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Keywords:

  • MOLECULAR PATHWAYS;
  • REMODELING;
  • BIOENGINEERING;
  • INJURY/FRACTURE HEALING;
  • THERAPEUTICS;
  • CHONDROCYTES;
  • CARTILAGE BIOLOGY

ABSTRACT

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Although bone has great capacity for repair, there are a number of clinical situations (fracture non-unions, spinal fusions, revision arthroplasty, segmental defects) in which auto- or allografts attempt to augment bone regeneration by promoting osteogenesis. Critical failures associated with current grafting therapies include osteonecrosis and limited integration between graft and host tissue. We speculated that the underlying problem with current bone grafting techniques is that they promote bone regeneration through direct osteogenesis. Here we hypothesized that using cartilage to promote endochondral bone regeneration would leverage normal developmental and repair sequences to produce a well-vascularized regenerate that integrates with the host tissue. In this study, we use a translational murine model of a segmental tibia defect to test the clinical utility of bone regeneration from a cartilage graft. We further test the mechanism by which cartilage promotes bone regeneration using in vivo lineage tracing and in vitro culture experiments. Our data show that cartilage grafts support regeneration of a vascularized and integrated bone tissue in vivo, and subsequently propose a translational tissue engineering platform using chondrogenesis of mesenchymal stem cells (MSCs). Interestingly, lineage tracing experiments show the regenerate was graft derived, suggesting transformation of the chondrocytes into bone. In vitro culture data show that cartilage explants mineralize with the addition of bone morphogenetic protein (BMP) or by exposure to human vascular endothelial cell (HUVEC)-conditioned medium, indicating that endothelial cells directly promote ossification. This study provides preclinical data for endochondral bone repair that has potential to significantly improve patient outcomes in a variety of musculoskeletal diseases and injuries. Further, in contrast to the dogmatic view that hypertrophic chondrocytes undergo apoptosis before bone formation, our data suggest cartilage can transform into bone by activating the pluripotent transcription factor Oct4A. Together these data represent a paradigm shift describing the mechanism of endochondral bone repair and open the door for novel regenerative strategies based on improved biology. © 2014 American Society for Bone and Mineral Research.


Introduction

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

There is a significant unmet clinical need to develop improved strategies for promoting vascularized bone regeneration. The Global Burden of Disease 2010 report recently published in The Lancet determined musculoskeletal diseases to be the second greatest cause of disability worldwide,[1] and in the United States the health-care cost burden associated with trauma has surpassed that of cardiovascular disease.[2] Bone grafting is an interventional technique used to restore structural integrity and promote bone repair in a diverse set of musculoskeletal conditions. Common indications for bone grafting include traumatic injuries sustained in motor vehicle accidents and war, fracture non-unions, spinal fusions, osteotomies from tumor resection, arthrodesis, and around joint replacements. Bone is the second most commonly transplanted tissue (behind blood) with an estimated 1.6 million grafting procedures performed annually.[3] The gold standard grafting technique remains transplantation of morsalized autologous bone. However, autologous grafting is associated with a number of clinical shortcomings, including limited availability of donor tissue, donor site morbidity, and increased potential for additional surgeries. Allografted bone, bone graft substitutes, and bone tissue engineering represent techniques to address inherent limitations of autografts. Despite recent progress in these technologies, poor graft integration and osteonecrosis remain problematic,[4] and clinical failure associated with bone allografts are estimated to be between 16% and 35%.[5]

In this study, we explore the idea of using cartilage tissue as a method to stimulate bone repair through endochondral ossification. We speculated that an underlying problem with current bone grafting technologies is that they promote bone regeneration through direct osteogenesis. However, long bone formation and fracture healing typically proceed via a cartilage intermediate, suggesting that an endochondral tissue therapy may be a more clinically and biologically appropriate treatment. Cartilage is physiologically adapted to function well under avascular conditions, and hypertrophic chondrocytes within endochondral cartilage stimulate vasculogenesis. These inherent biological features of cartilage propound that it may be a better therapeutic strategy for bone regeneration. We hypothesized that by promoting endochondral ossification using cartilage tissue grafts we could address the clinical shortcomings of current technologies to produce a well-vascularized bone regenerate that integrates with the host tissue.

The molecular details of endochondral ossification are understood predominantly in the context of long bone development.[6] First, mesenchymal cells condense and undergo chondrogenesis to form a cartilaginous model of the future bone. Chondrocytes within this anlagen organize into morphologically and functionally distinct domains that correspond to their maturation state (resting, proliferating, hypertrophic) and allow for longitudinal growth. Hypertrophic chondrocytes are believed to be the terminal maturation state of the chondrocyte. Upon maturation, chondrocytes undergo apoptosis, the cartilage matrix is degraded and replaced by the marrow cavity, and bone forms in the adjacent perichondrium to form the diaphysis of the long bones.[7] Hypertrophic chondrocytes are distinguished by their enlarged round shape and expression of type X collagen, matrix metalloproteinase-13 (MMP-13), and vascular endothelial growth factor (VEGF). These proteins are essential in recruiting osteoblast precursors from the surrounding tissue, promoting vascularization, and mineralizing of the extracellular matrix. Repair of nonstabilized fractures also occurs through a cartilage intermediate,[8] in a process that is presumed to follow endochondral ossification during skeletogenesis. In a murine tibia fracture, an avascular cartilage callus (∼day 5) precedes vascular invasion, mineralization, and remodeling into bone (∼days 10–28).

Despite many similarities between endochondral ossification during development and repair, the therapeutic application of cartilage tissue for promoting bone regeneration has not been established experimentally. The overall goal of this study was to utilize a translational murine model of segmental bone defects to validate cartilage grafts as a potential treatment strategy for bone repair. Further, by studying the molecular details during healing, we aimed to understand how the cartilage graft (donor tissue) and host tissue contribute to the repair process. An accurate assessment of how cartilage grafts facilitate bone regeneration is a necessary foundation for building an effective clinical strategy to promote endochondral repair. Design of scaffold material, cell/tissue source, and bioactive reagents will all depend on the fate of the chondrocytes, source of osteoblasts, and role of the vasculature during the repair process.

The data presented in this study demonstrate that cartilage grafts produce a well-vascularized and integrated bone regenerate in murine segmental tibial defects, implying that cartilage may be a better therapeutic effector for modulating bone regeneration than bone itself. Further, genetic labeling of the donor and host tissue revealed that the bone regenerate is donor/cartilage derived, suggesting that chondrocytes may directly transform into osteoblasts. This mechanism of repair significantly contrasts endochondral ossification during long bone development, requiring that we reevaluate the mechanisms of endochondral repair to improve bone regeneration strategies.

Materials and Methods

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Fractures

All animal studies were approved by the UCSF IACUC. Adult, male mice (10 to 14 weeks old) were anesthetized, and a standardized, closed non-stable fracture was made in the mid-diaphysis of the tibia using a custom-built apparatus designed to deliver a reproducible three-point bending fracture by controlling the weight (460 g) and distance (14 cm) of the force. Fractures were not stabilized to promote endochondral repair, and animals were provided with analgesics as needed.

Tissue grafts into the tibia defect model

A 2-mm segmental defect was created by an osteotomy in the mid-tibial diaphysis of immunocompromised mice (Nude Mice, Nu/J #2019; The Jackson Laboratories, Sacramento, CA, USA) (Fig. 1C). One of four different tissue grafts was transplanted into the defect: cartilage grafts, living cortical bone grafts (“isograft”), devitalized cortical bone grafts (“allografts”), or stem cell–derived cartilage pellets (see below). As a control, the critical-sized defect remained empty. Cartilage grafts were isolated from the central portion of a day 7 fracture callus (Fig. 1B) ex vivo by using a microscope to dissect out the cartilage and remove all noncartilaginous adherent tissues and the perichondrium. Isografts were created by simply replacing the osteotomized bone into the defect, and allografts were created by washing osteotomized bone in 70% EtOH and freezing at −80°C as previously described.[9-11] An 8.0 suture was used to secure the graft in place by closing the muscle. Tibias were externally stabilized with a customized circular fixator consisting of two 2-cm circular rings held concentrically by three threaded rods (Fig. 1C). This device provides rigid fixation as described previously.[12, 13] Animals survived for 1 to 6 weeks, with a minimum of 5 animals analyzed histologically at each time point and 8 animals analyzed at 4 weeks by micro-computed tomography (µCT) and biomechanical testing.

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Figure 1. Cartilage grafts from fracture callus. (A) X-ray image of unstabilized mid-diaphysial tibia fracture (arrow) created by three-point bending. (B) Safranin-O staining of fracture callus 7 days after injury. Grafts were isolated ex vivo; boxed region represents the approximate area from where grafts were harvested. (C) Grafts were transplanted into 2-mm segmental defects created in the mid-diaphysis of externally stabilized murine tibia. (D) Safranin-O staining of the cartilage graft (top), followed by DIG-probe in situ hybridization for sox-9, col2, col10, and osteocalcin (oc). (E) Quantitative RT-PCR shows gene expression in the cartilage graft (n = 6) relative to the whole fracture callus (n = 5). Data represent mean ± 95% confidence, with significance (*) of p < 0.005.

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µCT and biomechanical testing

A Scanco Medical AG (Bruttisellen, Switzerland) µCT was used to scan both the grafting area and fracture callus. Samples were rotated through 360° and the X-ray settings were standardized to 70 kV and 114 µA, with an exposure time of 0.14 seconds per frame to yield a nominal resolution of 10.5 μm. A 0.5-mm-thick aluminum filter was employed to minimize beam-hardening artifacts. Scan time for each sample was approximately 50 minutes. Bone mineral density was analyzed from 200 slices within the integration site or fracture callus, using a custom-made script. Briefly, bone mineral density (BMD) was measured by normalizing mineral content from the X-ray attenuation by bone volume. Integration was scored (0 = no integration; 1 = integration) based on µCT images to indicate incidence of integration. After scanning, the tibias were subjected to three-point bending using an ElectroForce 3200 testing machine (Bose Corp., Eden Prairie, MN, USA) to measure the integration strength. Only grafts that had integrated both proximally and distally were tested mechanically. Tibiae were placed on their lateral surface on the lower supports of the bending jig with supports located under the tibia-fibula junction and the tibial crest. A preload of 1 N was applied from above at the midpoint between the two lower supports to stabilize the bone. The load applied to the bone was measured by a 450-N load cell at a displacement rate of 2 mm/min. A load-displacement curve was generated for each bone and used to determine ultimate load. Statistical differences for integration success was tested using a pairwise comparison between cartilage, isograft, and allograft using the Fisher exact test. BMD and ultimate failure data were compared using an ANOVA followed by Tukey honestly significant difference (HSD) pairwise comparison. Any p values < 0.05 were considered significant.

Bone tissue embedding and histology

Tibiae from euthanized mice or in vitro cartilage grafts were collected and fixed in freshly made 4% paraformaldehyde (PFA, pH 7.2 to 7.4) for 24 hours at 4°C. Tibias were decalcified in 19% EDTA (pH 7.4) for 14 days at 4°C and then processed for paraffin or frozen histology. Histology staining to visualize bone and cartilage tissues was completed: Modified Milligan's Trichrome (bone = blue), Safranin-O/fast green (cartilage = red), or Hall and Brunt Quadruple stain (HBQ, bone = red, cartilage = blue).

Immunohistochemistry on bone tissue

X-gal staining was used to detect β-galactosidase on frozen sections that were post-fixed in 0.2% glutaraldehyde for 15 minutes, and then exposed to X-gal staining solution overnight at 37°C. Immunohistochemistry (IHC) for eGFP (A10259, Invitrogen, Carlsbad, CA, USA) was performed by treating sections with 0.3% Triton X (15 minutes), 0.05% trypsin (20 minutes, 37°C), 10 mM sodium citrate buffer (30 minutes, 100°C), 3% H2O2 in methanol (30 minutes), and blocking with 5% bovine serum albumin (BSA, 1 to 2 hours). Primary antibody was applied at a 1:100 dilution in 1% BSA (overnight, 4°C), samples were washed and then reacted the species specific secondary antibody (Santa Cruz Biotechnology, Santa Cruz, CA, USA), and detected by VectainStain ABC Kit (#PK-4000, Vector Laboratories, Burlingame, CA, USA) followed by 3, 3'-diaminobenzidine (DAB) with nickel ammonium sulfate and colbalt chloride. Antibodies to osteocalcin (#M173, Takara Bio, Otsu, Japan; 1:200) and Oct4A (#SC5279 Santa Cruz; 1:100) were applied to paraffin sections using the same protocol as eGFP with changes to antigen retrieval: osteocalcin, trypsin only; Oct4A, 40 minutes in EDTA (pH = 9.0, 1 mM) at 98°C.

In situ hybridization

In situ hybridization was performed on paraffin wax-embedded sections as previously described.[14, 15] Subclones of mouse sox-9, type II collagen (col2a1), type X collagen (col10a1), and osteocalcin (oc/bglp) were linearized for transcription of DIG-labeled antisense riboprobes.

In situ cell death detection

Detection of apoptosis using the Roche In Situ Cell Death Detection Kit (#116847959, Roche, Mannheim, Germany) according to the manufacturer's protocol. Sections were deparaffinized, treated with proteinase-K (20 μg/mL in 10 mM Tris HCl, 15 minutes), then reacted with the kit for 1 hour at 37°C in the dark. Positive controls were treated with DNase I before the TUNEL reaction, whereas negative controls were not reacted with vial #1 from the kit. Slides were mounted with VectaShield with Dapi (#H-1200, Vector). Apoptosis was visualized using a fluorescent microscope and photos merged with Safranin-O images from adjacent slides.

Expansion of human mesenchymal stem cells

Human mesenchymal stem cells (hMSCs) were isolated and expanded from iliac crest bone marrow aspirates of consented donors as previously described[16] or purchased from Lonza (Allendale, NJ, USA; #PT-2501). hMSCs were expanded in monolayer using DMEM supplemented with 10% FBS and fibroblast growth factor 2 (FGF-2, 10 ng/mL); cells were used after 3 to 12 population doublings (passage 1 to 4).

hMSC pellet culture and in vivo transplantation

hMSC-derived cartilage pellets were formed by putting 200,000 hMSCs into v-bottomed suspension plates, centrifuging for 5 minutes at 800g and culturing in a chondrogenic medium containing dexamethasone and TGF-β1 as previously described.[16] Pellets were cultured for 3 weeks at 37°C with 5% CO2 and either harvested for histology or transplanted into the tibia defect model (2 to 3 pellets per defect to create a tight fit).

Isolation and culture of human articular chondrocytes

Human articular chondrocytes (hACs) were obtained from the discard tissue of fresh osteochondral allografts (The Joint Restoration Foundation: male, 12 to 25 years, 6 donors). Chondrocytes were isolated by digesting cartilage tissue in 1% pronase (37°C, 1 hour), followed by 1 to 3 hours in a 0.4% collagenase type II solution (LS004176, Worthington Biochemical, Lakewood, NJ, USA). Cells were isolated from digested tissue using a 70-μM filter, spun down to remove collagenase solution, and directly encapsulated into scaffolds without expansion.

Poly(ethylene glycol) diacrylate scaffolds

Primary chondrocytes or expanded hMSC were photoencapsulated in a poly(ethylene glycol) diacrylate (PEGDA)-based semi-interpenetrating network with a scaffold composition consisting of 16% (w/v) PEGDA (6.0 kDa) and 32% (w/v) PEG-n-dimethyl ether (PEG, n = 2000, MW 88 kDa) to achieve a final cell density of 25 × 106 cells/mL as previously described.[17, 18] Hydrogels were cultured at 37°C, 5% CO2 for 6 weeks in defined chondrogenic medium.

mRNA isolation and quantitative RT-PCR

Murine mRNA was isolated from cartilage grafts or the entire fracture callus by homogenization in TRIzol. cDNA was reverse transcribed with Superscript III (#18080, Invitrogen) and RT-PCR was performed using SYBR Green. The primer sequences are included in Table 1. Relative gene expression was calculated by normalizing to the housekeeping gene (GAPDH, ΔCT) and then to gene expression of the entire fracture callus (ΔΔCT). Fold change was calculated as 2−ΔΔCT. Graphs represent mean ± 95% confidence of biological replicates for the cartilage grafts (n = 6) or the entire fracture callus (n = 5). mRNA isolation and cDNA synthesis from the hydrogels was performed as previously described.[19] Relative change in gene expression was calculated by normalizing to the housekeeping gene (18S, ΔCT) and then to gene expression of freshly isolated hACs or undifferentiated hMSCs before encapsulation (ΔΔCT). Fold change was calculated as 2−ΔΔCT. Graphs represent mean ± 95% confidence for a total of 6 different hAC and hMSC donors.

Table 1. Primer List
GeneSpeciesSyber GreenTaqman Assay ID#
ForwardReverse
GAPDHMouseTGATGACATCAAGAAGGTGGTGAAGCCTTGGAGGCCATGTAGGCCAT
Sox9MouseGCAGACCAGTACCCGCATCTCTCGTTCAGCAGCCTCCAG
Col2AMouseGGCTCCCAGAACATCACCTATCGGCCCTCATCTCTACATC
Col10AMouseAATCTGAAATGCAAGGTGCTAAGACTCAAATAGTCATTAAAGCAAA
Col1AMouseCCCAGAACATCACCTATCACTTGGTCACGTTCAGTTGGTC
OsteocalcinMouseCGCTCTGTCTCTCTGACCTCTCACAAGCAGGGTTAAGCTC
18SHumanhs9999999901_s1
ACANHumanhs00153936_m1
Col2AHumanhs00264051_m1
Col10AHumanhs00166657_m1
Col1AHumanhs00164004_m1
MMP-13Humanhs00233992_m1

MSC pellet/hydrogel histology and immunohistochemistry

Hydrogels were fixed in 10% neutral-buffered formalin, embedded in paraffin, and 5-μm sections cut onto silane-coated slides. Toluidine blue staining was used to visualize sulfated proteoglycans. IHC was used to detect collagens I (1:400, Anthony Hollander), II (1:200, II-II6B3, Developmental Studies Hybridoma Bank, Iowa City, IA, USA), X (1:200, Gary Gibson) as previously described.[17, 19, 20] Human mitochondria (1:100, #MAB1273, Millipore, Billerica, MA, USA) detection followed the same protocol, but slides were treated with 0.05% trypsin (20 minutes at 37°C) for antigen retrieval.

Osteoconductive in vitro cartilage explant culture

Cartilage from the fracture callus was harvested from the central portion of the day 7 fracture as described above, rinsed with sterile PBS, and then transferred into medium for in vitro culture at 37°C and 5% CO2. Control medium was a serum-free high-glucose DMEM containing 1% penicillin-streptomycin, 1% ITS+ Premix (#354352, BD Biosciences, San Jose, CA, USA), 1 mM sodium pyruvate, and 100 ng/mL ascorbate-2-phosphate. For osteogenic cultures, samples were grown under control conditions for 2 weeks, then the medium was supplemented with bone morphogenetic protein 2 (BMP-2: 10 nm, 100 nm, or 1 μm) and 10 mM β-glycerol phosphate (βGP). hMSC-derived cartilage pellets transferred to osteogenic conditions completed 3 weeks of standard chondrogenic culture before being transferred to osteogenic media for an additional 4 weeks of culture. Human vascular endothelial cell–conditioned media (HUVEC-CM) was collected from confluent plates of passage 2 to 3 HUVEC cells grown in Gibco Medium 200 with LVES supplement (#A1460901, Invitrogen). Media was collected every 3 days; batches were combined and filter sterilized.

Histomorphometric analysis

Area of mineralization or proteoglycan deposition within the cartilage explants was determined by staining every 25th section with Alizarin Red or Safranin-O. Images were captured from a Leica (Buffalo Grove, IL, USA) DM 5000 B light microscope, and number of pixels was determined by Adobe Photoshop (Adobe Systems, San Jose, CA, USA). The volume of each was calculated using the equation for a conical frustum:

  • display math

Ai and Ai + 1 are the area of total explant or mineralized explant extracellular matrix in sequential sections; h is the distance between sections (250 µm), and n is total number of sections analyzed per cartilage explant. Data represent the mean ± 95% confidence of the samples, and significance was tested using the Mann-Whitney U-test for nonparametric data, with p values < 0.05 determined to be significantly different. Biological replicates (n = 3–6) were used for in vitro quantifications.

Results

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Cartilage grafts produce well-vascularized and integrated bone regenerate

The ability of a cartilage graft to stimulate bone regeneration was tested in a murine model of a segmental bone defect. External stabilization was applied to the intact tibia as previously described,[12, 13] then a 2-mm osteotomy was created in the mid-diaphysis and cleared of bone debris in preparation for the graft (Fig. 1C). Cartilage for the graft was isolated from the central portion of a day 7 nonstabilized fracture callus by carefully dissecting the fracture callus ex vivo using a dissecting microscope (Fig. 1A, B). The phenotype of the cartilage graft was characterized using histology, in situ hybridization, and quantitative RT-PCR (Fig. 1). Safranin-O staining shows strong proteoglycan deposition in the graft and highlights that cellular morphology is predominantly that of normal chondrocytes with some hypertrophic chondrocytes in the middle (Fig. 1D). In situ hybridization confirms expression of sox-9 and col2a1 throughout the graft and col10a1 expression in hypertrophic chondrocytes in the central portion of the graft. Osteocalcin (oc/bglap) was not detected anywhere in the graft. Importantly, in regions that had low levels of Safranin-O staining, we observed cells expressing sox-9 but not oc/bglap (Fig. 1D and Supplemental Figs. S1 and S2). Further, we detected no evidence of a progenitor population using an antibody to Oct4A (Supplemental Fig. S3A, B). Quantitative RT-PCR analysis shows significantly more sox-9 and col2a1 and significantly less oc/bglap expression in the cartilage tissue graft compared with the entire fracture callus (Fig. 1E).

A 4-week healing time course of the defect shows that cartilage grafts facilitate endochondral bone repair in the mouse tibia (Fig. 2). Further, the trabeculated morphology of the bone at day 28 suggests the regenerate is highly vascularized (Fig. 2C). In size-matched empty defects, complete bridging of the bone was never observed (0/4); rather, defects were filled largely with granulation tissue (Supplemental Fig. S4). Empty defects display new bone regeneration predominantly at the periosteum with only small amounts of new bone in the osteotomized region.

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Figure 2. Cartilage grafts produce an integrated and vascularized bone regenerate in a segmental bone defect. (A–C) Safranin-O or (E–G) Masson's Trichrome staining at days (A, E) 7, (B, F) 14, or (C, G) 28 after implantation of the cartilage graft. (D) µCT image of tibia defect 4 weeks postsurgically. cb = cortical bone (host); graft = transplanted fracture callus cartilage.

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Integration between the cartilage-derived bone regenerate was evident histologically and tomographically (Fig. 2C, D). When the healing produced from cartilage grafts was compared with living cortical bone grafts (isograft) and devitalized cortical bone grafts (allografts), cartilage was as likely to integrate with the host bone as isograft, and both were far superior to allograft (Fig. 3A, B; a = p < 0.005). Integration strength, measured as ultimate failure by three-point bending, confirmed integration quality was not significantly different between cartilage and isograft, whereas integration was so poor with allograft that all grafts failed immediately without measurable force (Fig. 3C). Integration strength between the host and graft at 4 weeks did not reach the full strength of uninjured cortical bone or the repair of a day 14 nonstabilized fracture that is strengthened by the excess callus tissue (Fig. 3C; b = p < 0.005, c = p < 0.05). However, BMD of the isografts and cartilage grafts was not statistically different from the cortical bone controls, and, importantly, the cartilage graft had a BMD equivalent to that of a day 28 fracture callus (Fig. 3D; b = p < 0.02). BMD of the allograft was slightly higher than that of the days 14 and 21 fracture callus, and even cortical bone; increased BMD is seen in devitalized bone because it does not have the ability to remodel (Fig. 3D; b–d = p < 0.01).

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Figure 3. Integration of cartilage grafts is equivalent to isograft and superior to allograft after 4 weeks of healing. (A) Overall percentage that cartilage, isograft, or allograft integrated with the host tissue based on two potential integration sites per graft (7 to 8 animals, 14 to 16 integration opportunities; a = statistically different from cartilage and isograft, p < 0.005). (B) Graphical representation of integration for cartilage, isograft, and allograft. (C) Ultimate failure of grafts by three-point bending. Ultimate failure of uninjured cortical bone for both the grafts (eGFP) and host mice (immunocompromised/nude) are provided as a positive control alongside failure of nonstabilized fractures for reference (b = statistically different from d14 fx; c = different from cortical bone, p < 0.05). (D) Bone mineral density (BMD) of the fracture callus, grafts, and cortical bone (d = statistically different from d21 fx, p < 0.02).

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Bone regenerate is donor-derived

Based on the healing process observed above in the segmental defect and our previous studies of endochondral fracture repair, we presumed bone regeneration from the cartilage grafts occurred through endochondral ossification in a process analogous to embryonic long bone development. This model predicts the hypertrophic chondrocytes undergo apoptosis and are replaced by osteoblasts from the host. We tested this model by transplanting cartilage from LacZ+/+ Rosa26 mice into immunocompromised hosts and followed the fate of the regenerate using X-gal staining to detect β-galactasidase activity (x-gal background and staining specificity, Supplemental Fig. S5). In contrast to the established model, we found that the majority of the bone regenerate was donor, rather than host, derived (Fig. 4). Magnified views of the graft after 4 weeks of healing illustrate transition zones where there appears to be a morphological transition of the cells from chondrocytes (Fig. 4D, yellow arrow) to osteocytes (Fig. 4D, yellow star) within the donor tissue. Seamless integration is observed between the graft and host tissue, with donor and host cells embedded within a continuous matrix (Fig. 4C, F). Host-derived chondrocytes are also evident in the tissue at the interface between the graft and host tissue (Fig. 4D, green triangle), suggesting the host tissue was stimulated by bioactivity of the graft tissue or by mechanical stresses at the tissue interface.

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Figure 4. Donor (blue) versus host contribution to the bone regenerate. Cartilage grafts were obtained from Lac-Z+/+-Rosa26 reporter mice and transplanted into immunocompromised mice (SCID Beige) mice. Donor cells were labeled using X-gal staining to detect β-galactosidase activity from the Rosa26 mice at (A) 7, (B–E) 28, or (F–H) 42 days after cartilage engraftment. Special characters (D) donor osteocytes (yellow star), donor chondrocytes (yellow arrow), host chondrocytes (green triangle). cb = cortical bone (host); graft = transplanted fracture callus cartilage from LacZ+/+ mouse.

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The donor origin of the bone regenerate was confirmed using cartilage grafts obtained from eGFP mice and transplanted into immunocompromised mice in the same manner described above (Supplemental Fig. S6). Immunohistochemistry to GFP demonstrated that the regenerate remained donor derived after 28 days of healing, was highly trabeculated, and integrated with the host.

Cartilage grafts from human MSCs also promote bone regeneration by tissue transformation

To validate the use of cartilage grafts for endochondral bone repair in a more translationally relevant system, we generated cartilage pellets from human mesenchymal stem cells (hMSCs) and implanted these pellets into the murine segmental defect (Fig. 5). Chondrogenesis was induced using standard pellet cultures supplemented with TGF-β1 and dexamethasone.[21] Histological and immunohistochemical analysis of the hMSC-derived cartilage pellets at the time of implantation confirmed the tissue was strongly chondrogenic (Fig. 5A, B) with regions of hypertrophy marked both by morphology and collagen X protein (Fig. 5C). After 4 weeks in the tibial defects, transformation of the cartilage pellets into vascularized bone tissue was visualized with Masson's trichrome (Fig. 5D, E). Donor origin of the bone was confirmed with an antibody to human mitochondria (Fig. 5F). In the tibia, the pellet retains its rounded shape and integrates with the new periosteal bone formed at the cortex of the host bone.

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Figure 5. Human MSC-derived cartilage pellets transplanted into segmental bone defect regenerate bone. (A–C) hMSC pellets after 3 weeks of in vitro culture in chondrogenic conditions stained with (A) Safranin-O, or antibodies to (B) collagen II and (C) collagen X. (D, E) Masson's Trichrome staining of segmental defect 4 weeks after transplantation of hMSC-derived cartilage pellets. (F) hMSC-dervied pellet with antibody staining for human mitochondria in the trichrome positive bone.

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Tissue engineering cartilage grafts

Chondrogenic pellets grown by the method used in this study are inherently limited in size to approximately 1 mm in diameter, and we found that although the pellets integrated with the bone defect (Fig. 5E), multiple adjacent pellets do not integrate with each other (not shown). Consequently, scaling this methodology to larger-scale defects will require a different technology, such as the scaffold-based technologies developed for tissue engineering. We have previously developed a biocompatible technique for photoencapsulating hMSCs into synthetic PEGDA-based scaffolds that allow for chondrogenesis.[17, 19, 20] Using this scaffold technology, we tested the phenotype of cartilage that would develop from two candidate cell sources for tissue-engineered cartilage: hMSCs or healthy human chondrocytes (hACs). After 6 weeks of in vitro culture, sulfated proteoglycans and type II collagen were present in both hMSCs and hACs constructs, indicating their fundamental ability to generate cartilage tissue (Fig. 6). However, only hMSC-encapsulated constructs elaborate collagen I and X matrix proteins and show a robust increase in COL10A1 and MMP-13 gene expression.

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Figure 6. Engineering endochondral cartilage for tissue engineering. Human MSCs (A) or healthy human articular chondrocytes (hACs) (B) were photoencapsulated into PEGDA-based scaffolds and cultured in vitro with chondrogenic medium for 6 weeks. (C) Antibody specificity was verified using articular cartilage and the osteochondral interface. (D) Gene expression of the hMSC (n = 6) or hAC (n = 6) derived tissue engineered cartilage after 6 weeks of in vitro culture was compared with gene expression of each tissue at day 0. Graph represents mean ± 95% confidence.

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BMP and HUVEC-conditioned medium stimulate mineralization in vitro

To better understand the role of the vasculature in converting cartilage to bone in vivo, cartilage tissue grafts from the fracture callus were cultured in vitro under osteoinductive conditions. Fracture callus cartilage cultured in vitro without BMP-2 maintained strong proteoglycan staining, and the cells became hypertrophic with no evidence of mineralization in the matrix (Fig. 7A–C, M). When cultured with BMP-2 and β-glycerolphospate (βGP), the fracture callus cartilage mineralized, but cells maintained the hypertrophic chondrocyte morphology as opposed to becoming osteocyte-like (Fig. 7D–F, M). Increasing BMP-2 concentration in culture increased mineral content but did not change tissue morphology (Supplemental Fig. S7). Similar results were obtained when MSC-derived cartilage pellets were cultured in vitro with BMP-2/βGP (Supplemental Fig. S8).

image

Figure 7. HUVEC-conditioned medium is sufficient for producing mineralized cartilage in vitro. Alizarin red and Safranin-O/fast green staining of cartilage was obtained from the day 7 fracture callus and cultured in vitro in basal medium containing 10 nM dexamethasone and 100 µg/mL A2P for 2 weeks, then either maintained in that basal condition (A–C) or transferred to osteogenic supplements (rhBMP2,βGP,A2P) (D–F), HUVEC-CM (G–I), or HUVEC-CM with osteogenic supplements (J–L). Alizarin red and Safranin-O/fast green staining of cartilage grafts (M). Quantification of the percentage of mineralization (gray) and proteoglycan (black) in cartilage explants after 4 weeks of in vitro culture. Graph represents mean ± 95% confidence, with significance (*) of p < 0.005.

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Given the incomplete morphological transformation of the explanted cartilage to bone, we hypothesized that biological stimuli from the vascular endothelial cells may be essential. To test this hypothesis, we generated HUVEC-conditioned media to simulate the fracture callus microenvironment and added it to the fracture callus cartilage cultured in vitro, either with or without the addition of BMP-2/βGP. HUVEC-conditioned medium alone was sufficient to produce a mineralized matrix content in the explants that was not statistically different from osteogenic medium (Fig. 7G–I, M). Further addition of BMP-2/βGP to the HUVEC-conditioned medium produced a small, but not statistically significant, increase in the amount of mineralization in the cartilage explants (Fig. 7J–M). Explants cultured with HUVEC-conditioned media retained their hypertrophic morphology, perhaps because of an increase in the proteoglycan-rich cartilage matrix in the explant (Fig. 7M).

Endochondral fracture repair

Morphological conversion of chondrocytes into osteocytes was not completely recapitulated in vitro, suggesting the simple replacement of soluble bioactive molecules was not sufficient to support transformation. In vivo, the morphological transition from chondrocyte to osteocyte is clearly seen in the fracture callus in regions surrounding the invading vasculature (Fig. 8A–D). Hypertrophic chondrocytes in these transition areas stain for both the dense collagen fibers of bone matrix (HBQ red, Fig. 8A–D) and osteocalcin protein (Fig. 8E, F). Adjacent to the hypertrophic chondrocytes, cells appeared flatter and have the appearance of an osteocyte. To test if the switch in cell morphology was the result of apoptosis of hypertrophic chondrocyte and replacement by cells from the vasculature as suggested by current models, we used an In Situ Cell Death Kit. At the cartilage to bone transition region at day 7 (Fig. 8G) and day 10 (Fig. 8H) the majority of the apoptosis occurred in cells that histologically and morphologically identified as bone cells, as opposed to hypertrophic chondrocytes. These data suggest that apoptosis may simply facilitate remodeling of the hard callus to create marrow space. We next looked to see if progenitor programs were being activated in hypertrophic chondrocytes at cartilage to bone transition areas. Interstingly, we find Oct4A is expressed only in the hypertrophic chondrocytes near the invading vasculature (Fig. 8I, J and Supplemental Fig. S3C, D). We did not see evidence of Oct4A in chondrocytes or hypotrophic chondrocytes in areas without invading vasculature (Supplemental Fig. S3E, F) or in osteocytes in newly formed bone (Supplemental Fig. S3G, H).

image

Figure 8. Cartilage to bone transition zone in fracture callus. (A–D) Hall and Brunt Quadruple stain (HBQ, bone = red, cartilage = blue) of nonstabilized fracture callus 10 days post-injury. (E, F) Osteocalcin immunohistochemistry of nonstabilized fracture callus 10 days post-injury. (G, H) In situ cell death detection (GFP) merged with Safranin-O (cartilage = red, bone = blue) staining of day 7 (G) or day 10 (H) fracture callus. (I, J) Oct4A immunohistochemistry of day 10 fracture callus. Arrows point to Oct4A-positive hypertrophic chondrocytes. BV = blood vessel or bone marrow space.

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Discussion

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

The ability to regenerate a vascularized bone using tissue grafts is presently constrained by the clinical limitations of current technologies. Although autograft bone procedures produce reasonably successful outcomes, donor site morbidity and quantity of available tissue are significant problems.[22, 23] Clinical failures associated with allografts have been extensively documented in both animal and human studies. The main problem is that the tissue remains inert, producing limited revitalization and eventual loss in structural integrity of the graft.[5, 10, 24-26] A major effort has been made to develop novel technologies to promote vascularization during bone regeneration using gene therapy or growth factor delivery of VEGF.[11, 27-30] In this study, we investigated an alternative approach for stimulating vascularized bone regeneration using cartilage tissue grafts to promote endochondral bone repair. Using a translational murine model of segmental bone defects, we demonstrated that transplanted cartilage produces a highly trabeculated and well-vascularized bone regenerate that integrates well with the host tissue.

The specific advantage that cartilage grafts offer over bone transplantation is likely associated with the inherent bioactivity of chondrocytes and their adaptation for function under avascular conditions. As chondrocytes mature toward hypertrophy, they secrete VEGF to stimulate vascular invasion, MMP-13 to promote mineralization of the extracellular matrix, and BMPs to promote osteogenesis.[31-34] Capitalizing on this innate biological function may eliminate the need for complex engineering strategies to deliver bioactive stimuli. Further, chondrocytes are metabolically optimized for avascular conditions,[35] providing a therapeutic advantage for bone repair because grafting is often required in areas of vascular damage. Ischemia is detrimental to osteogenesis and significantly increases the rate of fracture non-union[36, 37] and contributes to current bone graft failures.[4]

The concept of using cartilage to promote endochondral bone regeneration has been effectively overlooked at a clinical level despite the clear biological relevance of this approach. In this study, we generated endochondral cartilage by creating unstabilized fractures in murine tibias and by inducing hMSCs toward chondrogenesis. Previous experiments have demonstrated the capacity of cartilage with a hypertrophic phenotype to form bone both in vitro[38-41] and after ectopic transplantation.[41-45] Interestingly, hyaline cartilage, which does not express collagens I or X, appears to be privileged from this transition to bone, and similar experiments have been unable to induce these “permanent” chondrocytes to undergo endochondral ossification.[[41, 43, 45, 46], and data not shown] Consequently, endochondral cartilage, or specifically the ability to undergo hypertrophic maturation, appears requisite in supporting bone regeneration. Functional bone organ regeneration by endochondral ossification has demonstrated that complete bone tissue can be produced from hMSC-derived cartilage,[47, 48] validating the concept explored here that recapitulating developmental sequences may be a useful tissue engineering strategy. Our data expand upon these studies to show that endochondral cartilage can promote effective healing of a segmental bone defect in a clinically relevant model. Specifically, the bone regenerate produced by the cartilage graft is highly vascularized and integrates much better than allograft bone and as well as isograft. The isograft and allograft models used for this study have been previously described in murine long bone defect studies,[9-11] provided for consistent graft dimensions for comparison across grafting groups, and have clinical relevance for the treatment of human bone defects with large segment grafting.

A key feature of our study is that we utilized genetic labels on the cartilage grafts to track the contribution of the donor versus host to the bone regenerate. Our hypothesis was that the cartilage graft would create a template for bone remodeling in a similar fashion to endochondral ossification during development. If healing occurred by this model, the transplanted chondrocytes would undergo apoptosis and the bone would be host derived. In contrast, we found the bone regenerate was largely donor derived. Our data were verified using two different genetically labeled mouse strains, Rosa26 (Lac-Z-positive) and eGFP, and a human mitochondria stain to identify origin of the transplanted hMSC-derived cartilage pellets. In line with our transplant results, both Scotti and colleagues[44] and Sacchetti and colleagues[49] have found that the ectopic bone formed from subcutaneously transplanted hMSC cartilage was donor derived. However, because of the potential heterogeneity in the cartilage tissue grafts, neither our study nor the previous works can conclusively refute the argument that the grafts contained committed osteocytes, or that a small population of donor-derived osteoprogenitor cells could repopulate the bone regenerate. To address the possibility that osteoblasts were present in the cartilage graft, we used qRT-PCR and in situ hybridization to confirm sox-9 expression throughout the graft and demonstrated that there was no molecular evidence of osteoblasts. Further, we saw no expression of the stem cell marker Oct4A in the cartilage graft before transplantation, suggesting progenitor cells were not present. Oct4A is an established marker of pluripotent stem cells, although its role in adult stem cell populations is not clear. Presently, there are no good markers for mouse MSCs and the absence of Oct4A cannot conclusively demonstrate osteoprogenitors were not present.

The donor fate of the bone regenerate prompted us to examine the fracture callus and evaluate whether endochondral repair may differ from the traditional view of endochondral ossification during development. In regions of the fracture callus where cartilage transitions to bone (“transition zone”), there is an obvious phenotypic maturation of chondrocyte to hypertrophic chondrocyte to bone near the vasculature. We found limited evidence of apoptosis in hypertrophic chondrocytes at these transition zones. This result is supported by other studies showing that the hypertrophic chondrocytes may not be destined for apoptosis.[50-54] This does not exclude the possibility that cells from the invading vasculature also contribute to bone formation,[55] but rather suggests that invading cells may not be the only,[56] or even primary, method by which bone is formed during endochondral repair.

This study is not the first to suggest that cartilage can become bone, but the idea remains nondogmatic, and it is presently not clear whether this happens routinely during development and repair or only under certain circumstances. Further, if cartilage to bone transformation does occur, it is unclear which genetic or molecular sequences enable this change. Although some reports suggest that cartilage “dedifferentiates” to a progenitor-like state before becoming bone,[51, 57] other data support a mechanism where chondrocytes can mature directly into osteocytes.[50, 56, 58-63] In our study, we found that hypertrophic chondrocytes at the transition zone of the fracture callus appear to express the progenitor marker Oct4A. The mechanism by which Oct4A acts is not completely understood: In this situation, the chondrocytes could be dedifferentiating to regain progenitor capabilities, similar to induced pluripotent cells and consistent with the mechanism described by Song and Tuan.[57] Alternatively, Oct4A may function differently in adult somatic cells and allow cellular reprogramming to enable a direct transition from cartilage to bone. A significant challenge toward learning the mechanistic details of transformation is the substantial overlap between markers of hypertrophic chondrocytes and osteoblasts.[64-67] Consequently, lineage tracing and clearly defining cartilage versus bone tissue is difficult both in vitro and in vivo and relies heavily on differences in morphology. Further work is required to elucidate the extent to which Oct4A may reprogram chondrocytes during fracture healing.

Based on these data, we hypothesized that the vasculature is essential in coordinating the phenotypic transformation from cartilage to bone. To isolate the vasculature from the experiment, we cultured the cartilage grafts in vitro under osteogenic conditions containing BMP-2. The fracture callus cartilage explants largely maintained their hypertrophic morphology and regions of calcification and cartilage matrix staining overlapped. For the hMSC pellets, regions of mineralization generally occurred in areas where there was diminished proteogylcan content and more flattened cells were observed. The ability of MSC-derived cartilage to generate these so-called “chondro-osseo rudiments” in vitro has previously been described, demonstrating that this superficial region has the histological, microstructural, and immunohistochemical characteristics of bone.[38]

To more accurately recapitulate the microenvironment of the fracture callus in vitro, and specifically introduce the paracrine effect of vascular endothelial cells, we cultured the cartilage explants with HUVEC-conditioned medium. In a fracture, the invading vascular endothelial cells provide MMP-9[68] and BMPs[33, 69] to stimulate alkaline phosphatase activity,[70] but the complete set of bioactive stimuli is not fully defined. We found that HUVEC-conditioned media was sufficient to promote mineralization of the cartilage matrix, but still did not observe substantial morphological change of the hypertrophic chondrocyte to osteocyte. Taken together, these in vitro experiments suggest that although mineralized cartilage can be produced with soluble factors from vascular endothelial cells, or BMP-2 treatment, they alone are not sufficient to produce a complete transition to bone.

Degradation of the cartilage matrix and mechanical stimulation were two components of the in vivo environment that were not replicated by our in vitro conditions. Previous data from our laboratory have shown that cartilage remodeling is critical during fracture repair and the absence of MMPs results in impaired healing.[32, 68] Similarly, our laboratory has shown that mechanical stability influences the fate of skeletal progenitor cells during bone healing, with full stabilization promoting intramembranous repair and movement enabling endochondral ossification.[13] The role of mechanical loading during the transition cartilage to bone in the fracture callus has not been previously explored. It is possible that mechanical stimulation is central to this transition and contributed to the sustained hypertrophic phenotype observed in vitro and enabled the transition in the segmental defect. The murine model used in this study, although providing external stabilization to facilitate alignment, will not completely eliminate sheer and compressive forces between the host and graft bone. The precise effect of loading in regulating the transformation of cartilage to bone, both at a molecular or kinetic level, were not explored in this study but are important questions to consider in larger animal models.

Designing improved clinical treatments for repair and regeneration of bone is a central focus of our laboratory. Our study substantiates the concept of using cartilage grafts as an alternative therapeutic approach for bone regeneration. Further, evidence that the cartilage directly converts into bone during repair may also require change in how we approach the treatment of impaired healing in fractures. Full realization of the cartilage graft technology described in this study is presently limited by the ability to scale this approach to clinically relevant defects; a next step for this study will be to optimize a scaffold for a tissue engineering strategy[19] and translate this approach into a large animal study. The importance of understanding the mechanism by which cartilage can regenerate bone for therapeutic purposes comes in designing an appropriate tissue engineering approach to accurately recapitulate the regeneration sequences.

Acknowledgments

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Some materials and reagents were generously gifted from colleagues: Jennifer West (Rice University, 6kDa PEGDA), Gary Gibson (Henry Ford, Collagen X antibody), Anthony Hollender (University of Bristol, Collagen I antibody), and Francesco Curcio (Oct4A antibody). We also thank colleagues Alfred Kuo, Tamara Alliston, Nathan Young, and Brian Hall for their thoughtful review and discussion of our data. Thank you to Frank Yang for his assistance with murine husbandry and fractures, Alexis Lainoff for preparation of some of the probes for in situ, and to Gina Baldoza and Anna Lissa Wi for daily lab function and grants administration.

Research reported in this publication was supported by the National Institute of Arthritis and Musculoskeletal and Skin Disease (NIAMS) of the National Institutes of Health (NIH) under the following award numbers: 5F32AR062469-02 (CSB), AR053645 (TM), and AR057344 (TM). Additional research support was provided by the Musculoskeletal Transplant Foundation (CSB: MTF Junior Investigator Award), the UCSF Graduate Education in Medical Sciences (GEMS) and Clinical and Translational Science Institute (CTSI), and OHSU Gerlinger Research Award (BJ, CSB).

Authors' roles: Study design: CSB, DPH, BH, TM, and RSM. Study conduct and data collection: CSB, DPH, AJT, FF, and HMB. Data interpretation: CSB, DPH, AJT, FF, HMB, BH, BJ, TM, and RSM. Drafting manuscript: CSB. Revising manuscript: CSB and RM. All authors approved of final version of the manuscript. CSB takes responsibility for the integrity of the data.

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  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information
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Supporting Information

  1. Top of page
  2. ABSTRACT
  3. Introduction
  4. Materials and Methods
  5. Results
  6. Discussion
  7. Disclosures
  8. Acknowledgments
  9. References
  10. Supporting Information

Additional Supporting Information may be found in the online version of this article.

FilenameFormatSizeDescription
jbmr2148-sm-0001-SupFig-S1.tif9498KSupplementary Figure S1.
jbmr2148-sm-0002-SupFig-S2.tif4397KSupplementary Figure S2.
jbmr2148-sm-0003-SupFig-S3.tif3850KSupplementary Figure S3.
jbmr2148-sm-0004-SupFig-S4.tif2483KSupplementary Figure S4.
jbmr2148-sm-0005-SupFig-S5.tif4270KSupplementary Figure S5.
jbmr2148-sm-0006-SupFig-S6.tif7266KSupplementary Figure S6.
jbmr2148-sm-0007-SupFig-S7.tif4198KSupplementary Figure S7.
jbmr2148-sm-0008-SupFig-S8.tif9157KSupplementary Figure S8.
jbmr2148-sm-0009-SupFigsLegend-S1.doc41KSupplementary Figures Legend.

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