in vitro. Then, a human medial knee meniscus digitally modeled using magnetic resonance images was bioprinted and evaluated over 8 weeks in vitro. Results show that the nanofiber-reinforced bioink allowed higher levels of cell proliferation within bioprinted strands, with a peak at day 7, while still maintaining a vast majority of viable cells at day 16. The cell metabolic activity on day 7 was 28.5% higher in this bioink compared to the bioink without nanofibers. Histology of the bioprinted meniscus at both 4 and 8 weeks showed 54% and 147% higher cell density, respectively, in external versus internal regions of the construct. Presence of collagen and proteoglycans was also noted in areas surrounding the hASC, indicating ECM secretion and chondrogenic differentiation.

Keywords Bioprinting, Knee Meniscus, Alginate, Nanofibers, Adipose-Derived Stem Cells, Musculoskeletal Soft Tissue


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1. INTRODUCTION The recent emergence of bioprinting is playing an important role in the advancement of tissue engineering and regenerative medicine (TERM) technologies. Traditionally, TERM strategies have relied on engineered scaffolds to provide the necessary structural organization for cells that are seeded onto these fabricated structures to attach, proliferate and form extracellular matrix (ECM). In addition to providing the 3D architectural template, the scaffolds attempt to mimic the porous native environment of the cells. Owing to their ability to create well-defined strand-pore architectures, computer-aided additive manufacturing processes such as selective laser sintering1,2, stereolithography3,4, fused deposition modeling5,6, and 3D-Bioplotting7-10 have been widely used for scaffold fabrication for over a decade. These processes overcome limitations associated with traditional scaffold fabrication methods such as gas-foaming, solvent casting, and electrospinning by allowing the creation of reproducible single or multi-material highly interconnected porous 3D-architectures with patient-specific geometry11,12. More recent strategies focus on bioprinting, which involves the automated fabrication of biologically active constructs using an appropriate suspension of living cells or cell aggregates within “bioinks”13. The overall geometric organization and fabrication of these constructs is driven by computeraided digital files, which provide precise control over the geometric and process variables and allow high repeatability in fabrication. In addition to clinical applications in regenerative medicine, such bioprinted tissue constructs can serve as in vitro models for disease modeling, pharmacokinetic and cell biology studies. The different types of bioprinting processes, their engineering principles, characteristics and potential applications have been well described in literature13-18. In comparison to molding processes which are commonly used to fabricate soft hydrogel constructs, bioprinting processes offer a major advantage in that the constructs can be patterned in porous lattice architectures similar to scaffolds to promote nutrient perfusion. The ability to crosslink the bioinks layer by layer during the fabrication process as opposed to bulk cross-linking after molding is also beneficial. Bioinks play a critical role in bioprinting, primarily acting as the medium for cells to be delivered spatially into desired 3D patterns18-22. The primary components of a bioink are the cells, the biopolymer



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and any additives mixed in specific concentrations to achieve the desired structural stability and functional characteristics. Both stem cells and differentiated cell types can be included in bioinks, depending upon the application. The biopolymers which constitute a major portion of bioinks serve as a sustaining medium for these cells by providing rheological (viscosity, surface tension) and physical properties (stiffness, anchorage for cells) to enable their stable ejection/deposition during the fabrication of tissue constructs. Most importantly, they support cell homeostasis by providing a supportive hydrated environment that enables nutrient diffusion, and can present biochemical, cellular, and physical stimuli similar to direct cellular processes23. Biopolymers used in bioprinting include alginate, agarose and agar, collagen,

poly(lactic-co-glycolic

acid)

(PLGA)

and

their

derivatives,

poly(ethylene

glycol)

dimethacrylate, chitosan, and hyaluronic acid among others24. Additives are included for several reasons that range from modulating the properties of biopolymers and enhancing the bioink gelation process to eliciting specific cell responses and providing necessary supplementary functionality that cannot be provided by cells themselves. Commonly used additives include growth factors, decellularized ECM, RGD peptides, nanomaterials, drug loaded microspheres and hydrogel crosslinkers. Alginate is one of the most widely investigated biopolymer hydrogel used in bioinks for TERM applications. It is a naturally derived anionic polysaccharide that gelates on binding with divalent cations25, and has been used as a construct/encapsulation material for a variety of cell/tissue types such as bone/osteoblasts26,

muscle/smooth

muscle

cells27,

aortic

valve

leaflet

interstitial

cells27,

chondrocytes/cartilage28, and pancreatic islets29. Since alginate by itself has limited bioactivity due to the lack of bioligands necessary for mammalian cell adhesion30,31, recent investigations have looked at tailormade composites of alginate and other hydrogels to improve cell responses and achieve tissue-specific characteristics of interest. For example, Lin et al. noted improved cell adherence and proliferation while using alginate containing 25% hydroxyapatite for fabricating bone constructs26. Park et al. and Avila et al. demonstrated improvement in encapsulated cell proliferation and viability by using alginate containing bacterial nanocellulose fibrils32-34.


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It is known that the cells respond to the stiffness and topography of the surfaces they attach to, and their activities are affected when suspended in a 3D-environment35,36. Thus, the structural organization of bioinks and their ability to provide anchor surfaces for encapsulated cells is important in order to promote optimal tissue formation within the constructs. This becomes especially critical in applications involving musculoskeletal soft tissues such as tendons, ligaments, knee menisci, and the annulus fibrosus (AF) of the intervertebral disc, which have a more pronounced fibrous cytoskeletal organization. The ECM of these tissues is composed primarily of collagen fiber (80-98%) networks to satisfy the essential load bearing requirements of the tissues37-40. Recognizing its importance, recent investigations have looked at the effects of nanofibrous organization of scaffolds on musculoskeletal and stem cell responses and engineered tissue morphology41-45. For example, Nerurkar et al., Baker and Mauck, and Lee et al. showed that AF cells, meniscus cells, and human fibroblasts respond favorably to aligned nanofibrous networks in electrospun scaffolds46-48,. A majority of such studies, however, focus on scaffolds as opposed to bioprinted constructs, and advances in the development of fibrous bioinks suitable for musculoskeletal soft tissues have been fairly limited. In a step towards this direction, this study focuses on the early development and feasibility analyses of a nanofibrous bioink for bioprinting soft tissue constructs. The primary goal was to study the formulation and characteristics of the alginate-based bioink with suspended polylactic acid (PLA) nanofibers and stem cells derived from human adipose tissue (hASC). The nanofiber dispersion and compressive mechanical properties of the acellular hydrogels were determined before proceeding to study the hASC responses in bioprinted structures. An overview of our approach is presented in Figure 1. First, we investigate the in vitro viability and proliferation of hASC encapsulated in the bioink when bioprinted in the form of strands. Strands are uniform cylindrical structures deposited on a substrate and are basic building blocks of any extrusion 3D-bioprinted construct. The adipose-derived stem cells used in this study have become an important source of cells for musculoskeletal tissue engineering given their ease of isolation and their ability to differentiate into multiple cell lineages such as osteogenic, chondrogenic, myogenic, neurogenic and adipogenic lineages in a controlled manner49-52. In the second part of the study,



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we provide a proof-of-concept for bioprinting a human medial knee meniscus using the hASCencapsulated bioink. This construct was designed using magnetic resonance imaging (MRI) scans of a single patient’s knee, bioprinted layer-by-layer on a 3D-Bioplotter, and evaluated over 8 weeks in in vitro culture. The knee menisci (lateral and medial) are C-shaped cartilaginous structures that transmit loads between the tibia and femur53. We selected this fibrous musculoskeletal soft tissue for demonstration because meniscal tears are one of the most commonly reported orthopaedic injuries, and approximately 1 million surgeries involving the meniscus are performed in the US every year54-56. This tissue is mostly avascular in nature and possesses relatively poor healing capabilities. Current treatment strategies for young, active patients with significant meniscal deficiency involve the use of allografts, but given their known limitations including size matching57-59, patient-specific tissue engineered and bioprinted constructs can serve as a potential alternative.

Figure 1. An overview of this study investigating the bioprinting of a PLA nanofiber-alginate hydrogel


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bioink with encapsulated human adipose-derived stem cells (hASC).

2. MATERIALS AND METHODS 2.1. PLA Nanofiber Production The PLA nanofibers were produced using the patented XanoShear process (Xanofi, Raleigh, NC) and characterized as described previously60. Briefly, PLA beads (4060D, Natureworks, Minnetonka, MN) were dissolved in tetrahydrofuran (THF; Brenntag Southeast, Durham, NC) to form the polymer solution. Separately, a viscous dispersion medium was created by mixing glycerin (Brenntag Southeast) and deionized (DI) water (antisolvent for the THF and PLA). The nanofibers were produced in a continuous process by injecting the polymer solution into a conduit of viscous dispersion media under laminar Poiseuille shear flow. The polymer droplets injected into the flow were stretched as they precipitated into individual nanofibers, and the final product was collected as a slurry. Finally, the PLA nanofiber wet cakes were separated from the slurry by pumping through a filter press and repeatedly washed with DI water to remove any excess THF and glycerin. The fiber diameter and length were characterized prior to sterilization and dispersion in alginate gels46. Briefly, single drops of 75% glycerin mixed with nanofibers were placed on microscope slides (n = 3) and cover slips were placed on top. A total of 10 phase-contrast images/slide were captured at random locations on the slide using a 5 mega-pixels charge-coupled device (CCD) camera on an Olympus BH-2 microscope (Olympus, Tokyo, Japan). The lengths and diameters of at least 100 fibers/slide were measured from these images using ImageJ (NIH, Bethesda, MD). The data are reported as mean ± standard deviation. Scanning electron microscope (SEM) images of the nanofibers were also obtained. Briefly, a thin sheet of PLA nanofibers placed on carbon tape was sputter-coated (Hummer VI Sputter System, Anatech USA, Union City, CA) with silver (12 mA of current at 75 millitorr absolute pressure for 2 minutes), and imaged (2000x magnification) at an excitation voltage of 10 kV using a NeoScope JCM-5000 SEM (Jeol Ltd., Tokyo, Japan).



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2.2. Preparation of Alginate and Alginate-Nanofiber Hydrogels A batch of 3 g sodium alginate salt (WillPowder, Miami Beach, FL) was subjected to ethanol sterilization at the beginning of the study. 15 mL of 100% ethanol was added to 3 g of alginate salt in a petri dish, mixed thoroughly by pipetting, and left inside a biosafety cabinet for 24 hours to allow the ethanol to evaporate. To prepare the 2.5% w/w alginate hydrogel (henceforth referred to as acellular Alg), 29.25 mL of sterile DI water was added to 0.75 g of the sterilized alginate salt. The mixture was then stirred using a sterile spatula and sonicated in an ultrasonic water bath (Ultrasonik 28X Degasser, Ney Ultrasonics, Jamestown, NY) at 60 Hz for 2 hours. A small portion of the PLA nanofibers wet cake was ethanol sterilized using a similar approach; 1.5 g of nanofibers were mixed with 15 mL of 100% ethanol in a petri dish inside a biosafety cabinet, and the alcohol was allowed to evaporate over 24 hours. The nanofibers were then washed multiple times with sterile phosphate-buffered saline (PBS; pH 7.4, no MgCl2 and CaCl2, Thermo Fisher Scientific, Waltham, MA). To prepare the alginate-nanofiber composite hydrogel, 2 mL of sterile DI water was added to 0.789 g of the sterilized PLA wet cake (containing approximately 19% PLA nanofibers by composition), and mixed by pipetting repeatedly until there were no agglomerations greater than 0.5 mm in diameter. To this solution, 0.75 g of sterilized alginate salt and 26.461 mL of DI water were added and stirred with a sterile spatula until there were no visible alginate salt clumps. Finally, the mixture was agitated in a vortex mixer for 5 minutes and sonicated at 60 Hz for 2 hours in the ultrasonic water bath to obtain 30 mL of 2.5% w/w alginate – 0.5% w/w PLA nanofiber hydrogel (henceforth referred to as acellular Alg-Nf). Note that acellular hydrogel formulations with higher concentrations of alginate and nanofibers were also screened for extrusion printing capabilities at the beginning of the study, but the selected formulation provided the most consistent 3D-bioplotted structures.

2.3. Characterization of Nanofiber Dispersion within Alginate-Nanofiber Hydrogel To characterize the dispersion of the PLA nanofibers within the hydrogel, a disk-shaped acellular Alg-Nf specimen (Ø 12 x 6 mm) was cast using an open silicone mold on a glass substrate. The specimen was


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paraffin embedded using standard protocol, and three slices of 100 µm thickness were sectioned from different zones along the disk thickness (top, middle and bottom). The sections were then imaged (10x) using a Leica S8 APO microscope (Leica Microsystems, Germany). Six images were captured at random locations on each slice and post-processed in ImageJ. Each image was converted into 16-bit color format and thresholded to isolate fibers from the image. The nanofiber area within the alginate matrix was measured and the fiber area fraction (fiber area/total area) was calculated (N = 18 images). Finally, the dispersion of PLA nanofibers was calculated using the following equation61. % = 100 −

Where U is the mean fiber area fraction and L is the % volume of PLA nanofibers in acellular Alg-Nf.

2.4. Compression Testing of Alginate and Alginate-Nanofiber Hydrogels The bulk compressive properties of acellular Alg and acellular Alg-Nf were tested (n = 5/hydrogel) via unconfined uniaxial compression following an established protocol62,63. Test specimens were cast and crosslinked as disks (Ø 10 x 7 mm), loaded into the Universal Testing Machine (ATS 1620, Applied Test Systems Inc., Butler, PA) while submerged in an aqueous environment, and equilibrated under a 0.02 N creep load for 300 s. The specimens were then compressed to 10% strain at a constant rate of 0.05%/s, and the ramp modulus was determined from the slope of the resulting stress-strain curve. Next, while maintaining a strain of 10%, a stress relaxation test was performed for 1000 s, at which point the equilibrium modulus was calculated from the resulting load and specimen geometry. Finally, a 1% sinusoidal strain was applied at 1 Hz for 10 cycles to determine the dynamic modulus from the slope of the resulting stress-strain response. The mean moduli of the two hydrogels were compared using Student’s t-tests and statistical significance was determined at p < 0.05 (JMP Pro, SAS, Cary, NC).

2.5. Cell Expansion and Bioink Formulation



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Cryopreserved human stem cells (hASC) derived from adipose tissue (StemPro® R7788115, Thermo Fisher Scientific) were plated and expanded in T-75 tissue culture flasks with MesenPro RS basal medium and growth supplement (Thermo Fisher Scientific) for a total of 14 days (37.5°C, 5% CO2). To begin, a total of 500,000 cells at passage 6 were plated on day 0, and complete media changes were performed every 72 hours. The cells were passaged on day 7 after reaching an approximate confluency of 70%. These cultures resulted in a total yield of 22 million cells by day 14. All bioprinting experiments were performed on day 14 using bioinks that were encapsulated with these cells. The cells were suspended in 20 mL media, which was then split into four equal (5 mL) volume portions in separate tubes. This step was introduced into the protocol to make the cell pellet sizes manageable and easier to mix into acellular Alg or acellular Alg-Nf during the following step. All tubes were centrifuged for 10 minutes at 500 × g to obtain the four individual cell pellets. One pellet was added to 4 mL of acellular Alg and uniformly dispersed by repeated pipetting to obtain the Alg-hASC bioink. Using the same procedure, the remaining three cell pellets were sequentially mixed with 12 mL of acellular Alg-Nf to create the Alg-Nf-hASC bioink. Both bioinks were formulated to the same concentration of 1.375 x 106 cells/mL.

2.6. Digital Modeling and Bioprinting of Strands Five 6-well plates configured as per the schematic in Figure 2 were used for this set of experiments. One individual six-well plate (3 wells/plate) was assigned to bioprinted strands (5 strands/well) for each of acellular Alg, acellular Alg-Nf, Alg-hASC, and Alg-Nf-hASC. These plates were used for cell proliferation assessment. The last six-well plate had two wells, each containing bioprinted strands of either Alg-hASC or Alg-Nf-hASC. These strands were utilized for the cell viability assay.


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Figure 2. Schematic of the 6-well plate assignments and configurations used for bioprinting strands and assessing the proliferation and viability of encapsulated hASC. For the Live/Dead assay, two strands were randomly picked from each bioink group.

The five strands, each 25 x 0.5 x 0.5 mm with a 3 mm inter-strand spacing, within a single well were first modeled as a single STL file in MagicsRP (Materialise NV, Leuven, Belgium), which was then processed in BioplotterRP positioning and slicing software (EnvisionTEC, Gladbeck, Germany). A sixwell plate was selected as the positioning platform, and the STL file and two duplicates were aligned to fit within three wells. The grouped file was then sliced to create a single layer to facilitate the printing of strands. The sliced file was finally processed in Visual Machines software in preparation for bioprinting on the 3D-Bioplotter (Manufacturer Series, EnvisionTEC). The acellular and cell-encapsulated bioinks were filled into 30 cc cartridges (Nordson EFD, Westlake, OH) with 18 gage (inner Ø = 0.81 mm) precision tip nozzles (Nordson EFD). The cartridges were loaded into the two low temperature print-heads, and the print-heads and stage temperature were set to 37°C. The nozzle-stage Z-offset was set to 0.3 mm, and bioprinting was performed at an extrusion pressure of 0.02 N/mm2. Due to the difference in viscosity of alginate with and without nanofibers, the



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acellular Alg and Alg-hASC were printed at a nozzle speed of 15 mm/second, while the acellular Alg-Nf and Alg-Nf-hASC were printed at 5 mm/second. Post-printing, the strands were first cross-linked in 1 mL/well solution of sterile 1% CaCl2 (DI water + CaCl2 dihydrate, Sigma-Aldrich, MO) for 5 minutes and then washed twice with sterile PBS (pH 7.4, with MgCl2 and CaCl2, Thermo Fisher Scientific). Finally, 3 mL of chondrogenic differentiation media (CDM)64 containing vitamin C and TGFβ3 (Biolegend, San Diego, CA) was added to each well, and the plates were incubated (37.5°C, 5% CO2). The media added to the plates assigned for cell proliferation assessment contained 10% alamarBlue reagent (Thermo Fisher Scientific).

2.7. Digital Modeling and Bioprinting of Medial Knee Meniscus MRI scans of the right knee joint of a de-identified 24-year-old male were obtained from the Department of Orthopaedics at the University of North Carolina-Chapel Hill. The DICOM files were processed in Mimics Research (v18, Materialise NV) to better distinguish the medial meniscus from its surrounding areas. Contours of the meniscus were drawn every third layer in the coronal and sagittal views using the 3D-LiveWire tool, and the selection was region grown to generate a mask in all layers. Manual adjustments were performed to remove noise and undesired features, and a 3D model smoothed for optimal surface quality was generated and exported as an STL file. Post-processing was performed in SolidWorks 2015 (Dassault Systèmes, Waltham, MA) and Magics (v19, Materialise NV) to create intercalating layers of circumferentially oriented and linear parallel strand geometries. The file was sliced into 20 layers (each 0.482 mm thick) and positioned inside a petri dish platform in BioplotterRP. Finally, the file was processed in Visual Machines with an inter-strand axial spacing of 0.8 mm in each layer. The Alg-Nf-hASC meniscus construct was 3D-bioplotted using previously described (Section 2.6) process parameters: nozzle diameter (inner Ø = 0.81 mm), print-head and stage temperature (37°C), Zoffset (0.3 mm), extrusion pressure (0.02 N/mm2), and nozzle speed (5 mm/second). Crosslinking was performed by micro-pipetting sterile 1% CaCl2 solution over the printed bioink once every two layers. Post-printing, the complete meniscus construct was washed twice with sterile PBS (pH 7.4, with MgCl2


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and CaCl2), and 25 mL of CDM containing 1 µL/mL of vitamin C and TGFβ3 was added to the petri dish. The construct was incubated (37.5°C, 5% CO2) and media changes were performed every 72 hours.

2.8. Characterization of hASC Viability and Proliferation in Bioprinted Strands The Live/Dead assay (Thermo Fisher Scientific) was used to evaluate the viability of hASC within the bioprinted Alg-hASC and Alg-Nf-hASC strands (n = 2 strands/bioink) after 16 days in culture. Briefly, the strands were isolated from the assigned six-well plate, stained with 2 µL of 2 µM calcien AM and 8 µL of 4 µM EthD-I in 4 mL sterile PBS (with calcium), incubated for 10 minutes, and analyzed using a Leica DM5500B fluorescent microscope (Leica Microsystems). The colorimetric alamarBlue (aB) assay (Thermo Fisher Scientific) was used to assess the hASC proliferation activity in the bioprinted strand wells (n = 3 wells/bioink) at days 1, 4, 7, 10, 13 and 16. Wells of acellular Alg and acellular Alg-Nf strands (n = 3 wells/hydrogel) were used as controls and data was normalized to the appropriate control. Each media change for the acellular controls as well the hASC-encapsulated bioinks contained 10% by volume of the aB reagent. At each time point, three 100 µL samples were pipetted from each experimental well into a standard 96 well plate, and the absorbance was measured using a microplate reader (Tecan, Männedorf, Switzerland) with excitation and emission wavelengths of 570 and 600 nm, respectively. The absorbance data was converted to and is reported as % aB reduction65 over the six time points. Statistically significant differences in % aB reduction between Alg-hASC and Alg-Nf-hASC over the six time points were determined by two-way analysis of variance (ANOVA) at a significance level of p < 0.05 (JMP Pro).

2.9. Histology of the Knee Meniscus A small section near a horn of the bioprinted meniscus construct was excised after 4 weeks in culture and analyzed for cell viability using the Live/Dead assay (same protocol as Section 2.8). Additional small samples were harvested from the same zone after 4 and 8 weeks in culture and fixed in a solution of 10% buffered formalin (VWR International, Radnor, PA) and 0.05 M CaCl2 dihydrate for 30 minutes. The



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fixed samples were then dehydrated in a series of increasing ethanol concentrations, incubated overnight in paraffin (Paraplast Plus, McCormick Scientific, St. Louis, MO) at 60°C, and embedded in paraffin blocks that were left to solidify at room temperature. Using a microtome, 8 µm slices were sectioned and mounted onto microscope slides. Standard protocols were followed for hematoxylin and eosin (H&E; Thermo Fisher Scientific), picrosirius red (American MasterTech, Lodi, CA), and Alcian blue 8GX (Acros Organics, Fair Lawn, NJ) staining. Briefly, fixed sections were washed twice with SafeClear II cleaning agent (Fisher Scientific, Waltham, MA), rehydrated in a series of ethanol solution baths with decreasing concentrations, stained, then dehydrated in sequentially increasing ethanol concentrations, and finally washed twice in SafeClear II. To prevent alginate dissolution, 0.05 M CaCl2 dihydrate was added to all water and stain baths. Coverslips were mounted using xylene substitute (Thermo Fisher Scientific), and sections were imaged using an inverted microscope. Figure 3 shows a schematic of the Live/Dead and histology sectioning zone and defines the internal and external regions which were imaged and analyzed. Samples of H&E images (n = 6/region/time point; N = 24) were used to quantify and compare cell densities in the two regions. A twoway ANOVA and Tukey’s HSD post-hoc tests were performed to determine statistically significant differences between regions and time points (JMP Pro). Statistical significance was determined at p < 0.05.


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Figure 3. Schematic of typical imaging areas for stained sections highlighting the external (orange) and internal (blue) regions. (View image in color for clarity.)

3. RESULTS AND DISCUSSION: 3.1. Nanofiber Dispersion Characterization: SEM images of PLA nanofibers are shown in Figure 4. The diameter and length of the nanofibers was 499 ± 307 nm and 67 ± 50 µm, respectively. Given the current lack of standards for characterizing dispersion of nanofibers in hydrogels for tissue engineered medical products, we adopted protocols modified from the ASTM D2663-14 standards61 to characterize the PLA nanofiber distribution within the alginate matrix.



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Figure 4. Scanning electron microscope image of PLA nanofibers produced using the patented XanoShear process. The diameter and length of the nanofibers was 499 ± 307 nm and 67 ± 50 µm, respectively.

Figure 5 shows representative images, before and after ImageJ thresholding, from three random sections around the top, middle, and bottom regions of the acellular Alg-Nf disc. The mean fiber area fraction was 32.6% and the corresponding dispersion index was 34.80 ± 14.28%. The difficulties associated with the uniform dispersion of nanofibers (e.g., carbon, cellulose, PLA) within polymer matrices have been well-documented61,66-68 and can be attributed to the predominance of the Van der Waal’s forces of attraction between surfaces at the sub-micron length scale. The dispersion index could be improved in the future by investigating mixing protocols that utilize surfactants/dispersants to modify the nanofiber surface properties as reported in literature69-72.


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Figure 5. Representative optical and ImageJ thresholded images of acellular Alg-Nf sections used in the characterization of PLA nanofiber distribution in the alginate matrix.

3.2. Compressive Characteristics of Bulk Hydrogels: The ramp, equilibrium, and dynamic moduli of acellular Alg and acellular Alg-Nf (n = 5/hydrogel) determined from their stress-strain response curves from the unconfined uniaxial compression are presented in Figure 6. All three mean moduli of acellular Alg-Nf were higher than those of acellular Alg, but these differences were not statistically significant (pramp = 0.6565, pequilibrium = 0.5116, pdynamic = 0.5330). The mean ramp and equilibrium moduli are comparable to the values reported in literature for alginate-based hydrogels and composites ranging in concentration from 1.5 to 3%73-79. The dynamic moduli are higher than those reported in literature, and these differences can be likely attributed to the differences in hydrogel formulation, crosslinking and testing protocols. These results confirm that addition of the PLA nanofibers did not adversely affect the properties of native alginate hydrogel. Future



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investigations should focus on the characterization, at both the bulk and matrix level, of compressive moduli of cell-seeded bioink constructs in culture over time to evaluate increases in biomechanical properties after ECM formation.

Figure 6.

Ramp, equilibrium and dynamic moduli

determined from unconfined uniaxial compression testing of the two hydrogels. The mean moduli of acellular AlgNf were higher than those if acellular Alg but the differences were not statistically significant.

3.3. hASC Viability and Proliferation in Bioprinted Strands: The width of the 3D-bioplotted Alg-hASC and Alg-NF-hASC strands before crosslinking was 852.33 ± 15.56 µm and 855.66 ± 38.83 µm, respectively. Representative optical microscope images of bioprinted strands show a uniform distribution of cells in the crosslinked matrices for both Alg-hASC and Alg-NfhASC bioinks (Figure 7a and b, respectively). Agglomerations of PLA nanofibers can also be observed


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in the Alg-Nf-hASC strands. The Live/Dead assay was performed for the qualitative assessment of hASC viability in these bioprinted stands while the aB assay was performed to quantify their cellular metabolic activity.

Figure 7. (a,b) Optical, and (c,d) Live/Dead fluorescence images (after day 16) of bioprinted Alg-hASC and Alg-Nf-hASC strands; live cells appear green, dead cells appear red. The white arrows in (b) and (d) indicate agglomeration of PLA nanofibers. (View image in color for clarity.)

Representative fluorescence images showing live and dead cells (green and red, respectively) within the Alg-hASC and Alg-NF-hASC strands after 16 days in culture are shown in Figure 7c, d. These images demonstrate that the vast majority of hASC encapsulated within both bioinks remained viable after bioprinting and 16 days in culture. Note that there was some background staining of the other bioink



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components while acquiring these images. For example, the two red clusters in Figure 7d are not dead cells but PLA nanofiber agglomerations. Results of the aB assay are shown in Figure 8. The mean % aB reduction in the bioprinted AlghASC and Alg-Nf-hASC strand wells (normalized to respective acellular controls) at days 1, 4, 7, 10, 13 and 16 is reported. The aB assay uses a non-toxic cell permeable compound called resazurin that, upon entering live cells, is reduced to resorufin. The amount of reduction, which can be determined based on the absorbance, is proportional to the metabolic activity of cells80. A decrease in % aB reduction reading may indicate a decrease in cell proliferation or a reduction in the overall metabolic activity which can occur during cell differentiation.

Figure 8. Results of alamarBlue assay from bioprinted Alg-hASC and Alg-Nf-hASC strands over 16 days in chondrogenic differentiation medium culture. Two-way ANOVA showed that the main effects of the type of bioink and days in culture on % aB reduction were significant (p < 0.0001), but their interactions were not (p = 0.2246). * indicates statistically significant difference (p < 0.001) in the % aB reduction compared to the previous time point.


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Alg-Nf-hASC consistently showed higher % aB reduction than Alg-hASC at every time point, but the general trends in the readings over the 16 day time period were similar for both bioinks. Two-way ANOVA showed that the main effects of both the type of bioink and days in culture on the % aB reduction were significant (both p < 0.0001). However, their interactions were not significant (p = 0.2246). Post hoc Tukey HSD analysis of % aB reduction between consecutive days in culture showed that the reduction was significantly different between days 1 and day 4 (p < 0.0001), but not between any other consecutive days. For both Alg-hASC and Alg-Nf-hASC, the highest increase in % aB reduction between consecutive days, which was observed between days 1 and 4, was 94.2% and 115.5%, respectively. At the peak of proliferation in both bioinks on day 7, the % aB reduction in Alg-Nf-hASC was 28.5% higher than that in Alg-hASC. Given that both bioinks were formulated with the same initial cell concentration (1.375 x 106 cells/mL) and the lack of statistically significant difference in % aB reduction between them on day 1 (p = 0.3877), these results indicate that the bioink with the PLA nanofiber suspension provided a more conducive environment for hASC and aided their proliferation. These results confirm that the hASC were able to endure the shear stresses during 3D-Bioplotting and continue to exhibit healthy metabolic activity over the following week. Multiple phenomena could have been responsible for the drop in the % aB reduction after the day 7 peak. First and foremost, static culturing conditions and media change intervals could, together, affect the nutrient diffusion into the strands over time. Secondly, although completely submerged within the media initially, we observed some strands partially floating on top, starting at day 10. This would have adversely affected the nutrient exchange process, which in turn could have caused reduction in cellular metabolic activity. Improving the culture setup and conditions can be expected to help overcome these limitations and improve the cell proliferation in future studies. Finally, given that the strands were cultured in CDM, the reduction in cellular metabolic activity can also be attributed, in part, to the chondrogenic differentiation of hASC. Xu et al. observed a similar reduction in metabolic activity of hMSC encapsulated within alginate during chondrogenesis, and reported a 76.9% reduction in proliferation on day 24 in comparison to day 081.



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3.4. hASC Viability and Histology of Bioprinted Meniscus: The bioprinted Alg-Nf-hASC meniscus construct is shown in Figure 9a. A distribution of cells and nanofiber agglomerations similar to the strands can be observed in the microscope image (Figure 9b). Note that the lower contrast in the meniscus images is due to differences in light penetration through the thicknesses of the multi-layered meniscus and the single-layered strands, and is not an indicator of variability in the quality of the bioink formulation.

Figure 9. (a) The Alg-Nf-hASC meniscus construct that was designed starting with a patient’s MRI images and bioprinted on a 3D-Bioplotter. (b) Microscope image of an excised section shows distribution of cells (white arrows) and agglomeration of nanofibers. (c) and (d) Live/Dead images of external and internal regions, respectively, of an excised section after 4 weeks in culture. (View image in color for clarity.)


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The hASC viability in the bioprinted meniscus construct after 4 weeks in culture was assessed using the Live/Dead assay, and there was a noticeable difference between the external and internal regions of the construct (Figure 9c and 9d, respectively). The markedly better viability in the external region of the construct is attributed to its proximity with the culture media. Given the static culturing conditions over 4 weeks, a drop in nutrient diffusion, and hence, lower cell viability, was expected within the core regions of the construct. After 4 and 8 weeks in culture, sections of the construct were randomly assigned to one of three staining protocols: H&E to visualize cytoplasmic, nuclear, and extracellular components82, picrosirius red to assess cell-secreted collagen83, and Alcian blue to evaluate chondrogenic differentiation of hASC by staining cell-secreted proteoglycans84. Representative images of internal and external regions for all three protocols at both the time points are presented in Figure 10. The varying levels of background staining of the alginate hydrogel matrix seen here are consistent with literature85-87. Furthermore, small crystalized artifacts seen in some images were potentially caused by the addition of CaCl2 in the stains and water baths to prevent the dissolution of the thin hydrogel sections. These images demonstrate that at both time points, outer peripheral regions of the construct yielded larger hASC populations when compared to the internal regions. This is consistent with observations from the Live/Dead assay and points to the limitations of static culturing and nutrient diffusion into and waste transportation out of the construct core. A two-way ANOVA showed statistically significant differences in cell densities between regions (p < 0.0001) quantified from H&E images across both sampled regions of the construct. The interaction between the factors was also significant, and posthoc Tukey’s HSD analyses show that the external regions were 53% more densely populated than the internal regions at 4 weeks (p = 0.0015). This difference was more pronounced, 147%, at 8 weeks (p < 0.0001). However, the cell density differences were deemed not significant within an individual region over the two time points (pinternal = 0.0854, pexternal = 0.1892). These results imply that static culturing over a longer time interval did not necessarily improve hASC proliferation and density in the anatomically



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sized meniscus construct, but the location and direct proximity to the media had a significant effect on the hASC population.

Figure 10. Representative microscope images of stained meniscus construct sections after 4 (a, b, e, f, i, j) and 8 (c, d, g, h, k, l) weeks in culture. H&E staining (a-d) allowed for visualization of cells and matrix within the construct. Picrosirius red (e-h) and Alcian blue (i-l) staining allowed for visualization of cell-secreted collagen and proteoglycans, respectively. Black arrows indicate the edge of the construct. Red arrows indicate instances of crystalized artifacts


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Picrosirius red staining (Figure 10e-h) demonstrated dense collagen formation immediately surrounding individual cells. A similar effect could be observed for the proteoglycans stained with Alcian blue (Figure 10i-l), indicating the differentiation of hASC into chondrocyte-like cells. However, interconnected networks of ECM were not evident indicating that the cell-secreted ECM components were unable to migrate across and interweave within the alginate matrix even after 8 weeks. Similar effects have been observed in previous studies with alginate and hyaluronic acid hydrogels88-90, indicating that bioinks prepared at high w/w hydrogel concentrations and relatively low cell seeding densities may “trap” secreted ECM and prevent formation of a homogenous tissue-like structure. Results of the viability and histology assays confirm that the hASC were able to sustain through the bioprinting process and remain viable in the construct, even though there were differences between the external and internal regions. Chondrogenic differentiation and ECM production, both of which are necessary for the formation of the meniscal tissue, were also evident. In future studies, the differences in viability between the internal and external regions could be likely overcome by dynamic culturing91-93 (e.g., inside a perfusion bioreactor or orbital shaker) and performing more frequent media changes. Another potential strategy that can be explored in order to improve the nutrient perfusion is the design of constructs with lattice architectures similar to polymer scaffolds. Additionally, using lower molecular weight alginates and/or decreasing the w/w concentration of alginate in the bioink could improve the diffusion kinetics of such thick constructs, and eventually lead to better cell viability and proliferation94,95 and better distribution of the newly formed ECM89. In conjunction, preparing the bioinks with higher cell seeding densities96,97 can potentially accelerate the formation of an interconnected ECM network. Finally, the impact of the nanofibers on the biochemical and biomechanical properties of the constructs should also be investigated. In particular, the alignment of the nanofibers may induce local mechanical anisotropy. Tuning of this variable could allow for directiondependent mechanical properties similar to musculoskeletal tissues. Together, these alternatives will promote the formation of engineered meniscus tissue that closely resembles the native structure and properties.



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4. CONCLUSION AND FUTURE DIRECTIONS: This study focused on the pneumatic extrusion-based 3D-bioprinting of a nanofiber-hydrogel bioink and the ability to engineer a viable, patient-specific musculoskeletal soft tissue construct. Results from the aB assay showed a significant difference in % aB reduction between the 3D-bioplotted Alg-hASC and Alg-NfhASC strands over 16 days in CDM culture, which indicates that the presence of PLA nanofibers in the alginate matrix aided hASC metabolic activity and proliferation. Results of the Live/Dead and histochemical staining assays on the 3D-bioplotted full-scale meniscus construct confirmed the ability of bioprinted hMSC to differentiate down the chondrogenic pathway and secrete a matrix consisting of proteoglycans and collagen, but also highlighted the limited nutrient perfusion through large constructs and the lack of diffusion of the cell-secreted matrix through the hydrogel. Future studies should address these fundamental issues to improve the cell viability within the bioprinted construct core and facilitate formation of an interwoven ECM network by investigating factors such as the hydrogel composition and concentration, nanofiber design and concentration, porous construct architecture, as well as dynamic culturing techniques. These factors are likely to impact several characteristics of interest including cell morphology, proliferation levels, ECM alignment, and biomechanical properties of engineered tissues. Taking advantage of non-intrusive imaging modalities such as MRI, computer aided design and manufacturing (CAD/CAM modeling), and additive manufacturing, patient-specific constructs can be engineered to fulfil specific needs. However, the manufacturability of different bioink compositions and the shape fidelity of the 3D-bioprinted constructs should be further investigated. From a bioprinting perspective, we envision the development of new process technologies that can manipulate and control the distribution and alignment of nanofibers within bioprinted constructs to stimulate the formation of anisotropically oriented ECM that mimics the native fibrous organization of musculoskeletal soft tissues. Once in vitro evaluations provide enough fundamental evidence of safety and efficacy, further exploration in animal models can lead the pathway for viable translation of such engineered constructs to human clinical treatments.


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Acknowledgements: The authors acknowledge the assistance of Dr. Harvey West (Center for Additive Manufacturing and Logistics, NC State University) for the mechanical testing experiments. The authors thank Xanofi Inc., Raleigh, NC for donating experimental materials and supplies. The study was also supported in part by grants from the Kenan Institute for Engineering, Technology and Science (Shirwaiker, Fisher and Spang), and NC State Comparative Medicine Institute – Functional Tissue Engineering pilot programs (Shirwaiker, Starly and Fisher).



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