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Oxygen-Releasing Microparticles for Cell Survival and Differentiation Ability under Hypoxia for Effective Bone Regeneration Ho Yong Kim, So Young Kim, Hye-Young Lee, Jin Ho Lee, Gyu-Jin Rho, Hyeon-Jeong Lee, Hee-Chun Lee, June-Ho Byun, and Se Heang Oh Biomacromolecules, Just Accepted Manuscript • DOI: 10.1021/acs.biomac.8b01760 • Publication Date (Web): 14 Jan 2019 Downloaded from http://pubs.acs.org on January 17, 2019

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Oxygen-Releasing Microparticles for Cell Survival and Differentiation Ability under Hypoxia for Effective Bone Regeneration

Ho Yong Kim,1,† So Young Kim,1,† Hye-Young Lee,† Jin Ho Lee,‡ Gyu-Jin Rho,§ Hyeon-Jeong Lee,§ Hee-Chun Lee,¶ June-Ho Byun,*,# Se Heang Oh*,†,ǁ

†Department of Nanobiomedical Science, Dankook University, Cheonan 31116, Republic of Korea. ‡Department

of Advanced Materials and Chemical Engineering, Hannam University, Daejeon

34054, Republic of Korea, §Department

of Theriogenology and Biotechnology, College of Veterinary Medicine, Gyeongsang

National University, Jinju 52727, Republic of Korea. ¶Department

of Veterinary Medical Imaging, College of Veterinary Medicine, Gyeongsang

National University, Jinju 52727, Republic of Korea. #Department

of Oral and Maxillofacial Surgery, Gyeongsang National University School of

Medicine, Gyeongsang National University Hospital, Institute of Health Sciences, Gyeongsang National University, Jinju 52727, Republic of Korea. ǁDepartment

of Pharmaceutical Engineering, Dankook University, Cheonan 31116, Republic of

Korea

ACS Paragon Plus Environment

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Keywords: Oxygen delivery; perfluorooctane; hollow particles; osteogenic differentiation; bone regeneration

ABSTRACT: Sufficient oxygen delivery into tissue-engineered three-dimensional (3D) scaffolds to produce clinically applicable tissues/organs remains a challenge for researchers and clinicians. One potential strategy to overcome this limitation is the use of an oxygen releasing scaffold. In the present study, we prepared hollow microparticles (HPs) loaded with an emulsion of the oxygen carrier perfluorooctane (PFO; PFO-HPs) for the timely supply of oxygen to surrounding cells. These PFO-HPs prolonged the survival and preserved the osteogenic differentiation potency of human periosteal-derived cells (hPDCs) under hypoxia. hPDCs seeded onto PFO-HPs formed new bone at a faster rate and with a higher bone density than hPDCs seeded onto phosphate buffered saline-loaded control HPs. These findings suggest that PFO-HPs provide a suitable environment for the survival and maintenance of differentiation ability of hPDCs at bony defects without vascular networks until new blood vessel ingrowth occurs, thus enhancing bone regeneration. PFO-HPs are a promising system for effective delivery of various functional cells, including stem cells and progenitor cells, to regenerate damaged tissues/organs.


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INTRODUCTION Insufficient oxygen supply to cells in tissue-engineered three-dimensional (3D) scaffolds may be the result of limited oxygen diffusion and/or slow ingrowth of blood vessels; this low oxygen environment (hypoxia) poses the biggest challenge to producing clinically applicable tissues/organs at relevant volumes.1-4 Cells grown in hypoxic environments produce more free radicals and require 15 times more glucose to produce the same amount of adenosine triphosphate (ATP) than cells grown in environments with normal oxygen levels. Thus, hypoxic cells fail to survive as a result of deoxyribonucleic acid (DNA) damage, cell cycle arrest, and rapid glucose depletion.5,6 Although the dissolved oxygen in blood has a limited diffusion distance (100–200 μm), it is effectively supplied to cells in native tissues via the fluid-rich extracellular matrix (ECM)7 and very dense capillary networks. However, in engineered tissues, the cells in the central regions of 3D scaffolds can be several hundred microns away from oxygen sources, such as cell culture media in vitro or blood in vivo, and thus numerous cells in the center experience hypoxia. As a result, heterogeneous tissues are generated, with necrotic cells in the central region and viable cells on the peripheral region. It has been commonly accepted that a maximum volume of 1 cm3 is the limit to prevent necrotic centers in non-vascularized tissue-engineered 3D scaffolds.1,8 To overcome the insufficient oxygen supply in scaffolds, several strategies have been studied extensively and produced promising outcomes. These include the use of angiogenic factors [vascular endothelial growth factor (VEGF), basic fibroblast growth factor (bFGF), and endothelial cells],9-13 and the use of tissue-engineered constructs with prevascular networks.14,15 Nevertheless, the slow ingrowth of blood vessels in scaffolds even with angiogenic factors16 and the lack of connection between


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the vascular network in the scaffold and the host system17 are challenges that need to be overcome to produce clinically applicable, large tissue masses. In recent years, scaffolds incorporating oxygen-generating materials, which supply oxygen throughout the whole scaffold regardless of dimension, have been shown to provide suitable environments for cell survival until blood vessel ingrowth occurs and have increasingly gained interest as an alternative strategy. These oxygen-generating materials include liquid hydrogen peroxide, which decomposes into oxygen and water,18 and solid peroxides such as sodium percarbonate19,20 and calcium peroxide,21-23 which dissociate into ions and hydrogen peroxide in aqueous solution and subsequently dissociate to form oxygen. However, it has been shown that an initial uncontrolled burst of oxygen production occurs (hyperoxia), which leads to increased levels of cytotoxic reactive oxygen species (ROS), peroxides, and peroxide by-products.24,25 Encapsulation of oxygen-generating peroxides in polydimethysiloxane (PDMS)26 or in poly(methyl methacrylate) (PMMA)27 or complexation of oxygen-generating peroxides with poly(vinyl pyrrolidone) (PVP)28 has been shown to temper the oxygen-producing hydrolytic reaction and to prevent the contact of cytotoxins with the cells. Hemoglobin-based oxygen carriers have been also considered as a strategy for cell survival in hypoxia.29-32 Recently, perfluorocarbons (PFCs), which have high solubility of oxygen and carbon dioxide and allow appropriate and controlled release of oxygen to cells without hypoxia and hyperoxia, have been used as oxygen carriers in tissue engineering.33 Initially, PFCs were embedded with target cells in alginic acidbased34 or fibrin-based35 hydrogels. Later, hydrogels were fabricated with fluorinated polymers, such as chitosan,36 hyaluronic acid,37 or fluorinated microparticles.38 Although in vitro experiments have shown that PFC-based oxygen carriers have the potential to solve the problem of insufficient oxygen delivery in tissue-engineered 3D scaffolds and to reduce the toxic side


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effects of oxygen-generating materials, only a few in vivo studies have evaluated whether PFCs can supply enough oxygen to maintain cell viability until blood vessel penetration to produce sound tissues and organs. In our previous study,39 we developed a PFC-based system with perfluorooctane emulsionloaded hollow microparticles (PFO-HPs). We demonstrated that the PFO-HPs provided sufficient oxygen to the surrounding cells for 10 days under hypoxia and that the initially seeded number of cells on PFO-HPs in vitro is maintained. Moreover, the implanted cells on PFO-HPs were survived for 14 days until blood vessel penetration in vivo. The aim of this study was to investigate whether PFO-HPs can provide a proper environment for cell survival at defective regions without vascular networks under hypoxia and for the maintenance of cell properties, such as the differentiation ability and target tissue regeneration (Figure 1). To the best of our knowledge, this is the first report of an approach using a PFC-based oxygen carrier for target tissue regeneration. Human periostealderived cells (hPDCs) with the ability to differentiate into osteoblasts40 were selected as model cells. The osteogenic differentiation of hPDCs that had been pre-incubated under hypoxia for 10 days on PFO-HPs was compared with the differentiation of those on control HPs in normoxia (normal level of oxygen). As an in vivo animal model, bone regeneration upon implantation of hPDCs/PFO-HPs or hPDCs/phosphate buffered saline (PBS)-HP in miniature pigs with mandibular defects was compared.

EXPERIMENTAL SECTION Materials. Poly-ε-caprolactone (PCL; MW 80,000 Da; Sigma-Aldrich, St. Louis, MO, USA), ethyl acetate (EA; Sigma-Aldrich), Pluronic F127 (MW 12,500 Da; Sigma-Aldrich), and


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poly(vinyl alcohol) (PVA; MW 85,000-124,000 Da; hydrolysis, 87-89%; Sigma-Aldrich) were used to prepare PCL hollow microparticles (PCL HPs). Perfluorooctane (PFO), Pluronic F68 (MW 8,400 Da), and L-α-phosphatidylcholine (egg yolk phospholipid; EYP) were purchased from Sigma-Aldrich to fabricate the PFO emulsion. Water was purified using an EXL®3 pure & ultrapure water system (Vivagen, Korea). The PCL HPs were sterilized with ethylene oxide and the PFO emulsion was autoclaved41 for in vitro cell culture and in vivo animal studies. Fabrication of PFO-HPs. PFO emulsion-loaded hollow microparticles (PFO-HPs) were fabricated by water-in-oil-in-water (W/O/W) emulsion solvent evaporation for the formation of HPs and infiltration of the PFO emulsion into the HPs as described previously (Figure S1).39 To prepare PCL HPs, a 1% (w/w) Pluronic F127 aqueous solution was added to a 5% (w/w) PCL EA solution with stirring at 400 rpm to emulsify the mixture. The emulsified mixture was added dropwise to a 0.5% (w/v) PVA aqueous solution with gentle stirring at 400 rpm, and the droplets were solidified for 10 min. The solidified PCL HPs were washed with excess water, separated by size (100–500 μm) using standard testing sieves (Chunggye Industrial Co., Korea), and freeze-dried. The morphology of the PCL HPs was observed by scanning electron microscope (SEM; S-4300; Hitachi, Japan). To prepare the PFO emulsion, a PFO solution, a 20% (w/v) Pluronic F68 solution, and a 4% (w/v) EYP solution [PFO/Pluronic F68/EYP, 6/2/2 (v/v/v)] were vigorously agitated using an ultrasonic wave homogenizer (Branson Sonifier model 185, Danbury, CT, USA; 30 sec ON and 20 sec OFF; 10 cycles; frequency, 18,000 Hz). The average size of the fabricated PFO emulsion was determined at room temperature using a particle size analyzer (APA5001SR, Malvern Instruments, UK). To load the PFO emulsion into the PCL HPs, the PFO emulsion was added to a syringe filled with PCL HPs. The PFO emulsion infiltrated into the PCL HPs by positive pressure when the syringe plunger was pushed in. The control PBS-loaded PCL HPs (PBS-HPs)


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were prepared with PCL HPs and PBS using the same procedure. Infiltration of the PFO emulsion or PBS into HPs was performed aseptically. The total loading amounts of oxygen per milliliter of PFO-HPs and PBS-HPs were 23.62 ± 1.01 μg and 12.98 ± 0.01 μg, respectively, as determined in our previous study.39 Cell Culture in Hypoxia. To investigate cell survival and cell responses under hypoxia using PFO-HPs and PBS-HPs, we used hPDCs isolated from periosteal tissues of patients. The cell isolation procedure42 was approved by the Ethics Committee of Gyeongsang National University Hospital (GNUH 2014-05-012). To saturate the PFO emulsion or the PBS in the HPs with oxygen, the HPs (200 μL/well) were placed in ultra-low-attachment 24-well PS plates (Corning, Corning, NY, USA) and incubated in an aseptic oxygen chamber (7.5 x 6.0 x 4.0 cm3; O2 flow rate, 5 mL/h) for 5 min. Immediately, 500 μL of cell suspension [passage 3; cell density, 2.0 x 105 cells/mL; DMEM supplemented with 10% fetal bovine serum (FBS; Gibco, Waltham, MA, USA), 100 U/mL penicillin (Gibco), and 100 μg/mL streptomycin (Invitrogen, Carlsbad, CA, USA)] was seeded onto each oxygen-saturated group of HPs and incubated in a regular incubator (21% O2, 5% CO2, 37°C) with shaking at 50 rpm for 12 h for cell adhesion onto the HPs. Then, the cell-adhered HPs were transferred to new ultra-low-attachment 24-well PS plates, 2 mL of culture medium was added to each well, and the plates were incubated in a hypoxic incubator (1% O2, 5% CO2, 37oC) for 14 days. During the cell culture period, the culture medium was not changed to prevent reoxygenation of the medium; however, excess culture medium was used to prevent cell starvation and the drop in pH caused by cell metabolism. The numbers of viable cells on the HPs after cell culture in a hypoxic environment for 0, 1, 3, 5, 7, 10, and 14 days were estimated using Cell Counting Kit-8 (CCK-8; Dojindo, Japan).


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To estimate the responses of cells adhered to PFO-HPs and PBS-HPs and cultured in a hypoxic environment, caspase3/7 activity, which is an indicator of apoptotic cell death,43 and RNA expression of hypoxia-inducible factor 1α (HIF-1α)44,45 were detected using the Caspase-Glo® 3/7 assay (Promega, Madison, WI, USA) and quantitative real-time polymerase chain reaction (qRTPCR) analysis, respectively. For the qRT-PCR analysis, RNA was extracted from cells cultured for 0, 1, 3, 5, 7, 10, and 14 days under hypoxia using TRIzol reagent (Molecular Research Center, Cincinnati, OH, USA), and RNA was converted into cDNA using a First Strand cDNA synthesis kit (Applied Biosystems Inc., Foster City, CA, USA). Prepared cDNA and probe [HIF-1α, catalog no. Hs00153153_m1; glyceraldehyde 3-phosphate dehydrogenase (GAPDH), catalog no. Hs02758991-g1 (ThermoFisher Scientific, Waltham, MA, USA)] were mixed with Taqman® universal PCR master mix (Applied Biosystems Inc.), and qRT-PCR was performed using the StepOnePlusTM real-time PCR system (ThermoFisher Scientific). Data were normalized to GAPDH expression levels. To visualize HIF-1α expression and cell distribution in each HP group, immunocytochemical staining was conducted on cells cultured for 0, 3, 5, 7, 10, and 14 days under hypoxia. The cells, which were adhered on HPs, were washed briefly with PBS, fixed with 4% paraformaldehyde (in PBS) for 20 min, permeabilized with 0.1% Triton X-100 solution (in PBS), and blocked with 10% bovine serum albumin (BSA; in PBS). Subsequently, the cells were incubated with anti-HIF-1α antibody (1:100 dilution; Santa Cruz Biotechnology, Dallas, TX, USA) for 12 h at 4°C followed by incubation with goat anti-rabbit IgG (1:10,000 dilution; Jackson Immuno Research, West Grove, PA, USA) secondary antibody for 1 h at room temperature. The actin cytoskeleton and nuclei were visualized using Alexa 568-labeled phalloidin (ThermoFisher Scientific) and 4′,6-


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diamidino-2-phenylindole (DAPI; Vector Laboratories, Burlingame, CA, USA), respectively. Images were obtained using a confocal microscope (LSM 700, Carl Zeiss, Germany). Osteogenic Differentiation of hPDCs underwent Hypoxia. To investigate whether PFO-HPs provide a suitable environment for the maintenance of osteogenic differentiation potency of hPDCs under hypoxia, hPDCs seeded on PFO-HPs were incubated under hypoxia for 10 days. Then, the culture medium was replaced with 1 mL of osteogenic medium [DMEM supplemented with 10% FBS, 50 µg/mL L-ascorbic acid 2-phosphate (Sigma-Aldrich), 10 nM dexamethasone (Sigma-Aldrich), and 10 mM β-glycerophosphate (Sigma-Aldrich)40,42] and cultured in a regular incubator under normoxia for 4 weeks (Hyp-Nor/PFO-HP). To compare osteogenic differentiation with hPDCs that have not undergone hypoxia (control group), hPDCs (passage 3) seeded on PBSHPs (Nor/PBS-HP) were also cultured in osteogenic medium in a regular incubator. The overall cell culture scheme is shown in Figure 1. Quantification of Alkaline Phosphatase (ALP) Activity and Calcium Deposition. At 0, 1, 2, and 4 weeks of cell culture in normoxia, hPDCs from each group (Hyp-Nor/PFO-HP and Nor/PBS-HP) were rinsed with PBS, harvested, and assayed for ALP enzyme activity using a TRACP & ALP activity kit (Takara, Japan). ALP enzyme activity was quantified by measuring absorbance at 405 nm using a plate-reader (Spark 10M, Tecan, Switzerland). ALP activity was normalized to cellular DNA content, which was measured using a PicoGreen dsDNA quantitation kit (Molecular Probes, Eugene, OR, USA). To estimate calcium deposition by hPDCs at 0, 1, 2, and 4 weeks after cell culture, the hPDCs from each HP group were rinsed with Tyrode’s balanced salt solution and decalcified using 300


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μL of 0.6 N HCl for 24 h at room temperature. The calcium content in the resulting solution was measured using a calcium colorimetric assay kit (calcium C-test Wako, Japan). Quantitative RT-PCR Analysis. At 0, 1, 2, and 4 weeks after cell culture, RNA extraction and cDNA synthesis were performed as described in section 2.3. Prepared cDNA and probe [Runx2, catalog no. Hs00231692-m1; osteocalcin, catalog no. Hs00609452-g1, GAPDH, catalog no. Hs02758991-g1 (ThermoFisher Scientific)] were mixed with Taqman® universal PCR master mix, and qRT-PCR was performed using the StepOnePlusTM real-time PCR system. Data were normalized to GAPDH expression levels. Immunocytochemical Analysis. To visualize Runx2 and osteocalcin expression in each HP group, immunocytochemical staining was conducted at 0, 1, 2, and 4 weeks after cell culture. The cells, which were adhered on HPs, were washed briefly with PBS, fixed with 4% paraformaldehyde (in PBS) for 20 min, permeabilized with 0.1% Triton X-100 solution (in PBS), and blocked with 10% BSA in PBS. The cells were then incubated with anti-Runx2 antibody (1:100 dilution; Abcam, Cambrige, MA, USA) and anti-osteocalcin antibody (1:100 dilution; R&D Systems, Minneapolis, MN, USA) for 12 h at 4C. The cells were subsequently incubated with DyLight 488 (1:200 dilution; Abcam; for RUNX2) and NorthernLights 557 (1:200 dilution; R&D Systems; for osteocalcin) secondary antibodies for 1 h at room temperature. Cell nuclei were stained with DAPI, and images were obtained using a confocal laser microscope (LSM 700). Animal Study. To determine whether PFO-HPs can supply sufficient oxygen for the survival of hPDCs until blood vessel ingrowth occurs and whether PFO-HPs can facilitate new bone formation by differentiation of hPDCs into bone cells, we used miniature pigs with mandibular defects as animal models. This animal study was approved by the Animal Center for Biomedical


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Experimentation at Gyeongsang National University (GNU-160913-P0047). Three miniature pigs that were 6-12 months old and weighed approximately 25 kg were used. Under general anesthesia with 4 mg/kg of azaperone and 10 mg/kg of tiletamine-zolazepam, the mandibular body and ramus were exposed through a submandibular incision, and three bony defects (3 mm depth and 12 mm diameter) were created for each animal. The specimens were divided into the following three groups: hPDC-seeded PFO-HPs (hPDCs/PFO-HP), hPDC-seeded PBS-HPs (hPDCs/PBS-HP; no oxygen supply), and PFO-HPs without hPDCs (PFO-HP). The hPDCs were seeded onto PFO-HPs and PBS-HPs, as described above in section 2.3. Each of the three specimens (~0.35 mL) were implanted into each bony defect (Figure S2) in each pig, and the wounds were closed in two layers using 3-0 vicryl and 3-0 nylon sutures (Ethicon, Somerville, NJ, USA). First-generation cephalosporin was injected into the pigs intramuscularly twice a day for 7 days. Radiographic images were taken and evaluated at 0, 4, 7, and 10 weeks after implantation. The miniature pigs were under general anesthesia with tiletamine-zolazapam during the plain radiograph and computed tomography (CT) scans. CT images were acquired with a helical dual slice CT scanner (Somatom Emotion Duo; Siemens AG, Germany) with a slice thickness of 1.0 mm and a pitch of 1. Images were collected on a Lucion image post processing system (Infinitt Technology, Korea). Mandible regions of interest were reformatted with 3D reconstruction to evaluate the healing process of the bony defects in the mandible. Hounsfield units (HU) were also obtained from CT scans to evaluate the mineral density of the regenerated bone. Pixel data from DICOM CT images were used to calculate HU by confining elliptical regions of interest (ROI) to regenerated bone areas to prevent averaging the volumes from adjacent cortical bone. HU values were determined for elliptical ROIs (0.4 cm2) on CT images based on the growth rate of the miniature pig and the area of the regenerated bone. Ten weeks after implantation, the animals were sacrificed and the


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mandibles were stained with hematoxylin and eosin (H&E) for histological observations using a light microscope (CKX41, Olympus, Japan). Statistical Analysis. Data obtained were expressed as mean ± standard deviation. Statistical analyses were performed using IBM SPSS statistics 22 (IBM, Armonk, NY, USA). Data were evaluated by one-way analysis of variance (ANOVA) with Tukey’s multiple comparison test. We considered p values < 0.05 to be statistically significant.

RESULTS AND DISCUSSION Morphology of PCL HPs. The PCL HPs that function as reservoirs for the oxygen-carrying PFO emulsion were prepared by the W/O/W emulsion solvent evaporation technique, and the PFO emulsion was prepared by ultra-sonication.39 The prepared PCL HPs were 100–500 μm spherical structures with an empty inner space like a balloon and a membranous shell ~10 μm in thickness (pore size, ~2 μm), and the PFO emulsion had the average size of ~300 nm (Figure 2). In our previous study,39 we reported that the pores in the membrane allow penetration of the PFO emulsion into the inner hollow space under positive pressure, and that the emulsions are stably entrapped in the PCL HPs. It has been shown that the PFO emulsion has high oxygen solubility due to van der Waals interactions between PFO and oxygen,46 and the oxygen loaded in the PFO emulsion can be liberated in low-oxygen environments according to Henry's law, which explains that the solubility of a gas is proportional to the partial pressure.47 These phenomena are interpreted that the PFO emulsion can provide oxygen to surrounding cells in low-oxygen environments without releasing a burst of oxygen (hyperoxia), which increases the ROS level. Therefore, we hypothesized that our PFO-HPs would provide a suitable environment for hPDC survival and


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maintenance of osteogenic differentiation potency in bone defects until establishment of blood vessel ingrowth, thus enhancing bone regeneration. Cell Survival in Hypoxia. The proliferation of hPDCs that were seeded onto PFO-HPs and PBS-HPs and cultured in a hypoxic incubator for 14 days was compared using the CCK-8 assay [Figure 3(A)]. The number of viable hPDCs on PFO-HPs increased up to ~2.5 fold after 5 days, indicating that PFO-HPs provided a suitable oxygen environment for cell survival and proliferation for 5 days of hypoxia. After 5 days, the viable cell number decreased over time because of the limited oxygen in the PFO emulsion. However, more cells (5.0 x 104 cells/well) remained on PFOHPs after 14 days of culture than were initially seeded (3.8 x 104 cells/well), whereas the number of cells on PBS-HPs decreased sharply after 3 days of hypoxic culture and subsequently increased only slightly due to the low oxygen in the medium, which could not support hPDC survival, proliferation, and adaptation in a hypoxic environment.48 In a previous study,39 we verified that the PFO-HPs supply notably greater oxygen to cell (fibroblasts) than PBS-HPs in a hypoxic incubator. The PFO-HPs allowed suitable oxygen concentration for cell growth by 3 days (oxygen concentration in medium, >5.0 mg/L) then gradually decreased, and eventually the oxygen was exhausted by 10 days (3.2 mg/L). To evaluate the apoptotic activation of hPDCs on PFO-HPs and PBS-HPs in a hypoxic environment, caspase 3/7 activity was measured [Figure 3(B)]. It has been well established that an increase in caspase 3/7 activity leads to apoptosis.43,49 Apoptosis is a primary form of cell death under hypoxia.50 In the PFO-HP group, caspase 3/7 activity increased continuously and plateaued at day 10, but the activity differences at various time points during the first 5 days did not differ significantly, which indicated that the PFO-HPs supplied sufficient oxygen to prevent apoptosis for 5 days. However, in the PBS-HP group, caspase 3/7 activity increased dramatically at day 1,


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continued to increase gradually, and plateaued at day 5. Caspase 3/7 activity was significantly higher in the PBS-HP group than in the PFO-HP group for the first 7 days, but afterwards, caspase 3/7 activity did not differ significantly indicating that after 10 days of culture, the oxygen environment was similar for both the PFO-HPs and PBS-HPs due to depletion of the oxygen in the PFO emulsion. The higher cell proliferation and lower apoptotic activity of PFO-HPs at the initial stage compared with PBS-HPs suggested that no burst of oxygen release occurred causing hyperoxia, which induces cell growth arrest and apoptosis,51 or oxygen bubbles, which induces cell damage.52 To evaluate the oxygen responses of hPDCs on PFO-HPs and PBS-HPs, expression of the hypoxia-related biomarker HIF-1α during cell culture was measured using qRT-PCR [Figure 3(C)] and immunocytochemical analysis (Figure 4). HIF-1α expression was found to be reduced and delayed in PFO-HPs compared with that in PBS-HPs. The results were similar to the caspase 3/7 activity results. In addition, the viable cell distribution for each HP group was consistent with the CCK-8 results [Figure 3(A)]. These findings suggested that PFO-HPs provided a better oxygenated environment for hPDC survival compared with PBS-HPs with no oxygen carrier. Osteogenic Differentiation of hPDCs underwent Hypoxia. During tissue regeneration using tissue engineering techniques, cells introduced into 3D scaffolds inevitably experience hypoxic conditions until blood vessel ingrowth into the scaffold occurs. During this period, cell survival and maintenance of cell differentiation potency determine whether target tissue regeneration will succeed or fail. To investigate whether PFO-HPs provided a suitable environment for the maintenance of the osteogenic differentiation potency of hPDCs under hypoxia, osteogenic differentiation of hPDCs on PFO-HPs (pre-incubated under hypoxia for 10 days, then incubated under normoxia; Hyp-Nor/PFO-HP) was compared with osteogenic differentiation of normal


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hPDCs on PBS-HPs (incubated under normoxia; Nor/PBS-HP). In the Hyp-Nor/PFO-HP and Nor/PBS-HP groups, ALP activity increased gradually, plateaued at 2 weeks, and then declined (Figure 5 and Figure 6), indicating osteogenic differentiation of hPDCs.53 In addition, calcium deposition (mineralization) by hPDCs as they differentiated into osteoblasts increased continuously over time (Figure 5). The differences in ALP activity and calcium deposition between the Hyp-Nor/PFO-HP and Nor/PBS-HP groups were not significant, which indicated that the osteogenic differentiation potency of hPDCs on PFO-HPs did not change even after incubation under hypoxia for 10 days. To further evaluate the osteogenic differentiation potency of hPDCs in the Hyp-Nor/PFO-HP and Nor/PBS-HP groups, qRT-PCR was used to analyze expression levels of the osteogenic differentiation markers, Runx2 (early differentiation marker) and osteocalcin (late differentiation marker),54 after 0, 1, 2, and 4 weeks of culture. The expression levels of Runx2 and osteocalcin [Figure 6(A)] were consistent with the ALP activity and calcium deposition results. Immunocytochemical analysis and qRT-PCR data showed that Runx2 and osteocalcin expression levels did not differ significantly between the Hyp-Nor/PFO-HP and Nor/PBS-HP groups [Figure 6(B)], and the cells in both groups proliferated (as estimated by counting cell nuclei) during cell culture. Expression of the Runx2 (early differentiation marker) was brightest at 1 week then gradually weakened, whereas expression of the osteocalcin (late differentiation marker) was detected initially at 2 weeks and gradually increased up to 4 weeks suggesting osteogenic differentiation. These findings provided clear evidence that the osteogenic differentiation potency of the hPDCs on PFO-HPs that underwent hypoxia can be preserved. Therefore, we propose that these potent hPDCs on PFO-HPs may enhance the formation of new bone due to sufficient osteogenesis by hPDCs after blood vessel ingrowth [Figure 1(B)]. It has been established that new


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blood vessels take 1-2 weeks to infiltrate 3-mm-thick tissue-engineered 3D scaffolds at a maximum infiltration rate of ~400 μm/day.55 Based on our in vitro experiments and the literature, the minimum scaffold thickness for PFO-HPs to maintain the initial cell number and to preserve the osteogenic differentiation potency of hPDCs until blood vessel ingrowth occurs is ~8 mm [10 days x ~400 μm/day x 2 directions = ~8,000 μm (8 mm)]. Recent studies have shown that the thickness limits of tissues prepared using tissue engineering techniques is ~2 mm.16 Animal Study. To examine whether PFO-HPs can supply sufficient oxygen for the survival of hPDCs and the maintenance of osteogenic differentiation potency to facilitate new bone formation, miniature pigs with mandibular bone defects were used as animal models. After implantation of the specimens [hPDC-seeded PFO-HPs (hPDCs/PFO-HP), hPDC-seeded PBSHPs (hPDCs/PBS-HP), and PFO-HPs without hPDCs (PFO-HP)] into the bone defects, new bone formation was observed by plain radiography and CT at 0, 4, 7, and 10 weeks. Reconstructed bone was clearly observed in all of the defects [Figure 7(A-C)] and was confirmed by HU measurements [Figure 7(D) and Figure S3]. The continuous regeneration of bone indicated that PCL HPs embedded with PFO emulsion did not cause any unfavorable tissue reactions. The hPDCs/PBSHP group showed greater bone regeneration than the PFO-HP group without hPDCs, but the difference was not significant. The hPDCs/PFO-HP group formed new bone at a faster rate and with higher bone density at each time point than the hPDCs/PBS-HP and PFO-HP groups. At 10 weeks after implantation, newly regenerated bone was detected in the defects by H&E staining. The bony defects in all of the groups were almost completely filled with new bone; however, the bony defects in the hPDC/PFO-HP group displayed more effective bone reconstruction based on the extent of the Haversian system (osteon), which houses blood vessels and is the basic unit of bone [Figure 7(E)]. These findings support our hypothesis that PFO-HPs provide a suitable


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oxygenated environment for cell survival and maintenance of differentiation potency until blood vessel ingrowth occurs, and enhances bone target tissue regeneration via osteogenesis by the hPDCs on the PFO-HPs. In general, when bone is damaged, stem cells and progenitor cells are recruited to the damaged site from circulating blood, bone marrow, endosteum, and periosteum, and bioactive molecules are synthesized by the host and recruited cells such as platelet-derived growth factor (PDGF), bone morphogenetic proteins, FGF, and VEGF.56,57 Regeneration of bone with suitable volume and mechanical properties is precisely orchestrated, and it is well known that the delivery of exogenous cells, which can differentiate into bone cells, is a promising strategy to overcome the problem of insufficient numbers of cells for the generation of new bone tissue at defect sites. Nakahara et al.58 and Gerstenfeld et al.59 showed that the periosteum-derived cells used in this study are a better source of mesenchymal stem cells than bone marrow and that these periosteum-derived cells play pivotal roles in bone regeneration by differentiating into bone cells. To maximize the effect of exogenous cells on bone regeneration, it is essential to ensure cell survival and maintenance of cell properties, such as osteogenic differentiation potency, at defective regions without vascular networks until blood vessel ingrowth occurs. Thus, we propose that PFOHPs can provide sufficient oxygen to surrounding cells to prevent cell death and preserve cell properties under hypoxia, and thus are suitable tissue-engineering scaffolds for effective bone regeneration. To more clearly understand the improved effects of hPDCs adhered to PFO-HPs on bone regeneration, further osteogenesis and paracrine action studies on hPDCs in host tissues will be required.56,60

CONCLUSIONS


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In this study, we investigated whether PFO-HPs, which release oxygen, can provide a proper environment for hPDC survival and maintenance of osteogenic differentiation potency under hypoxia in vitro and for new bone formation at defective sites without a vascular network in vivo. Our PFO-HPs prolonged cell survival under hypoxia and preserved the osteogenic differentiation potency of hPDCs on PFO-HPs even after 10 days of hypoxia. In vivo, the hPDCs/PFO-HP group displayed a faster rate of new bone formation and a higher bone density than the hPDCs/PBS-HP and PFO-HP groups, which suggested that the PFO-HPs provided a suitable environment for cell survival and maintenance of differentiation potency until blood vessel ingrowth occurred, and thereby, enhanced osteogenesis and bone regeneration by hPDCs. Thus, PFO-HPs are a promising delivery system for various functional cells, including stem cells and progenitor cells, to produce clinically applicable tissues/organs.


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Figure 1. Schematic showing the expected events in the PFO-HP group during in vitro cell culture, which includes hypoxia followed by normoxia (A) and in vivo bone regeneration (B). Oxygen release from PFO-HPs allows for the survival and maintenance of the differentiation potency of cells during hypoxia leading to bone regeneration in bony defects.


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Figure 2. Scanning electron microscope (SEM) images showing the gross appearance and the surface/cross-sectional morphology of a PCL HP.


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Figure 3. Proliferation (A), caspase 3/7 activity (B), and HIF-1α expression (C) of hPDCs cultured on PBS-HPs and PFO-HPs under hypoxia (1% oxygen); n = 3; *p < 0.05, PBS-HP group vs. PFOHP group.


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Figure 4. Immunocytochemical analysis of hPDCs cultured on PBS-HPs and PFO-HPs under hypoxia; blue, cell nucleus; red, phalloidin; green, HIF-1α.


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Figure 5. Osteogenic differentiation of hPDCs underwent hypoxia for 10 days (on PFO-HPs) and normal hPDCs (on PBS-HPs). Quantitative analysis of ALP activity (A) and calcium deposition (B); n = 3; *p < 0.05.


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Figure 6. Expression of osteogenic biomarkers in hPDCs exposed to hypoxia for 10 days (on PFOHPs) and normal hPDCs (on PBS-HPs). (A) qRT-PCR expression analysis of Runx2 and osteocalcin; n = 3. (B) Immunocytochemical analysis of Runx2 (green) and osteocalcin (red) expression; blue, cell nuclei.


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Figure 7. Representative bone regeneration data from implantation of hPDCs/PFO-HP, hPDCs/PBS-HP, and PFO-HP into mandibular defects in miniature pigs. Radiographs (A), CT scans; dashed circle, initial defect or reconstructed bone (B), reconstructed 3D CT images (C), HU measurements; n = 3; *p < 0.05 (D), and histological sections; arrows, Haversian system; asterisks, PCL HPs (E) of the reconstructed bone at 10 weeks after implantation.


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ASSOCIATED CONTENT Supporting Information The supporting information includes the surgical procedure and measurements of HUs (PDF).

AUTHOR INFORMATION Corresponding Authors *E-mail: [email protected] (S.H. Oh) *E-mail: [email protected] (J.H. Byun)

Author Contributions The manuscript was written through contributions of all authors. All authors have given approval to the final version of the manuscript. 1 These authors contributed equally.

Notes The authors declare no competing financial interest or no conflict of interests.

Acknowledgments This research was supported by grants from the Korea Health Technology R&D Project through the Korea Health Industry Development Institute (KHIDI) funded by the Ministry of Health & Welfare, Korea (HI15C0607), and the Basic Science Research Program through the National Research Foundation of Korea (NRF) funded by the Ministry of Science, ICT and Future Planning (NRF-2015R1A5A2008833) and the Ministry of Education (2018R1D1A1A02085564).


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