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ARTHRITIS & RHEUMATOLOGY
Vol. 67, No. 5, May 2015, pp 1274–1285
DOI 10.1002/art.39049
C 2015, American College of Rheumatology
V
Use of Recombinant Human
Stromal Cell–Derived Factor 1a–Loaded
Fibrin/Hyaluronic Acid Hydrogel Networks to Achieve
Functional Repair of Full-Thickness Bovine Articular Cartilage
Via Homing of Chondrogenic Progenitor Cells
Yin Yu,1 Marc J. Brouillette,1 Dongrim Seol,1 Hongjun Zheng,1 Joseph A. Buckwalter,2
and James A. Martin1
Results. Use of rhSDF-1a dramatically improved
CPC recruitment to the chondral defects at 12 days. After
6 weeks under chondrogenic conditions, cell morphology,
proteoglycan density, and the ultrastructure of the repair
tissue were all similar to that found in native cartilage.
Compared with empty controls, neocartilage generated
in rhSDF-1a–containing defects showed significantly
greater interfacial strength, and acquired mechanical
properties comparable to those of native cartilage.
Conclusion. This study showed that stimulating
local CPC recruitment prior to treatment with chondrogenic factors significantly improves the biochemical
and mechanical properties of the cartilage tissue
formed in chondral defects. This simple approach may
be implemented in vivo as a one-step procedure by
staging the release of chemokine and chondrogenic
factors from within the hydrogel, which can be achieved
using smart drug-delivery systems.
Objective. Articular cartilage damage after joint
trauma seldom heals and often leads to osteoarthritis.
We previously identified a migratory chondrogenic progenitor cell (CPC) population that responds chemotactically to cell death and rapidly repopulates the
injured cartilage matrix, which suggests a potential
approach for articular cartilage repair. This study was
undertaken to determine whether recombinant human
stromal cell–derived factor 1a (rhSDF-1a), a potent
CPC chemoattractant, would improve the quality of
cartilage regeneration, hypothesizing that increased
recruitment of CPCs by rhSDF-1a would promote
the formation of cartilage matrix upon chondrogenic
induction.
Methods. Full-thickness bovine chondral defects
were filled with hydrogel, composed of fibrin and
hyaluronic acid and containing rhSDF-1a. Cell migration was monitored, followed by chondrogenic induction. Regenerated tissue was evaluated by histology,
immunohistochemistry, and scanning electron microscopy. Push-out tests and unconfined compression tests
were performed to assess the strength of tissue integration and the mechanical properties of the regenerated cartilage.
Stem cell–based tissue-engineering treatments
using bone marrow–derived mesenchymal stem cells
(BM-MSCs) (1), as well as adipose-derived mesenchymal stem cells (AD-MSCs) (2), for adult human articular cartilage repair have drawn great attention and
been extensively studied (3). Although substantial success has been achieved with BM-MSCs, low cell yields
and phenotypic alterations during prolonged in vitro
cultivation remain problematic. Moreover, the chondrogenic activity of BM-MSCs varies according to the
subject’s age and the presence of osteoarthritis (OA).
AD-MSCs are more readily acquired than BM-MSCs,
but generate repair tissue with mechanical properties
that are inferior to hyaline cartilage.
Supported by the US Department of Defense (grant
W81XWH-10-1-0702a) and the American Arthritis Society.
1
Yin Yu, MS, Marc J. Brouillette, MS, Dongrim Seol, PhD,
Hongjun Zheng, PhD, James A. Martin, PhD: University of Iowa,
Iowa City; 2Joseph A. Buckwalter, MD: University of Iowa and Iowa
City VA Medical Center, Iowa City, Iowa.
Address correspondence to James A. Martin, PhD, 1182 Medical Laboratories, University of Iowa, Iowa City, IA 52242. E-mail:
james-martin@uiowa.edu.
Submitted for publication June 30, 2014; accepted in revised
form January 20, 2015.
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SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
Even in youth, the chondrogenic potential of
mesenchymal stem cells may be inferior to native chondrocytes, especially in an in vivo environment without
growth factor supplementation and in the presence of
proinflammatory cytokines. Stem cells may also display
a hypertrophic phenotype upon chondrogenic induction, which is undesirable for restoring an articular surface (4).
In addition to mesenchymal stem cells, pluripotent progenitor cells from multiple joint tissues, including the synovium (5), infrapatellar fat pad (6), and
meniscus (7), have shown chondrogenic potential.
However, it remains to be seen if any of these strategies consistently regenerate stable hyaline cartilage
that is well integrated with the surrounding matrix and
biologically and mechanically similar to native cartilage.
In addition, risks posed by these cell-based therapies,
such as pathogen transmission and tumorigenesis, as
well as the complexity of the ethical and regulatory
issues involved have limited the clinical implementation
of these therapies (8,9).
Articular cartilage tissue engineering by induction of cell homing without cell transplantation is a
provocative alternative that has already achieved notable success (10). In fact, a series of studies have identified subpopulations of stem/progenitor cells in articular
cartilage as well as repair tissue from patients with latestage OA (11–13). These cells, often referred to as
chondrogenic progenitor cells (CPCs), respond to various chemokines and cytokines and migrate toward
damaged cartilage tissue (14). They also exhibit other
characteristics of stem/progenitor cells, including an
apparent potential for repairing cartilage defects
(13–15). CPCs are thought to be a great candidate for
regenerative therapy in OA (16). However, to date, no
studies have demonstrated the regeneration of articular
cartilage using CPCs.
Stromal cell–derived factor 1a (SDF-1a) is a key
chemokine regulating stem cell migration and homing to
sites of tissue damage, where the cells participate in
tissue or organ regeneration. SDF-1a exerts its effects
through binding to the cell surface receptor CXCR4
(17,18). Recently, Seol and colleagues reported that
expression levels of SDF-1a and CXCR4 were highly
up-regulated in a migratory progenitor cell population
found on articular cartilage surfaces within a few days
after focal impact (14), and progenitor cells responded
vigorously to SDF-1a, which suggests that SDF-1a plays
a role in in situ articular cartilage repair by recruiting
endogenous stem/progenitor cells.
Fibrin and hyaluronic acid (HA) are classic biomaterials for articular cartilage regeneration. Their
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unique biocompatibility and highly hydrated structure
can mimic natural tissue and deliver biochemical cues
(19,20). A composite interpenetrating polymer network
(IPN) hydrogel composed of fibrin and HA has been
shown to exhibit mechanical properties that are far
superior to either polymer alone. The excellent cell
affinity of fibrin and delayed degradation of HA results
in mutually beneficial effects on the synthesis of cartilage extracellular matrix (ECM) (21).
In the present study, we attempted to repair
full-thickness articular cartilage defects in a bovine
osteochondral explant model by first enhancing the
recruitment of migratory progenitor cells to the IPN
using recombinant human SDF-1a (rhSDF-1a), followed by treatments to initiate chondrogenic differentiation. We hypothesized that, when compared with
controls lacking one or both factors, these sequential
manipulations would result in near-complete restoration of the cartilage matrix within the defect and
would improve integration with the host tissue.
MATERIALS AND METHODS
IPN hydrogel fabrication, drug release, and biocompatibility. The IPN hydrogel consisted of HA–thrombin
(solution A) and fibrinogen (solution B). For solution A,
10 mg/ml hyaluronate (GelOne; Zimmer) was mixed with the
same volume of 40 units/ml thrombin (TISSEEL; Baxter
Healthcare). Solution B was composed of 25 mg/ml fibrinogen
(TISSEEL; Baxter Healthcare) in Dulbecco’s phosphate buffered saline (DPBS; pH 7.4) with or without 400 ng/ml (or
200 ng/ml) rhSDF-1a (R&D Systems). To form the IPN, solution A and solution B were gently mixed together at a ratio of
1:1 under a temperature of 4 C. The final concentrations of
HA, thrombin, fibrinogen, and rhSDF-1a were 2.5 mg/ml,
10 units/ml, 12.5 mg/ml, and 200 ng/ml, respectively.
Cylindrical-shaped IPN hydrogel disks (2 mm in
thickness and 4 mm in diameter) were fabricated in a plastic mold and kept in DPBS for future use. The protein
release kinetics of rhSDF-1a were determined with the use
of a previously reported protocol (22). Briefly, each IPN
disk was placed in a well of a 24-well plate with 400 ml of
DPBS, and then cultured at 37 C. Supernatants were collected at each time point (days 2, 4, 6, 8, 10, 12, and 14).
DPBS (400 ml) was added to replenish each well and samples were placed back for cultivation until the next time
point. Enzyme-linked immunosorbent assay was used for
quantification, in accordance with the manufacturer’s instructions (MyBioSource).
To test the biocompatibility of the IPN, CPCs were
isolated in a manner as previously described (14) and were
encapsulated in IPN hydrogel disks (5 3 106 cells/ml) for an
in vitro viability assay using Live/Dead cell staining (23). The
stainings were performed at different time points after encapsulation (days 1, 7, and 21).
SDF-1a and CXCR4 receptor expression. To assess
the expression of SDF-1a and its receptor, CXCR4, upon
cartilage focal injury, immunofluorescence staining was used
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for cell surface markers, using a monoclonal anti–SDF-1a
antibody (Abcam) and anti-CXCR4 antibody (Santa Cruz
Biotechnology). A goat anti-mouse fluorescent secondary
antibody (Alexa Fluor 488) was used for fluorescent labeling and detection (Jackson ImmunoResearch) using confocal microscopy. Staining was performed on CPCs cultured
in monolayer or normal chondrocytes, as well as on cryosections of impacted articular cartilage or unimpacted fresh
cartilage tissue, as previously described (14). The expression levels of SDF-1a and CXCR4 were also compared
between CPCs and normal chondrocytes by real-time
reverse transcription–polymerase chain reaction (RT-PCR),
utilizing a previously described method (24). Each realtime RT-PCR experiment was done with at least 3 replicate
samples, and target gene expression is presented as messenger RNA (mRNA) levels normalized to the values for
b-actin.
IPN scaffold implantation, cell migration, and in
vitro chondrogenesis. Osteochondral explants (12-mm diameter and 8–10-mm thickness) were harvested from the bovine
femoral condyle (12–18 months of age, 9 animals in total).
After 2 days in the pre-equilibrium culture, full-thickness
chondral defects (4 mm in diameter and ;2 mm in thickness)
were created as previously described (14). Thereafter, the
explants were maintained in culture overnight, before IPN
implantation. The IPN (;60 ml), with or without the addition
of rhSDF-1a (100 or 200 ng/ml), was implanted into defects
slightly over the surface of the explants, which were then
placed back into culture.
To monitor cell migration, confocal microscopy was
performed essentially as previously described (15). Cell numbers were quantified by averaging automated cell counts from
6 random images (203 magnification) using ImageJ software
(25). The DNA content in the IPN hydrogel was quantified
following previously described procedures (15). Empty IPN
gel was used as a blank control.
Upon initiation of cell migration on day 12, the
explants were incubated in chondrogenic medium (Dulbecco’s modified Eagle’s medium containing 10 ng/ml transforming growth factor b1, 100 ng/ml insulin-like growth
factor 1, 0.1 mM dexamethasone, 25 mg/ml L-ascorbate, 100
mg/ml pyruvate, 50 mg/ml ITS1 [insulin–transferrin–
selenium] Premix, and antibiotics) in an atmosphere of 5%
CO2 at 37 C for up to 6 weeks. Regenerated tissue, together
with host cartilage, was harvested from the explants and analyzed for ECM formation using Safranin O–fast green staining of either cryosections (3 weeks) or paraffin-embedded
sections (6 weeks).
Immunohistochemical, biochemical, and ultrastructural evaluations of cartilage repair. For immunohistochemical analysis, deparaffinized sections from tissue samples obtained at 6 weeks were stained with type II
collagen and aggrecan antibodies (Developmental Studies
Hybridoma Bank). A goat anti-mouse secondary antibody
(Vector Laboratories) was used for detection. The reaction
products were visualized using the Vectastain ABC kit and
the DAB Peroxidase Substrate kit (both from Vector Laboratories), in accordance with the manufacturer’s instructions. In addition, staining for lubricin, an articular
cartilage superficial zone protein, was performed using a
rabbit polyclonal antibody, followed by detection with a
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goat anti-rabbit secondary antibody (Vector Laboratories.).
All negative controls were performed using the same staining but without use of the primary antibodies. The dimethylmethylene blue (DMMB) dye binding assay was used
for quantifying sulfated glycosaminoglycan (sGAG) content,
as previously described (15).
We also compared the water content between cartilage repair tissue and native cartilage, while blank IPN
hydrogel was used as a negative control. The wet weight of
all samples was measured with a bench-top scale (MettlerToledo). Dry weight was measured after overnight lyophilization at 245 C (Lobconco). Water content was determined using the following calculation: water content 5 ([wet
weight 2 dry weight]/wet weight) 3 100%. The cartilage
tissue, as well as freshly fabricated IPN gel, was harvested
at 6 weeks after chondrogenesis. Scanning electron microscopy samples were processed using previously described
methods (26), and all scanning electron microscopy tests
were performed at the University of Iowa Central Microscopy Research Facility.
Biomechanical assessment of tissue repair and
material properties of the regenerated tissue. In order to
evaluate integration (interfacial) strength between the
repair and host cartilage tissues, we performed push-out
tests using 6-week–cultured samples from both the rhSDF1a–treated group (n 5 9) and the untreated group (n 5 6).
A customized cartilage-fixation device rigidly held samples
to measure integration strength. Upon harvesting, the
specimens were then placed in the fixation device while a
LabVIEW (National Instruments)–controlled stepper motor
(Ultra Motion) depressed a cylindrical indenter (3.8 mm
in diameter) connected to a load cell (1 kg, 1 KHz sample
rate; Honeywell) at a constant velocity of 0.1 mm/second
(see Figure 5B). The test proceeded through the fullthickness of the tissue, and the integration strength was
determined, calculated as the maximum force recorded
divided by the area of integration.
To further characterize the mechanical properties of
the regenerated cartilage tissue, we used a materials testing
machine (MTS Systems) to perform stress–relaxation tests on
the regenerated cartilage as well as on the native cartilage tissue harvested from the explants. Briefly, the thickness of the
cartilage was measured using a laser measurement system
(Keyene Corporation of America), and the samples were then
placed in an unconfined chamber. A nonporous platen was
brought into contact with the tissue surface and the tissue was
compressed to 20% strain at a velocity of 1 mm/second or
2 mm/second. A 10N load cell, which recorded the load as
compression to 20% strain, was held for 20 minutes (see Figure 6C). Maximum stress, equilibrium stress, Young’s modulus
(ratio of stress to strain, a measure of cartilage elasticity), and
maximum force were recorded or calculated. This test was
applied to regenerated cartilage that had formed in defects
filled with SDF-1a–loaded IPN hydrogels (n 5 9 explants) following culture for 6 weeks. Native cartilage samples were harvested from the tibial plateau or femoral condyle cartilage
(n 5 8 explants each) of healthy bovine knee joints.
Statistical analysis. All data are presented as the
mean 6 SD. Data were analyzed by Student’s t-test, using
GraphPad Prism software (version 6). P values less than 0.05
were considered statistically significant.
SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
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Figure 1. Fabrication and characterization of the interpenetrating polymer network (IPN) hydrogel. A, Schematic drawing of IPN hydrogel fabrication. The fibrin hydrogel and hyaluronic acid (HA) polymer were blended and crosslinked to form the IPN. B, Macroscopic view of the IPN
scaffold. C and D, Scanning electron microscopy images showing interpenetrated polymer fibers (C) and interconnected pores (arrowheads) (D).
E–H, Continuous release of recombinant human stromal cell–derived factor 1a (rhSDF-1a) protein from the IPN scaffold in Dulbecco’s phosphate buffered saline at different time points over 14 days. Bars show the mean 6 SD of 4 explants per time point. I–K, Viability of the encapsulated chondrogenic progenitor cells (green fluorescence) on day 1 (I), day 7 (J), and day 21 (K), with minimal numbers of dead cells presented
(red fluorescence). L, Sustained average cell viability of $90% over 21 days. Results are the mean 6 SD of 6 explants per time point.
Bars 5 5 mm in B; 4 mm in E–G; 500 mm in I–K.
RESULTS
Fabrication and characterization of the IPN
scaffold. The IPN hydrogel could be readily formed by
thrombin-initiated crosslinking of fibrinogen to become
fibrin fibers, and was fully polymerized with a defined
shape under physiologic temperatures (37 C), with the
HA network fully penetrating the pores among the
fibrin fibers (Figure 1A). After polymerization, the IPN
scaffold displayed an opaque appearance and a welldefined disk shape (Figure 1B). Scanning electron
microscopy images showed that the HA network was
fully distributed within the fibrin fibers with great
homogeneity and interconnected pores, both from the
surface (Figure 1C) and the cross-section (Figure 1D).
This porous structure allows the cells to attach and
migrate both along the surface and within the implanted
IPN scaffold.
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Figure 2. Expression of rhSDF-1a and in vitro cell migration. A, Top, Immunofluorescence staining shows positivity (red fluorescence) for
SDF-1a and CXCR4 in monolayer-cultured chondrogenic progenitor cells (CPCs), whereas normal chondrocytes (NCs) are largely negative for
both markers and positive only for nuclear DAPI staining (blue fluorescence). Positive SDF-1a staining is also evident in impacted cartilage tissue, but not in unimpacted healthy cartilage. Bottom, Reverse transcription–polymerase chain reaction analyses show profound up-regulation of
SDF-1a (.13-fold) and CXCR4 (.3.5 fold) in CPCs compared with normal chondrocytes. Bars 5 200 mm. B, Schematic drawing shows the
experimental design of IPN implantation into bovine osteochondral defects. C, Left, Stacked confocal images from different time points show
dramatic cell migration in response to rhSDF-1a (100 or 200 ng/ml), in a concentration- and time-dependent manner, as compared with the
empty IPN control cultured in Dulbecco’s phosphate buffered saline alone. Bars 5 500 mm. D, Top, Quantification of high-magnification confocal images on day 12 (Day 12H) (n 5 8 per group) confirms that a significantly higher number of CPCs migrated in response to rhSDF-1a as
compared with empty IPN controls. Bottom, Quantification of double-stranded DNA (dsDNA) (n 5 8 per group) also suggests a much higher
dsDNA content in the rhSDF-1a–loaded IPNs as compared with controls. Results are the mean 6 SD. * 5 P , 0.05. NS 5 not significant (see
Figure 1 for other definitions).
The IPN scaffold maintained its integrity in
DPBS as long as 2 weeks without noticeable changes
(Figures 1E–G). The time-dependent release curve
showed that rhSDF-1a could be released over 14 days
(Figure 1H), with a sustained daily protein concentration of .2.0 ng/ml and a continuous releasing
trend.
CPCs were encapsulated in the IPN scaffold to
check their biocompatibility, expressed in terms of cell
viability. Confocal images showed a minimal number of
dead cells (red fluorescence), while most of the cells
were viable (green fluorescence) (Figures 1I–K). The
initial encapsulation process yielded a cell viability of
91.6 6 2.4 (mean 6 SD number of live cells) on day 1,
and the cell viability continued to remain at a high
level ($90%) up to 21 days (Figure 1L). These data
suggest that IPN scaffolds are easy to fabricate, are
able to support sustained release of rhSDF-1a, and are
biocompatible.
SDF-1a/CXCR4 expression and rhSDF-1a–
guided CPC migration. Immunofluorescence staining
showed high expression of SDF-1a protein in CPCs,
with .90% cells staining positive for SDF-1a
(Figure 2A, top right panel). In contrast, SDF-1a
protein expression was barely detectable in normal
chondrocytes (Figure 2A, top left panel). A similar pattern of expression was observed for CXCR4 (Figure
2A, middle panels). In impacted cartilage, expression
of SDF-1a was also significantly increased, throughout
the full depth of the tissue, compared with that in uninjured freshly isolated cartilage (Figure 2A, bottom panels), with stronger expression on the superficial and
middle zones. RT-PCR analyses revealed that SDF-1a
and CXCR4 mRNA expression was 13-fold and
SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
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Figure 3. Histologic and quantitative analyses of cartilage tissue regeneration. A–L, Safranin O–fast green staining of regenerated cartilage tissue. Stronger Safranin O–positive staining and more organized proteoglycan deposition are evident in the rhSDF-1a–treated group (IPN 1 SDF)
compared with untreated controls (IPN) at both 3 weeks (3W) (A–F) and 6 weeks (6W) (G–L). Also, at 3 weeks, cells in both groups display
the spindle shape characteristic of migrating CPCs (C and F), while at 6 weeks, the cells are more chondrocyte-like (spherical shape) (I and L).
Bars 5 1 mm in A, D, G, and J; 200 mm in B, E, H, and K; 50 mm in C, F, I, and L. M, Determination of sulfated glycosaminoglycan (sGAG)
content (normalized to wet weight), water content, and cell density in rhSDF-1a–treated cartilage compared with untreated cartilage. Results
are the mean 6 SD of 8 extracts per group. * 5 P , 0.05. HT 5 host tissue; RT 5 regenerated tissue (see Figure 1 for other definitions).
3.5-fold higher, respectively, in CPCs compared with
normal chondrocytes (P 5 0.0004).
Upon creation of the full-thickness articular cartilage defect and implantation of the IPN in the absence
of rhSDF-1a (in DPBS alone; empty IPN control) or in
the presence of rhSDF-1a (100 ng/ml or 200 ng/ml), we
monitored cell migration by confocal microscopy at different time points thereafter (Figure 2B). As clearly
shown in Figure 2C, in explants implanted with the
empty IPN control, very few cells migrated into the
defect area over 12 days, and the migrated cells were
mainly at the defect edge, leaving the majority of the
defect empty. For explants implanted with rhSDF-1a–
loaded IPN, a significant number of cells migrated from
the peripheral area to the center of the defect on day 7,
and more cells had migrated by day 12. Cell migration
occurred in an rhSDF-1a concentration–dependent
manner, with an increased number of migrating cells
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Figure 4. Immunohistochemical examination of cartilage tissue for articular cartilage–specific proteins. Immunohistochemical staining was carried out for type II collagen (COL2A) (A–C), aggrecan (AGC) (D–F), and lubricin (LUB) (G–I) in the rhSDF-1a–treated group compared with
the untreated controls (empty IPN) and the negative controls (without primary antibodies). Insets are lower-magnification views; boxed areas in
insets are shown at higher magnification in the main views. Bars 5 200 mm; 1 mm in insets. See Figure 1 for other definitions.
being observed in response to the higher dose of
rhSDF-1a (200 ng/ml) both on day 7 and on day 12.
Thus, 200 ng/ml rhSDF-1a was used in subsequent
studies of full-thickness cartilage repair.
To further quantify the effect of rhSDF-1a on
the migration of progenitor cells, higher-magnification
confocal images from day 12 (Day 12H in Figure 2C)
were used for automated cell counting. The IPN loaded
with rhSDF-1a (200 ng/ml) attracted .250% as many
cells as that in the IPN scaffold without rhSDF-1a
(P , 0.0001). Similarly, the double-stranded DNA
(dsDNA) content of the cartilage tissue on day 12 was
increased more than 2-fold in the presence of IPN
loaded with rhSDF-1a (200 ng/ml) as compared with
the empty IPN control (Figure 2D), whereas the
dsDNA levels were not significantly higher in the
rhSDF-1a (100 ng/ml)–loaded group compared with
empty controls. These observations suggest that exoge-
nous rhSDF-1a could act as a chemotactic cue for the
initiation of homing of progenitor cells to repopulate
full-thickness cartilage defects filled with IPN.
Histologic and immunohistochemical features
of the repaired cartilage tissue. We carried out histologic evaluations of the repaired cartilage defects to
identify production of cartilage ECM at the end of 3
weeks and 6 weeks. Three weeks after chondrogenic
induction, a substantially higher amount of proteoglycan deposition was observed in the rhSDF-1a–loaded
IPN scaffold, which displayed strong positive staining
for Safranin O (Figure 3D), as compared with the
empty IPN scaffold, which mainly displayed fast green
staining only (Figures 3A and B). Stronger Safranin O
staining was observed on the superficial zone of the
regenerated cartilage tissue and gradually decreased
into the deep zone (Figure 3E). Most of the migrated
cells still displayed a spindle-like morphology at 3
SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
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Figure 5. Assessment of cartilage tissue integration. A, Typical macroscopic appearance of repair tissues formed in defects with or without the addition of recombinant human stromal cell–derived factor 1a (rhSDF-1a). Left, The defect is still clearly visible in the untreated tissue, but not in the
rhSDF-1a–treated tissue. Center, Safranin O staining shows continuous proteoglycan-rich matrix in repair tissue with seamless connection to host
cartilage tissue in rhSDF-1a–treated defects, while untreated defects contain matrix that shows spotty Safranin O staining and poor adhesion to
native cartilage. Right, In rhSDF-1a–treated tissue, type II collagen shows well-organized, strong-intensity staining in the entire matrix of the interfacial area, while in the untreated tissue, staining presents only partially at the tissue interface. Bars 5 4 mm (left panels); 200 mm (middle and right
panels). B, Apparatus (top) and schematic diagram (bottom) for the push-out test. C, Comparison of stress and peak force between groups. Both
stress and peak force are significantly higher (.20-fold; P , 0.0001 and P 5 0.0004, respectively) in rhSDF-1a–treated tissue (n 5 9) compared with
untreated tissue (n 5 6). Results are the mean 6 SD. * 5 P , 0.05. D, Scanning electron microscopy images showing continuous cell ingrowth from
the surface (I) and in cross-section at the tissue interface (III), and also interconnected extracellular matrix (II) with entangled collagen fibers (IV).
Broken line marks the boundary between the host tissue (HT) and the regenerated tissue (RT).
weeks (Figures 3C and F), more similar to CPCs than
to chondrocytes (14).
Six weeks after chondrogenic differentiation,
both the empty IPN scaffold and the rhSDF-1a–loaded
IPN scaffold showed increased proteoglycan deposition
and stronger staining for Safranin O (Figures 3G and J)
compared with these features at 3 weeks. The rhSDF1a–loaded IPN scaffold yielded evenly distributed cells
and more intense Safranin O–positive staining for both
pericellular and interterritorial ECM throughout nearly
the whole depth of the regenerated tissue (Figure 3K).
In contrast, the empty IPN scaffold showed rather
uneven cell distribution with moderately positive Safranin O staining, mainly in the pericellular ECM (Figure
3H). The cells took on a chondrocyte-like spherical
morphology at 6 weeks, a sign of complete differentiation (Figures 3I and L), and cells in the rhSDF-1a–
loaded IPN scaffold had more similarity to host chondrocytes (Figure 3L).
Further quantification of sGAG by DMMB
assay showed that the rhSDF-1a–loaded IPN scaffold
yielded nearly 8-fold higher sGAG content than did
the empty IPN scaffold (P 5 0.0055) (Figure 3M, left
panel). Moreover, the regenerated cartilage tissue
from the rhSDF-1a–loaded IPN scaffold had significantly lower water content compared with the empty
IPN scaffold (P 5 0.0242) (Figure 3M, middle panel).
Quantification of cell density showed over twice as
many cells in the rhSDF-1a–loaded IPN group as in
the empty IPN control group (P , 0.0001) (Figure
3M, right panel). Interestingly, we observed a higher
cell density in cartilage repair tissue compared with
native cartilage on histologic images (results not
shown), and cell density in the repair tissue gradually
decreased from the superficial and middle zones to
the deep zone. This may be attributable to the fact
that most CPCs are located in the upper third of the
ECM (14).
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Figure 6. Biomechanical characterization of the regenerated cartilage (REGC) tissue. A, Scanning electron microscopy images showing the morphology of the cells and pattern of extracellular matrix fibers on the surface and the cross-section of host cartilage and regenerated cartilage tissue. B, Comparison of sulfated glycosaminoglycan (sGAG) content and water content between regenerated cartilage, host cartilage, and empty
interpenetrating polymer network (IPN) gel as blank control. Results are the mean 6 SD. C, Top, Apparatus and schematic diagram for the
stress–relaxation test. Bottom, Gross appearance of the 3 different cartilage tissues under the stress–relaxation test. D, Stress–strain curve for the
3 different cartilage tissues under loading rates of 1 mm/second and 2 mm/second. E, Maximum force, equilibrium stress, maximum stress, and
Young’s modulus in the 3 different cartilage tissues under loading rates of 1 mm/second and 2 mm/second. Results are the mean 6 SD of 8–9
different samples for each group. * 5 P , 0.05. NS 5 not significant; TPC 5 tibial plateau cartilage; FCC 5 femoral condyle cartilage.
Immunohistochemical analyses showed intense
positive staining for type II collagen as well as aggrecan
throughout the repair tissue from the rhSDF-1a–
loaded IPN, nearly identical to that in native cartilage
tissue (Figures 4C and F). In contrast, repair tissue
from the empty IPN displayed uneven and isolated
areas of type II collagen and aggrecan staining, which
was mainly pericellular and in the superficial zone
(Figures 4B and E).
The rhSDF-1a–loaded IPN scaffold yielded
regenerated tissue with strong positive staining for
lubricin, which was found mainly in the superficial
zone while relatively fewer positively stained cells were
observed in the middle and deep zones (Figure 4I).
These characteristics were all largely similar to that in
native cartilage. In contrast, repair tissue from the
empty IPN scaffold displayed disordered lubricin staining that was cluttered within the ECM (Figure 4H). A
great continuity of all 3 types of staining across the
surface of both native tissue and repair tissue was also
observed in the presence of the rhSDF-1a–loaded IPN
(insets of Figures 4C, F, and I), indicating a possible
SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
potential of this technique to restore the defective
articular cartilage surface. All negative controls
showed only light background staining (Figures 4A, D,
and G).
Integration of repair tissue with native cartilage. Macroscopic, ultrastructural, and histologic analyses of the junction between the defect and the host
tissue at 6 weeks showed that rhSDF-1a–loaded defects
were nearly seamlessly integrated with the host cartilage, a milestone of successful repair. Defects in the
absence of rhSDF-1a were not well integrated. Images
of Safranin O–fast green and type II collagen staining
showed a significantly improved repair–host tissue connection upon rhSDF-1a treatment, with subsequent
chondrogenesis (Figure 5A).
Push-out tests showed a dramatically different
integration strength between the rhSDF-1a–treated
and untreated groups. Both the stress and the peak
force were significantly higher in the rhSDF-1a–
treated group than in the untreated control group
(mean 6 SD stress 158.0 6 26.04 kPa versus 7.56 6 1.34
kPa; mean 6 SD peak force 3.23 6 0.53N versus
0.15 6 0.03N) (Figure 5C). In addition, scanning electron microscopy images of the rhSDF-1a–treated
group showed integration of the regenerated tissue
with the host cartilage, both in terms of cell ingrowth
and crosslinking of ECM fibers. The defect line was
largely closed by interconnected ECM fibers from both
the native tissue and the rhSDF-1a–treated regenerated tissue (Figure 5D).
Biochemical and mechanical properties of the
regenerated cartilage tissue. We further compared the
ultrastructure, sGAG content, water content, and various material properties between the regenerated cartilage and native cartilage. Scanning electron microscopy
images showed that cells in the regenerated tissue were
not as closely connected with the surrounding ECM as
were cells in the host cartilage. Moreover, cell density
was relatively higher in the regenerated tissue in comparison with the native tissue (Figure 6A, top panels).
Collagen fibers formed a less compacted network in
the regenerated cartilage compared with the native cartilage (Figure 6a, bottom panels), which may result in
mechanical properties that could differentiate regenerated from native cartilage.
The DMMB assay showed that the sGAG content was significantly increased in the regenerated tissue compared with cartilage tissue cultured with the
empty IPN control scaffold (P 5 0.0016). However,
sGAG content in the regenerated tissue was not significantly different from that in the host cartilage (P 5
0.2607). Similarly, water content was significantly
1283
decreased in the regenerated tissue compared with that
in the empty IPN control scaffold (P 5 0.0016), but was
not significantly different from that in the host cartilage
(Figure 6B).
In terms of mechanical properties (maximum
stress, equilibrium stress, Young’s modulus of elasticity,
and maximum force), the regenerated cartilage showed
higher values than did the tibial plateau cartilage and
lower values than did the femoral condyle cartilage, at
2 different testing speeds (Figures 6D and E). The
properties of the cartilage tissue filled with the empty
IPN control gel were too low to measure using our current testing system.
In the regenerated cartilage, the Young’s modulus was a mean 6 SD 746.7 6 82.3 kPa at a testing speed
of 1 mm/second and 965.4 6 78.9 kPa at a testing speed
of 2 mm/second, values that were notably higher than
those in cartilage from the tibial plateau (mean 6 SD
475.6 6 42.9 kPa and 542.8 6 46.1 kPa, respectively).
The Young’s modulus in the regenerated cartilage
reached only 70% of the values in the femoral condyle
cartilage. These results indicate that the mechanical
properties of the regenerated cartilage, measured using
physiologic loading rates, were well within the range for
native bovine cartilage. Notably, the regenerated cartilage
showed an increase in Young’s modulus with higher loading speed, similar to that in the tibial plateau and femoral
condyle cartilage.
DISCUSSION
The development of novel cartilage repair
strategies based on stimulating endogenous cell homing is of substantial clinical interest. In this study,
we show for the first time that full-thickness cartilage defects can be repaired entirely by endogenous
progenitor cells from articular cartilage, without
requiring cells from other sources (22,27,28). The
results demonstrated that the potential of intrinsic
cartilage healing can be enhanced by a 2-step strategy, first by initiating progenitor cell chemotaxis with
rhSDF-1a, followed by stimulation of chondrogenesis
with growth factors.
The expression of SDF-1a and CXCR4 upon cartilage injury supports the involvement of the SDF-1a/
CXCR4 axis in migration of CPCs to the site of a cartilage defect. SDF-1a also significantly increased progenitor cell migration from the surrounding cartilage into
IPN scaffolds, clearly demonstrating its ability to direct
progenitor cell homing. These results are consistent with
those from a number of published studies (7,29–32).
1284
Subsequent chondrogenic induction further stimulated
type II collagen and aggrecan deposition, resulting in
proteoglycan-rich cartilage matrix. The distribution of
lubricin staining, which tended to be stronger toward the
cartilage surface, suggests the potential for regenerating
stratified articular cartilage with zone-specific properties.
The regenerated cartilage and native cartilage showed
great similarities, in terms of sGAG and water content,
as well as in terms of ultrastructural collagen fiber alignment and cell–ECM interaction, all of which are essential
elements needed to support articular cartilage function.
The bonding of engineered cartilage with surrounding native tissue determines integration strength
(33). Our study showed that rhSDF-1a dramatically
increased the integration strength of treated cartilage,
compared with that of untreated controls. The average
stress value in the regenerated cartilage, 158.0 6 26.04
kPa, was more than 3 times higher than that reported
in comparable studies (34–36). This may indicate that
increased CPC migration enhanced tissue integration,
which is consistent with the results in a study by Lu
et al showing that cell migration at the interface of
engineered cartilage and surrounding cartilage could
result in dramatically stronger host–graft tissue integration after autologous chondrocyte implantation (37). It
is also worth noting that the collagen fiber networks of
the regenerated and host tissues in the fully treated
defects were extensively entangled with each other,
which might explain the gain in integration strength.
Regeneration of mechanically functional cartilage
tissue is key to the success of any cartilage repair strategy.
Although cartilage engineered from primary chondrocytes
has been found to reach physiologic equivalence with
native cartilage in terms of compressive moduli, these values in cartilage engineered from stem/progenitor cells
have been found to be no more than 50% of the values in
native cartilage. In our study, the Young’s modulus of the
tissue formed in large full-thickness chondral defects
exceeded that of the tibial plateau cartilage and rivaled
that of the femoral condyle cartilage. Moreover, this was
accomplished in a relatively short time in comparison with
other studies. Further improvement of mechanical performance may require loading stimulation, which has been
shown to enhance the Young’s modulus of engineered cartilage (38). For in vivo translation, the IPN gel may not be
able to withstand initial mechanical stresses like repetitive
loading, and therefore certain immobilization procedures
may be needed during the early stages of neocartilage
development, after which physiologic loading would be
beneficial for further maturation.
Although the results are promising, there are certainly limitations within this study. Healthy young cows
YU ET AL
may have a regenerative capacity that is superior to that
in aged animals. This may limit the translatability of our
strategy, especially for elderly patients with OA, since
CPCs from OA patients may have limited chondrogenic
potential, due either to an altered phenotype or to the
unfriendly environment in which the cells reside. Various
inflammatory factors, such as interleukin-1b, tumor
necrosis factor a, or nitric oxide, could also inhibit the
migration activity of CPCs in OA (39). More refined
strategies could be developed, particularly strategies that
would not only incorporate chemotactic factors for cell
homing but also modify scaffolds by introducing antiinflammatory agents, which would certainly have profound
benefits for cartilage neogenesis. In terms of translation
in vivo, approaches of efficient delivery and retention of
these factors at sites of damage will need to be carefully
designed. This can be achieved by encapsulating chemokines, growth factors (40), or genetic materials (41)
within polymer microspheres to achieve sustained or
multiphase release from the scaffold.
We have developed a cartilage repair strategy that
exploits the regenerative potential of endogenous chondrogenic progenitor cells. The matrix formed by these cells
is similar in composition to native cartilage and strongly
adheres to surrounding tissues. Regenerated cartilage tissue possesses mechanical properties within the recognized
physiologic range of functional native cartilage. Optimization of this strategy could lead to a new, minimally invasive
single-step procedure for cartilage repair.
ACKNOWLEDGMENTS
The authors would like to thank Mr. Jianqiang Shao (University of Iowa Central Microscopy Research Facility) for providing help with the scanning electron microscopy imaging, and Dr.
Anneliese Heiner (University of Iowa Orthopaedic Biomechanics
Laboratory) for providing help with the mechanical testing.
AUTHOR CONTRIBUTION
All authors were involved in drafting the article or revising it
critically for important intellectual content, and all authors approved
the final version to be published. Dr. Martin had full access to all of
the data in the study and takes responsibility for the integrity of the
data and the accuracy of the data analysis.
Study conception and design. Yu, Martin.
Acquisition of data. Yu, Brouillette.
Analysis and interpretation of data. Yu, Brouillette, Seol, Zheng,
Buckwalter.
REFERENCES
1. Pittenger MF, Mackay AM, Beck SC, Jaiswal RK, Douglas R,
Mosca JD, et al. Multilineage potential of adult human mesenchymal stem cells. Science 1999;284:143–7.
2. Erickson IE, Kestle SR, Zellars KH, Farrell MJ, Kim M,
Burdick JA, et al. High mesenchymal stem cell seeding densities
SDF-1a FOR ARTICULAR CARTILAGE REPAIR UPON CHONDROGENIC INDUCTION
3.
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
in hyaluronic acid hydrogels produce engineered cartilage with
native tissue properties. Acta Biomater 2012;8:3027–34.
Tuan RS. Stemming cartilage degeneration: adult mesenchymal
stem cells as a cell source for articular cartilage tissue engineering [editorial]. Arthritis Rheum 2006;54:3075–8.
Johnstone B, Alini M, Cucchiarini M, Dodge GR, Eglin D,
Guilak F, et al. Tissue engineering for articular cartilage repair:
the state of the art. Eur Cell Mater 2013;25:248–67.
De Bari C, Dell’Accio F, T ylzanowski P, Luyten FP. Multipotent mesenchymal stem cells from adult human synovial membrane. Arthritis Rheum 2001;44:1928–42.
Wickham MQ, Erickson GR, Gimble JM, Vail TP, Guilak F.
Multipotent stromal cells derived from the infrapatellar fat pad
of the knee. Clinical Orthop Relat Res 2003;412:196–212.
Shen W, Chen J, Zhu T, Chen L, Zhang W, Fang Z, et al.
Intra-articular injection of human meniscus stem/progenitor cells
promotes meniscus regeneration and ameliorates osteoarthritis
through stromal cell-derived factor-1/CXCR4-mediated homing.
Stem Cells Transl Med 2014;33:387–94.
Fodor WL. Tissue engineering and cell based therapies, from
the bench to the clinic: the potential to replace, repair and
regenerate. Reprod Biol Endocrinol 2003;1:102.
Prockop DJ. Repair of tissues by adult stem/progenitor cells
(MSCs): controversies, myths, and changing paradigms. Mol
Therapy 2009;17:939–46.
Lee CH, Cook JL, Mendelson A, Moioli EK, Yao H, Mao JJ.
Regeneration of the articular surface of the rabbit synovial joint
by cell homing: a proof of concept study. Lancet 2010;376:440–8.
Dowthwaite GP, Bishop JC, Redman SN, Khan IM, Rooney P,
Evans DJ, et al. The surface of articular cartilage contains a progenitor cell population. J Cell Sci 2004;117:889–97.
Alsalameh S, Amin R, Gemba T, Lotz M. Identification of mesenchymal progenitor cells in normal and osteoarthritic human
articular cartilage. Arthritis Rheum 2004;50:1522–32.
Koelling S, Kruegel J, Irmer M, Path JR, Sadowski B, Miro X,
et al. Migratory chondrogenic progenitor cells from repair tissue
during the later stages of human osteoarthritis. Cell Stem Cell
2009;4:324–35.
Seol D, McCabe DJ, Choe H, Zheng H, Yu Y, Jang K, et al.
Chondrogenic progenitor cells respond to cartilage injury.
Arthritis Rheum 2012;64:3626–37.
Seol D, Yu Y, Choe H, Jang K, Brouillette MJ, Zheng H, et al.
Effect of short-term enzymatic treatment on cell migration and
cartilage regeneration: in vitro organ culture of bovine articular
cartilage. Tissue Eng Part A 2014;20:1807–14.
Dealy CN. Chondrogenic progenitors for cartilage repair and
osteoarthritis treatment [editorial]. Rheumatology (Sunnyvale)
2012;S3:e001.
Hattori K, Heissig B, Tashiro K, Honjo T, Tateno M, Shieh JH,
et al. Plasma elevation of stromal cell-derived factor-1 induces
mobilization of mature and immature hematopoietic progenitor
and stem cells. Blood 2001;97:3354–60.
Lagasse E, Connors H, Al-Dhalimy M, Reitsma M, Dohse M,
Osborne L, et al. Purified hematopoietic stem cells can differentiate into hepatocytes in vivo. Nat Med 2000;6:1229–34.
Klein TJ, Rizzi SC, Reichert JC, Georgi N, Malda J, Schuurman
W, et al. Strategies for zonal cartilage repair using hydrogels.
Macromol Biosci 2009;9:1049–58.
Slaughter BV, Khurshid SS, Fisher OZ, Khademhosseini A,
Peppas NA. Hydrogels in regenerative medicine. Adv Mater
2009;21:3307–29.
Rampichova M, Filova E, Varga F, Lytvynets A, Prosecka E,
Kolacna L, et al. Fibrin/hyaluronic acid composite hydrogels as
appropriate scaffolds for in vivo artificial cartilage implantation.
ASAIO J 2010;56:563–8.
Sukegawa A, Iwasaki N, Kasahara Y, Onodera T, Igarashi T,
Minami A. Repair of rabbit osteochondral defects by an acellular technique with an ultrapurified alginate gel containing stromal cell-derived factor-1. Tissue Eng Part A 2012;18:934–45.
1285
23. Yu Y, Zhang Y, Martin JA, Ozbolat IT. Evaluation of cell viability and functionality in vessel-like bioprintable cell-laden tubular channels. J Biomech Eng 2013;135:91011.
24. Seol D, Choe H, Zheng H, Jang K, Ramakrishnan PS, Lim TH,
et al. Selection of reference genes for normalization of quantitative real-time PCR in organ culture of the rat and rabbit intervertebral disc. BMC Res Notes 2011;4:162.
25. Schneider CA, Rasband WS, Eliceiri KW. NIH Image to
ImageJ: 25 years of image analysis. Nat Methods 2012;9:671–5.
26. Swords WE, Chance DL, Cohn LA, Shao J, Apicella MA, Smith
AL. Acylation of the lipooligosaccharide of Haemophilus influenzae and colonization: an htrB mutation diminishes the colonization
of human airway epithelial cells. Infect Immun 2002;70:4661–8.
27. Zhang W, Chen JL, Tao JD, Jiang YZ, Hu CC, Huang L, et al.
The use of type 1 collagen scaffold containing stromal cell-derived
factor-1 to create a matrix environment conducive to partialthickness cartilage defects repair. Biomaterials 2013;34:713–23.
28. Mendelson A, Frank E, Allred C, Jones E, Chen M, Zhao W, et al.
Chondrogenesis by chemotactic homing of synovium, bone marrow,
and adipose stem cells in vitro. FASEB J 2011;25:3496–504.
29. Thevenot PT, Nair AM, Shen J, Lotfi P, Ko CY, Tang L. The
effect of incorporation of SDF-1a into PLGA scaffolds on stem
cell recruitment and the inflammatory response. Biomaterials
2010;31:3997–4008.
30. Schantz JT, Chim H, Whiteman M. Cell guidance in tissue engineering: SDF-1 mediates site-directed homing of mesenchymal
stem cells within three-dimensional polycaprolactone scaffolds.
Tissue Eng 2007;13:2615–24.
31. Shen W, Chen X, Chen J, Yin Z, Heng BC, Chen W, et al. The
effect of incorporation of exogenous stromal cell-derived factor1a within a knitted silk-collagen sponge scaffold on tendon
regeneration. Biomaterials 2010;31:7239–49.
32. Kitaori T, Ito H, Schwarz EM, Tsutsumi R, Yoshitomi H, Oishi S,
et al. Stromal cell–derived factor 1/CXCR4 signaling is critical for
the recruitment of mesenchymal stem cells to the fracture site during
skeletal repair in a mouse model. Arthritis Rheum 2009;60:813–23.
33. Obradovic B, Martin I, Padera RF, Treppo S, Freed LE,
Vunjak-Novakovic G. Integration of engineered cartilage.
J Orthop Res 2001;19:1089–97.
34. Diekman BO, Christoforou N, Willard VP, Sun H, SanchezAdams J, Leong KW, et al. Cartilage tissue engineering using
differentiated and purified induced pluripotent stem cells. Proc
Natl Acad Sci 2012;109:19172–7.
35. Tam HK, Srivastava A, Colwell CW Jr, D’Lima DD. In vitro
model of full-thickness cartilage defect healing. J Orthop Res
2007;25:1136–44.
36. Theodoropoulos JS, De Croos JN, Park SS, Pilliar R, Kandel RA.
Integration of tissue-engineered cartilage with host cartilage: an in
vitro model. Clin Orthop Rel Res 2011;469:2785–95.
37. Lu Y, Xu Y, Yin Z, Yang X, Jiang Y, Gui J. Chondrocyte migration affects tissue-engineered cartilage integration by activating the signal transduction pathways involving Src, PLCg1, and
ERK1/2. Tissue Eng Part A 2013;19:2506–16.
38. Bian L, Fong JV, Lima EG, Stoker AM, Ateshian GA, Cook
JL, et al. Dynamic mechanical loading enhances functional properties of tissue-engineered cartilage using mature canine chondrocytes. Tissue Eng Part A 2010;16:1781–90.
39. Joos H, Wildner A, Hogrefe C, Reichel H, Brenner RE. Interleukin-1b and tumor necrosis factor a inhibit migration activity
of chondrogenic progenitor cells from non-fibrillated osteoarthritic cartilage. Arthritis Res Ther 2013;15:R119.
40. Eswaramoorthy R, Chang CC, Wu SC, Wang GJ, Chang JK, Ho
ML. Sustained release of PTH(1-34) from PLGA microspheres
suppresses osteoarthritis progression in rats. Acta Biomater
2012;8:2254–62.
41. Ha CW, Noh MJ, Choi KB, Lee KH. Initial phase I safety of
retrovirally transduced human chondrocytes expressing transforming growth factor-b-1 in degenerative arthritis patients.
Cytotherapy 2012;14:247–56.