ArticlePDF Available

Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells without Exogenous Stimuli

Authors:

Abstract and Figures

Stem cell fate has been linked to the mechanical properties of their underlying substrate, affecting mechanoreceptors and ultimately leading to downstream biological response. Studies have used polymers to mimic the stiffness of extracellular matrix as well as of individual tissues and shown mesenchymal stem cells (MSCs) could be directed along specific lineages. In this study, we examined the role of stiffness in MSC differentiation to two closely related cell phenotypes: osteoblast and chondrocyte. We prepared four methyl acrylate/methyl methacrylate (MA/MMA) polymer surfaces with elastic moduli ranging from 0.1 MPa to 310 MPa by altering monomer concentration. MSCs were cultured in media without exogenous growth factors and their biological responses were compared to committed chondrocytes and osteoblasts. Both chondrogenic and osteogenic markers were elevated when MSCs were grown on substrates with stiffness <10 MPa. Like chondrocytes, MSCs on lower stiffness substrates showed elevated expression of ACAN, SOX9, and COL2 and proteoglycan content; COMP was elevated in MSCs but reduced in chondrocytes. Substrate stiffness altered levels of RUNX2 mRNA, alkaline phosphatase specific activity, osteocalcin, and osteoprotegerin in osteoblasts, decreasing levels on the least stiff substrate. Expression of integrin subunits α1, α2, α5, αv, β1, and β3 changed in a stiffness- and cell type-dependent manner. Silencing of integrin subunit beta 1 (ITGB1) in MSCs abolished both osteoblastic and chondrogenic differentiation in response to substrate stiffness. Our results suggest that substrate stiffness is an important mediator of osteoblastic and chondrogenic differentiation, and integrin β1 plays a pivotal role in this process.
Content may be subject to copyright.
RESEARCH ARTICLE
Substrate Stiffness Controls Osteoblastic and
Chondrocytic Differentiation of Mesenchymal
Stem Cells without Exogenous Stimuli
Rene Olivares-Navarrete
1
, Erin M. Lee
2
, Kathryn Smith
3
, Sharon L. Hyzy
1
,
Maryam Doroudi
4
, Joseph K. Williams
5
, Ken Gall
3,6
, Barbara D. Boyan
1,2
*, Zvi Schwartz
1,7
1Department of Biomedical Engineering, Virginia Commonwealth University, Richmond, Virginia, United
States of America, 2Wallace H. Coulter Department of Biomedical Engineering, Georgia Institute of
Technology, Atlanta, Georgia, United States of America, 3Medshape, Inc., Atlanta, Georgia, United States of
America, 4School of Biology, Georgia Institute of Technology, Atlanta, Georgia, United States of America,
5Children’s Healthcare of Atlanta, Atlanta, Georgia, United States of America, 6Department of Mechanical
Engineering and Materials Science, Duke University, Durham, North Carolina, United States of America,
7Department of Periodontics, University of Texas Health Science Center at San Antonio, San Antonio,
Texas, United States of America
These authors contributed equally to this work.
*bboyan@vcu.edu
Abstract
Stem cell fate has been linked to the mechanical properties of their underlying substrate,
affecting mechanoreceptors and ultimately leading to downstream biological response. Stud-
ies have used polymers to mimic the stiffness of extracellular matrix as well as of individual
tissues and shown mesenchymal stem cells (MSCs) could be directed along specific line-
ages. In this study, we examined the role of stiffness in MSC differentiation to two closely
related cell phenotypes: osteoblast and chondrocyte. We prepared four methyl acrylate/
methyl methacrylate (MA/MMA) polymer surfaces with elastic moduli ranging from 0.1 MPa
to 310 MPa by altering monomer concentration. MSCs were cultured in media without exoge-
nous growth factors and their biological responses were compared to committed chondro-
cytes and osteoblasts. Both chondrogenic and osteogenic markers were elevated when
MSCs were grown on substrates with stiffness <10 MPa. Like chondrocytes, MSCs on lower
stiffness substrates showed elevated expression of ACAN, SOX9, and COL2 and proteogly-
can content; COMP was elevated in MSCs but reduced in chondrocytes. Substrate stiffness
altered levels of RUNX2 mRNA, alkaline phosphatase specific activity, osteocalcin, and
osteoprotegerin in osteoblasts, decreasing levels on the least stiff substrate. Expression of
integrin subunits α1, α2, α5, αv, β1, and β3 changed in a stiffness- and cell type-dependent
manner. Silencing of integrin subunit beta 1 (ITGB1) in MSCs abolished both osteoblastic
and chondrogenic differentiation in response to substrate stiffness. Our results suggest that
substrate stiffness is an important mediator of osteoblastic and chondrogenic differentiation,
and integrin β1 plays a pivotal role in this process.
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 1 / 18
a1111111111
a1111111111
a1111111111
a1111111111
a1111111111
OPEN ACCESS
Citation: Olivares-Navarrete R, Lee EM, Smith K,
Hyzy SL, Doroudi M, Williams JK, et al. (2017)
Substrate Stiffness Controls Osteoblastic and
Chondrocytic Differentiation of Mesenchymal Stem
Cells without Exogenous Stimuli. PLoS ONE 12(1):
e0170312. doi:10.1371/journal.pone.0170312
Editor: Andre van Wijnen, University of
Massachusetts Medical School, UNITED STATES
Received: July 27, 2016
Accepted: January 2, 2017
Published: January 17, 2017
Copyright: ©2017 Olivares-Navarrete et al. This is
an open access article distributed under the terms
of the Creative Commons Attribution License,
which permits unrestricted use, distribution, and
reproduction in any medium, provided the original
author and source are credited.
Data Availability Statement: All relevant data are
within the paper and its Supporting Information
files.
Funding: Research reported in this publication was
supported by the National Institute of Arthritis and
Musculoskeletal and Skin Diseases of the National
Institutes of Health (http://www.niams.nih.gov/)
under Award Numbers AR054339 received by KG
and BDB and AR052102 received by BDB. The
funders had no role in study design, data collection
and analysis, decision to publish, or preparation of
Introduction
Millions of medical devices are implanted in Americans every year. These devices have a wide
array of mechanical, chemical, and morphological properties. In vivo, implant surface proper-
ties including roughness, chemistry, energy, and topography affect bone-to-implant contact
[14]. In vitro studies suggest that this is in part by stimulating osteoblastic differentiation of
mesenchymal stem cells (MSCs) during bone healing [5].
Several reports have shown that MSCs are sensitive to substrate properties, such as surface
roughness, stiffness, chemistry, and energy, and differentiate along specific lineages in response
to these cues [610]. Substrate material properties play a role in inducing MSC differentiation
into osteoblasts [1113], even in the absence of exogenous factors or media supplements fre-
quently used to stimulate osteogenesis in cultures grown on tissue culture polystyrene (TCPS)
[5]. The specific role of stiffness has been more difficult to determine. Efforts to recapitulate the
mechanical properties of extracellular matrix have suggested that specific stiffness can contrib-
ute to stem cell fate [14,15], but whether osteoblast differentiation is mediated by specific stiff-
ness is not clear. Many studies were performed on metal and polymer substrates with lower or
higher moduli range than native moduli of bone where such biomaterials generally are placed.
Moreover, few studies have examined whether the effects of stiffness and chemistry are unique
to osteoblastic differentiation or if other mesenchymal lineage fates might be induced as well.
Cells use mechanoreceptors to detect substrate stiffness via a mechanism that involves
integrin-dependent signaling [14]. We have shown that integrin expression in MSCs and oste-
oblasts is modulated by surface properties, with α5β1 being expressed on smooth titanium and
titanium alloy substrates and α2β1 being expressed on microtextured surfaces. Whereas α5β1
is associated with attachment and proliferation [16], α2β1 signaling is required for osteoblast
differentiation [17]. Integrin β1 has been shown to mediate effects of other material and envi-
ronmental stimuli on cell response [18,19] and has been demonstrated to play a role in chon-
drogenic differentiation [20,21].
Many studies examining how these properties modulate differentiation of multipotent cells
like MSCs have focused on a single lineage fate. Relatively little is known about how changes in
the chemical and mechanical microenvironment of these cells might differentially modulate
phenotypic expression along multiple lineages [14,22]. In vivo, MSCs reside in tissues of vary-
ing stiffness and participate in tissue regeneration with stiffness changing as the repair tissue
matures. This suggests that cells at different states within a lineage may respond differentially
as they commit to a specific fate. To begin to examine this, we developed a series of polymer
substrates with varying stiffness but without major changes in surface chemistry [23]. We
found that a relatively high stiffness of 850 MPa was able to induce maturation of osteoblast-
like MG63 cells. In the present study, we took advantage of methacrylate/methylmethacrylate
polymer networks in which stiffness could be controlled by varying the amount of monomer
[24], to investigate how stiffness mediates MSC commitment to two related lineages, osteo-
genic and chondrogenic, and compared MSC responses to those of committed osteoblasts and
chondrocytes.
Materials and Methods
Polymer synthesis
Polymer substrates with elastic moduli orders of magnitude apart were synthesized to examine
the effects of stiffnesses in ranges beyond those reported in the current literature and with
moduli relevant to clinical applications. To accomplish this, we varied the weight ratio of
methyl acrylate (MA) and methyl methacrylate (MMA) crosslinked with 10% poly(ethylene
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 2 / 18
the manuscript. KS is an employee of MedShape,
Inc. KS and KG have equity in MedShape, Inc.
MedShape, Inc. provided support in the form of
salary for author KS, but did not have any
additional role in the study design, data collection
and analysis, decision to publish, or preparation of
the manuscript.
Competing Interests: KS is an employee of
MedShape, Inc. KS and KG have equity in
MedShape, Inc. This does not alter our adherence
to PLOS ONE policies on sharing data and
materials.
glycol) dimethacrylate (PEGDMA) [24]. Copolymer solutions consisting of MA, MMA, and
PEGDMA MW~750 were obtained from Sigma-Aldrich and used as received. The weight
ratio of MA to MMA was varied while the crosslinking concentration of PEGDMA was held
constant at 10 wt% to produce four copolymer networks (by wt. % of MA): 18MA, 29MA,
40MA, and 72MA. 0.5 wt.% 2,2-dimethoxy-2-phenylacetophenone (DMPA) was used as the
photoinitiator (Sigma-Aldrich). Each solution was mixed manually in a glass vial and injected
between two glass slides using a glass pipette. Slides were separated with two 1mm glass spac-
ers. The samples were placed in a UV chamber (Model CL-1000L Ultraviolet Crosslinker; λ=
365nm; energy = 2000x100μJ/cm 2) for 30 minutes. Discs were laser-cut from the polymerized
sheets to a diameter such that the disc of each composition would swell to fill the bottom of a
well in a 24-well cell culture plate when incubated in cell culture media. All discs were post-
cured in an oven at 90˚C for 90 minutes and boiled in distilled water for 30 minutes to remove
excess monomer. Finally, discs were sterilized by UV light (λ= 254nm) for 90 minutes.
Mechanical testing
Tensile strain-to-failure tests to determine toughness and elastic modulus were performed on
a universal testing machine (MTS Insight 2) using a 2kN load cell and a strain rate of 5%/s.
ASTM D632 Type IV Dogbone samples were laser-cut with a 20 mm gauge length and 2.8 mm
gauge width, and their edges were sanded to remove any laser-induced defects. Samples were
soaked in phosphate buffered saline (PBS) for 24 hours prior to testing, removed from PBS,
patted with a paper towel to remove excess PBS, and their dimensions measured using digital
calipers. Following this, the samples were loaded in tensile grips, submerged in a PBS bath at
37˚C, and held at 37˚C for 10 min to allow for thermal equilibration.
Toughness was calculated as the area under the stress-strain curve in units of MJ/m
3
. Elastic
modulus was determined by calculating the slope of the linear portion of the stress-strain
curve (n = 4). Dynamic mechanical analysis (DMA) in tensile loading was used to determine
the rubbery modulus of the networks corresponding to the degree of crosslinking (TA Q800
DMA, Newcastle, DE). Rectangular samples of 1 x 5 x 15 mm
3
were laser cut from polymer
sheets, and their edges were sanded to remove any defects from the laser. The samples were
thermally equilibrated at -75˚C for 2 minutes and then heated to 200˚C at a rate of 5˚C/min-
ute. Testing was performed in cyclic strain control at 0.2% strain with a preload force of 0.001
N and a force track setting of 150%. The glass transition temperature (T
g
) was defined as the
peak of the tan delta curve, and the rubbery modulus was measured as the storage modulus
value taken 20˚C beyond the lowest point in the rubbery plateau (n = 3).
Surface characterization
Surface wettability was determined by performing contact angle measurements using the ses-
sile drop method (Rame
´-hart Model 250 goniometer, Mountain Lakes, NJ) (n = 3). FTIR-ATR
spectra were obtained on discs using a Bruker Optics Tensor Spectrometer (Billerica, MA)
with a KBr crystal. Ten scans were performed on each sample at a 1 Hz frequency, and peak
wavenumbers were determined using OMNIC software (Thermo Electron Corporation,
Madison, WI). Three spectra were obtained for three separate discs for each composition.
Cell studies
Human MSCs and human osteoblasts (HOBs, Lonza) were obtained from Lonza (Walkersville,
MO). Human auricular chondrocytes were isolated from pediatric ear cartilage obtained under
an IRB-approved protocol at Children’s Hospital of Atlanta and Georgia Institute of Technol-
ogy. Informed consent was from the guardian and was in written form. The chondrocytes were
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 3 / 18
isolated as described previously [25], cultured to confluence, and stored at -80˚C until used for
this study. Auricular chondrocytes were chosen for their applications in tissue engineering,
including their ability to proliferate and maintain their phenotype in culture [2630]. In addi-
tion, we were interested in the modulation of phenotype along closely related lineages. Accord-
ingly, we used auricular chondrocytes rather than articular or growth plate chondrocytes, to
better identify specific stiffness modulating differentiation to an osteoblast or chondrocyte
lineage.
Cells in passage two were used for all studies. Expression of cartilage cell phenotype at this
passage was verified by gene expression of SOX9, ACAN, COL2, and COMP (S1 Fig), as
described in the following paragraph. We did not assess expression of mRNA for elastin, a
marker of the auricular chondrocyte phenotype, as our intent was to examine the general
properties of osteoblasts v. chondrocyte lineage commitment.
All cells were grown plated at a density of 10,000 cells/cm
2
on copolymer surfaces and cul-
tured in Dulbecco’s modified Eagle’s medium (Corning, Manassas, VA) supplemented with
10% fetal bovine serum (Life Technologies, Carlsbad, CA) and 1% penicillin-streptomycin
(Life Technologies). Cells were fed with this medium for 24 hours post-plating and every other
day. After 7 days of culture, cells were incubated with fresh medium for 12 hours. RNA was
isolated (TRIzol, Life Technologies) and quantified using a NanoDrop spectrophotometer
(Thermo Scientific, Waltham, MA). To create a cDNA template, 500 ng of RNA was reverse
transcribed using a High Capacity Reverse Transcription cDNA kit (Life Technologies). To
quantify expression of RUNX2 mRNA in MSCs and HOBs, cDNA was used for real-time PCR
with gene-specific primers (S1 Table) using the StepOnePlus Real-time PCR System and
Power SYBR1Green PCR Master Mix (Life Technologies). Fluorescence values were quanti-
fied as starting quantities using known dilutions of cells cultured on tissue culture polystyrene
(TCPS). mRNA expression was normalized to glyceraldehyde 3-phosphate dehydrogenase.
Total cell number and cellular alkaline phosphatase specific activity were measured in the cell
lysate as previously described [31].
Secreted osteocalcin (OCN, Biomedical Technologies, Stoughton, MA) and osteoprotegerin
(R&D Systems, Minneapolis, MN) were measured to determine osteogenic differentiation.
Immunoassays were normalized to total cell number. Chondrogenic differentiation was deter-
mined by measuring the expression of mRNAs for aggrecan (ACAN), cartilage oligomeric
matrix protein (COMP), and collagen type II (COL2) as described above. Cartilage matrix pro-
duction was assessed using an Alcian blue assay (Sigma-Aldrich, St. Louis, MO) to measure
sulfated glycosaminoglycans. In brief, cell layers were fixed with 10% neutral buffered formalin
for 10 minutes at room temperature. Cells were washed twice with PBS, then incubated with
3% acetic acid for 10 minutes. Proteoglycans were stained with 1% Alcian blue in 3% acetic
acid (pH 2.5) for 30 minutes at room temperature. Cell layers were washed twice, and Alcian
blue was extracted with dimethyl sulfoxide. Absorbance was measured at 650 nm [32].
For all experiments, MSCs, HOBs, and chondrocytes were grown at the same time with the
same culture media to limit variability. To visualize cell shape, MSCs, HOBs and chondrocytes
were plated on copolymer surfaces at a density of 5,000 cells/cm
2
and allowed to spread for 24
hours in culture medium as described. Cell layers were fixed in 4% paraformaldehyde for 20
minutes and permeabilized in 0.05% Triton X-100 in PBS for 5 minutes. To visualize F-actin,
cells were incubated for 1 hour with Alexa Fluor 488-labeled phalloidin (Life Technologies).
At the end of the incubation period, cells were washed with PBS and incubated with Hoechst
33342 (Invitrogen) for 10 minutes. Finally, cultures were washed with 0.05% Triton X-100 in
PBS, mounted on glass coverslips with Fluoro-Gel with Tris buffer (Electron Microscopy Sci-
ences, Hatfield, PA) and imaged (Zeiss LSM 510 Non-Linear Optics with META Multiphoton
Excitation, Carl Zeiss Microscopy, Thornwood, NY).
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 4 / 18
To examine integrin expression, MSCs, HOBs, and chondrocytes were plated on copolymer
surfaces at a density of 10,000 cells/cm
2
on copolymer surfaces and cultured in the same cul-
ture medium as described above. Cells were fed 24 hours post-plating and every other day
thereafter. After 7 days, cells were incubated with fresh media for 12 hours and gene expression
for integrin subunits α1 (ITGA1), α2 (ITGA2), α5 (ITGA5), αv (ITGAV), β1 (ITGB1), and β3
(ITGB3) measured as described above.
Permanently silenced ITGB1 MSCs were generated to examine integrin-dependent MSC dif-
ferentiation on surfaces of varying stiffness. MSCs were transduced with shRNA lentiviral trans-
duction particles (SHCLNV_NM_002211, TRCN 0000029645, Mission1, Sigma-Aldrich) to
silence ITGB1. MSCs plated at 20,000 cells/cm
2
were cultured overnight. Cells were incubated
with particles (multiplicity of infection 5.0) overnight. Transduced cells were selected with cul-
ture media containing 0.25 μg/ml puromycin, yielding cells with 85% knockdown of mRNA.
Quantification of mRNA levels of SOX9 and RUNX2, cell number, alkaline phosphatase specific
activity, and secreted OCN and OPG were performed as described above and compared to
wild-type cells. In preliminary experiments, there was no difference found between wild-type
cells and cells containing empty vectors.
Statistics
Data are shown as mean +/- SEM of six independent cultures from a representative experi-
ment. All experiments were repeated. Using ANOVA with post-hoc Bonferroni’s modification
of Student’s t-test a value of P <0.05 was considered statistically significant.
Results
The similar chemical makeup of the monomers (Fig 1A) yielded networks (Fig 1B) with simi-
lar surface chemistries as indicated by their FTIR-ATR spectra–the position of the major
bonds (O-CH
3
, C = O, and C-O-C) did not shift between the different compositions (Fig 1C).
Although these networks had similar surface energy as evidenced by similar contact angles
(Fig 1D), their elastic moduli, measured in PBS at 37˚C, spanned multiple orders of magni-
tude, (18MA>29MA>40MA>72MA) (Fig 1E). The toughness of the MA-MMA copolymers
closely mimics the reported toughness of hard biological tissues including dentin and cortical
bone (Fig 2).
Stiffness affected the structural organization of cytoskeletal filaments. MSCs grown on
MA-MMA copolymer surfaces were longer on the less stiff surfaces, with multiple contact
points on the 40MA surface (Fig 3A–3D). Unlike MSCs grown on surfaces with lower stiffness,
F-actin appeared to be reduced in MSCs grown on 18MA surfaces (Fig 3A). HOBs were
noticeably smaller on the 18MA surface (Fig 3E) and more spread out on the less stiff surfaces
(Fig 3F–3H). Chondrocytes on 18MA, 29MA, and 40MA had similar morphology with long
extensions and few points of contact (Fig 3I–3K), and began to spread wider on the least stiff
surface (Fig 3L). However, there were no notable differences in the F-actin organization of
HOBs or chondrocytes on the substrates examined (Fig 3E–3L).
We then compared the effects of stiffness on osteoblast phenotype in mMSCs and mature
osteoblasts. MSCs and HOBs were grown without exogenous growth factors typically used to
induce differentiation (see Materials and Methods). mRNA levels of transcription factor
RUNX2 showed MSCs were more sensitive to stiffness than osteoblasts. RUNX2 mRNA in
MSCs increased as substrate stiffness decreased, an effect not present in OBs; however, the
highest RUNX2 levels were seen on 72MA surfaces in both cell types (Fig 4A and 4B). Cell
number was significantly higher on the 29MA and 72MA surfaces for MSCs whereas in HOBs
peak numbers were found on 72MA, followed by 29MA and 40MA (Fig 4C and 4D). Alkaline
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 5 / 18
phosphatase specific activity, an early marker for cells in the osteoblast lineage, was greatest in
MSCs grown on 40MA followed by 72MA while HOBs grown on these stiffnesses had lower
alkaline phosphatase activity compared to 18MA (Fig 4E and 4F). mRNAs for matrix proteins
osteocalcin and osteoprotegerin, associated with more mature osteoblasts, were similarly
affected. Peak levels of these proteins occurred in MSCs grown on 40MA (Fig 4G and 4I). Con-
versely, there was a decrease in osteocalcin expression in HOBs grown on the least stiff sub-
strates (Fig 4H), and osteoprotegerin mRNAs were lower in HOBs grown on all but 18MA
(Fig 4J).
Chondrogenic differentiation of MSCs and chondrogenic markers in chondrocytes were
responsive to substrate stiffness as well. MSCs and chondrocytes were grown for 7 days on the
polymer surfaces without exogenous growth factors to induce differentiation (see Materials
and Methods). SOX9, ACAN, COMP, and COL2 in MSCs increased as substrate stiffness
decreased and were greatest on 72MA (Fig 5A, 5C, 5E and 5G), whereas levels of SOX9 were
Fig 1. Characterization of MA-MMA networks. (A) Monomers used to create networks of increasing stiffness. (B)
Example of crosslinked network. (C) Representative FTIR-ATR spectra for each network indicating similar surface
chemistry for each. (D) Glass transition temperature (Tg) and contact angle of crosslinked networks. (E) DMA of
crosslinked networks (n = 3).
doi:10.1371/journal.pone.0170312.g001
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 6 / 18
equally high in chondrocytes grown on 40MA and 72MA (Fig 5B). ACAN levels in chondro-
cytes mimicked MSCs for 40MA and 72MA (Fig 5C and 5D), but contrary to MSCs the level
of COMP decreased on those same surfaces (Fig 5E and 5F). COL2 mRNA increased in chon-
drocytes as stiffness decreased, and the effect was similar to MSCs (Fig 5G and 5H). Alcian
blue staining to detect sulfated glycosaminoglycans was similar for both MSCs and chondro-
cytes, with an increase in staining in cells grown on both 40MA and 72MA surfaces with the
greatest intensity in cells grown on the least stiff surface (Fig 5I and 5J).
Integrin expression was surface-dependent in all three cell types after seven days in culture.
In MSCs, levels of integrin subunits ITGA1 and ITGA2 were higher on 29MA and 72MA sub-
strates than on 18MA, with peak levels occurring in MSCs grown on the 40MA (4.7 MPa stiff-
ness, Fig 6A and 6B). Surface stiffness altered mRNA levels of ITGA5 and ITGB1 in MSCs, with
higher levels in MSCs grown on the least stiff surfaces (Fig 6C and 6E). Levels of IGTAV were
significantly higher only on the 40MA and 72MA surfaces, peaking on the 40MA surfaces (Fig
6D). Conversely, ITGB3 increased as substrate stiffness increased (Fig 6F). In HOBs, ITGA1 lev-
els were higher on all but the stiffest surface, peaking on the 40MA (Fig 6G), but levels of ITGA2
were highest in cells grown on 29MA and lower in those grown on the copolymers with lower
stiffness (Fig 6H). HOBs grown on the surfaces with lower stiffness had higher levels of ITGA5
but lower levels of ITGAV (Fig 6I and 6J). The mRNA level for ITGB1 in HOBs was lowest on
the stiffest surface (Fig 6K). A decrease in the levels of ITGB3 could be seen for HOBs grown on
both 40MA and 72MA surfaces with the greatest decrease from those on the least stiff surface
(Fig 6L). Finally, in chondrocytes, ITGA1 mRNAs increased in cells grown on the 40MA surface
with higher levels in those grown on the 72MA surface (Fig 6M). The levels of ITGA2 were
lower in chondrocytes grown on surfaces with lower stiffness (Fig 6N). Chondrocytes had simi-
lar levels of ITGA5 on all surfaces but were significantly higher from those grown on the least
stiff surface (Fig 6O). The levels of both ITGAV and ITGB1 increased in chondrocytes grown on
Fig 2. Toughness vs. elastic modulus for load-bearing biological tissues (green) and MA-MMA
networks (black). Modified from data from [23].
doi:10.1371/journal.pone.0170312.g002
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 7 / 18
Fig 3. Cytoskeleton arrangement was altered by substrate stiffness. Representative staining of F-actin by phalloidin (green)
and nuclei by DAPI (blue) in human MSCs (A-D), HOBs (E-H), and chondrocytes (I-L) cultured on surfaces of varying stiffness.
(Scale bars: 100 μm for A,B,D; 50 μm for all others.)
doi:10.1371/journal.pone.0170312.g003
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 8 / 18
Fig 4. Osteoblastic differentiation on MA-MMA networks. (A-B) mRNA levels for osteoblast-specific
marker RUNX2. (C-J) MSC and HOB response to substrate stiffness seen in cell number and osteogenic
protein levels. *P<0.05 vs. 18 MA, #P<0.05 vs. 29 MA, $P<0.05 vs. 40 MA.
doi:10.1371/journal.pone.0170312.g004
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 9 / 18
Fig 5. Chondrogenic differentiation on MA-MMA networks. Levels of chondrogenic mRNA (A-H) and
quantification of proteoglycan staining in MSCs and chondrocytes (I-J) cultured on surfaces of varying
stiffness. *P<0.05 vs. 18 MA, #P<0.05 vs. 29 MA, $P<0.05 vs. 40 MA.
doi:10.1371/journal.pone.0170312.g005
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 10 / 18
Fig 6. Integrin expression is stiffness- and cell-type- dependent. Comparison of integrin mRNA levels in
MSCs, OBs, and chondrocytes cultured on surfaces of varying stiffness. *P<0.05 vs. 18 MA, #P<0.05 vs. 29 MA,
$P<0.05 vs. 40 MA.
doi:10.1371/journal.pone.0170312.g006
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 11 / 18
surfaces with lower stiffness (Fig 6P and 6Q) and, as in MSCs, a similar stiffness-dependent
decrease in the level of ITGB3 was observed (Fig 6R).
Because MSCs tended to be the most sensitive to the varied stiffness, we wanted to deter-
mine how silencing integrin β1 (ITGB1) in these cells would modulate this response. Silencing
ITGB1 abolished the stiffness-dependent expression of mRNA for transcription factors for
chondrocytes (SOX9) and osteoblasts (RUNX2) (Fig 7A and 7B). Silencing ITGB1 also abol-
ished the increase in cell number on decreasingly stiff surfaces seen in WT MSCs (Fig 7C).
shITGB1-MSCs had lower alkaline phosphatase specific activity compared to their wild-type
counterparts, and the levels decreased with decreasing stiffness (Fig 7D). Osteocalcin levels
that were highest on 40MA in WT MSCs were lower in the silenced cells on all stiffness and
lowest on 72MA surfaces (Fig 7E). Finally, the increase of osteoprotegerin in wild-type cells
grown on 40MA was also abolished in shITGB1-MSCs (Fig 7F).
Discussion
Multipotent stem cells from various sources have previously been shown to differentiate in
response to varying topographies and stiffness [5,6,3335]. In most cases, this differentiation
has been enhanced with different induction media resulting in a very complex process that
potentially masks effects of surface features, chemistry, or stiffness. In our study, we eliminated
all but one of these variables, stiffness, in order to tease out its effects on MSC differentiation
along two lineage fates: chondrogenic and osteogenic. Our results show that stiffness alone can
Fig 7. Integrin-dependent osteoblast differentiation. Levels of chondrogenic (A) and osteoblastic (B)
mRNA. (C-F) Cell number and osteogenic protein levels in wild type human MSCs (WT) and silenced integrin
β1 MSCs (shITGB1) cultured on surfaces of varying stiffness. *P<0.05 vs. 18 MA, #P<0.05 vs. wild type.
doi:10.1371/journal.pone.0170312.g007
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 12 / 18
direct differentiation and that different stiffness favors expression of a cartilage cell phenotype
v. expression of an osteoblast phenotype, but no one stiffness produces an exclusive outcome.
We did not observe significant morphological changes in MSCs, HOBs, or chondrocytes on
our polymer networks as others have demonstrated [6,36] although we did see some stiffness-
dependent cytoskeletal arrangement. Significant changes in expression of differentiation
markers do not necessarily correlate with outward changes in MSC morphology over the short
time course of our study [37]. Similarly, we did not observe morphological changes in the
chondrocyte and osteoblast cultures, although gene expression for differentiation markers was
affected.
Gene expression in the cultures did vary with cell type and with substrate stiffness. MSCs
exhibited increasing RUNX2 expression with decreasing stiffness, whereas expression in the
committed HOB cells was less sensitive to substrate. Levels of this factor are correlated with
osteoblastic differentiation of MSCs [3840], suggesting that less stiff substrates induced osteo-
blastic differentiation. Whether this reflects in vivo differentiation on osteoclast resorbed bone
surfaces, which are primarily collagen and non-collagenous proteins like osteopontin rather
than stiffer fully mineralized bone [41,42], isn’t clear. The lower RUNX2 in HOBs on certain
substrates suggests that the cells are less active osteoblasts [43,44] than on other substrates.
We had expected MSCs to behave more like HOBs on the stiffer surface and more like
chondrocytes on the less stiff surfaces but MSC responses were observed on the less stiff poly-
mers (40MA and 72MA) for both lineages, indicating the culture was a mixed population
exhibiting both osteoblastic and chondrogenic markers: more osteoblastic markers on the
slightly less soft surface and more chondrogenic markers on the softest surface. The observa-
tion that MSCs had the highest osteoblast response on the next to least stiff (40MA) networks
and the highest chondrogenic response on the least stiff (72MA) networks is an indicator that
there exists an optimal substrate stiffness to promote osteoblast differentiation and that it is
not simply ‘the harder, the better.’
HOB expression levels of osteoblastic genes increased only on the stiffer surfaces; on the
softer surfaces, they not only did not have this increase but also may have begun to dedifferenti-
ate. This suggests that maintenance of an osteoblastic phenotype may require a stiffer microen-
vironment typical of mineralized bone. To achieve a stable osteoblast phenotype in MSCs
grown on TCPS requires extensive time in culture to develop multi-layered nodules and
requires the use of media supplements for as long as three weeks to support mineral formation
within the nodules [45]. Our study did not examine the long-term effects of MSCs on the softer
72MA substrate to determine if stiffness alone would support stable osteoblastic differentiation
and matrix mineralization. Continued culture on the softer substrate in the absence of media
supplements could have an inhibitory effect on downstream osteoblastic differentiation.
The differentiated HOBs behaved differently than the MSCs on the varied stiffness, suggest-
ing that as differentiation progresses, substrate stiffness continues to influence cell maturation.
Metal and ceramic implants generally have moduli much higher than native bone [23,4648];
any polymer scaffold or bone substitute must consider stiffness as a critical factor. Most pub-
lished work examining cell response to substrate stiffness use polymers such as hydrogels,
which have moduli orders of magnitude low er than 100 kPa, far below biological tissues such
as dental tissue or cortical bone, which are at or near common implant sites [19,33,35,4952].
In contrast, chondrocytes, which exist in a hydrogel environment in vivo [53] behaved very
similarly to MSCs. It is important to note that this study was performed using auricular chondro-
cytes, which may have different responses to substrate stiffness than growth plate or articular
chondrocytes. Several reports have demonstrated that auricular chondrocytes proliferate well in
culture, maintain their phenotype, and are suitable for tissue-engineered constructs [2629].
Moreover, other reports have demonstrated that they are able to heal defects in articular cartilage
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 13 / 18
to a similar extent as articular chondrocytes [30]. Given these applications in tissue engineering,
we chose to use them as a cell source for this experiment.
Our results indicate that differential expression of integrins in response to surface stiffness
plays a crucial role in determining cell response, and that integrin signaling controls MSC differ-
entiation. Because integrins are a cell’s primary response to substrate stiffness due to ligand bind-
ing [5456], and change according to differentiation [57], it follows that depending on a cell’s
phenotype it would have more or less sensitivity to substrate stiffness. Expression of ITGA1,
ITGA2, and ITGA5 in particular in MSCs was much more sensitive to stiffness than in either
osteoblasts or chondrocytes. Others have reported a similar increase in ITGA5 in murine fibro-
blasts though no substrates stiffer than 55 kPa were examined [54]. Sanz-Ramos et al. examined
integrin expression in rat chondrocytes on surfaces of 2–20 Pa stiffness under normoxia and
hypoxia conditions and saw a decrease in ITGA2 and ITGAV, but no differences in ITGA1,
ITGB1, or ITGB3 under normoxia [56], whereas we did. The softness of the substrates examined
could account for the difference in our data compared to theirs, as changes in ITGB3 expression
were not seen until MSCs were grown on stiffer substrates. The silencing of ITGB1 completely
abolished this sensitivity at one week, faster than the 2–3 weeks reported previously [55]. The
abolition of the SOX9 response in MSCs by ITGB1 silencing is likely due to cellular stiffness and
diffusion changes, as others have seen increased activation of ITGB1 on softer substrates [18,58].
Conclusion
Our results show that stiffness can direct the fate of MSCs and suggest that over a very small
range, induce bone or cartilage formation–or both, such as our 40MA networks, which showed
an enhancement of bone and cartilage markers. Once cells commit to an osteoblast lineage,
stiffness has an entirely different effect, suggesting that softer substrates could halt further oste-
oblast maturation. We were able to enhance chondrocyte markers in mature chondrocytes
while the same networks inhibited osteoblast markers in mature osteoblasts. In addition we
show that multiple integrins and in particular integrin β1 play a vital role in MSC sensitivity to
stiffness. Understanding the importance of this mechanical property unlocks its useful poten-
tial for exploitation to control cell fate.
Supporting Information
S1 Fig. Expression of cartilage cell phenotype. Levels of chondrogenic mRNA cultured on
TCPS for 7 days. #P<0.05 vs. MSCs.
(TIF)
S1 Table. Primer sequences used for Real-time PCR analysis of gene expression.
(DOC)
Acknowledgments
We would like to thank Children’s Healthcare of Atlanta for their contributions to this
research.
Author Contributions
Conceptualization: RON KG BDB ZS KS.
Formal analysis: SLH BDB ZS.
Funding acquisition: KG BDB.
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 14 / 18
Investigation: RON EML SLH KS MD.
Methodology: ZS SLH MD KS.
Project administration: RON SLH BDB ZS.
Resources: JKW.
Supervision: RON KG BDB ZS.
Validation: ZS SLH MD.
Visualization: SLH EML.
Writing – original draft: RON EML BDB.
Writing – review & editing: RON EML KS SLH MD JKW KG BDB ZS.
References
1. Schwarz F, Wieland M, Schwartz Z, Zhao G, Rupp F, Geis-Gerstorfer J, et al. Potential of chemically
modified hydrophilic surface characteristics to support tissue integration of titanium dental implants.
2009; 88: 544–57. doi: 10.1002/jbm.b.31233 PMID: 18837448
2. Calvo-Guirado JL, Gomez Moreno G, Aguilar-Salvatierra A, Mate Sanchez de Val JE, Abboud M, Nem-
covsky CE. Bone remodeling at implants with different configurations and placed immediately at differ-
ent depth into extraction sockets. Experimental study in dogs. Clin Oral Implants Res. 2015; 26: 507–
515. doi: 10.1111/clr.12433 PMID: 24888507
3. Sartoretto SC, Alves AT, Resende RF, Calasans-Maia J, Granjeiro JM, Calasans-Maia MD. Early
osseointegration driven by the surface chemistry and wettability of dental implants. J Appl Oral Sci.
2015; 23: 279–287. doi: 10.1590/1678-775720140483 PMID: 26221922
4. Lee HJ, Yang IH, Kim SK, Yeo IS, Kwon TK. In vivo comparison between the effects of chemically modi-
fied hydrophilic and anodically oxidized titanium surfaces on initial bone healing. J Periodontal Implant
Sci. 2015; 45: 94–100. doi: 10.5051/jpis.2015.45.3.94 PMID: 26131369
5. Olivares-Navarrete R, Hyzy SL, Hutton DL, Erdman CP, Wieland M, Boyan BD, et al. Direct and indirect
effects of microstructured titanium substrates on the induction of mesenchymal stem cell differentiation
towards the osteoblast lineage. Biomaterials. 2010; 31: 2728–2735. doi: 10.1016/j.biomaterials.2009.
12.029 PMID: 20053436
6. Engler AJ, Sen S, Sweeney HL, Discher DE. Matrix elasticity directs stem cell lineage specification.
2006; 126: 677–89.
7. Seib FP, Prewitz M, Werner C, Bornhauser M. Matrix elasticity regulates the secretory profile of human
bone marrow-derived multipotent mesenchymal stromal cells (MSCs). Biochem Biophys Res Commun.
2009; 389: 663–667. doi: 10.1016/j.bbrc.2009.09.051 PMID: 19766096
8. Xue R, Li JY, Yeh Y, Yang L, Chien S. Effects of matrix elasticity and cell density on human mesenchy-
mal stem cells differentiation. J Orthop Res. 2013; 31: 1360–1365. doi: 10.1002/jor.22374 PMID:
23606500
9. Her GJ, Wu HC, Chen MH, Chen MY, Chang SC, Wang TW. Control of three-dimensional substrate
stiffness to manipulate mesenchymal stem cell fate toward neuronal or glial lineages. Acta Biomater.
2013; 9: 5170–5180. doi: 10.1016/j.actbio.2012.10.012 PMID: 23079022
10. Angele P, Muller R, Schumann D, Englert C, Zellner J, Johnstone B, et al. Characterization of esterified
hyaluronan-gelatin polymer composites suitable for chondrogenic differentiation of mesenchymal stem
cells. J Biomed Mater Res A. 2009; 91: 416–427. doi: 10.1002/jbm.a.32236 PMID: 18985778
11. Marletta G, Ciapetti G, Satriano C, Perut F, Salerno M, Baldini N. Improved osteogenic differentiation of
human marrow stromal cells cultured on ion-induced chemically structured poly-epsilon-caprolactone.
Biomaterials. 2007; 28: 1132–1140. S0142-9612(06)00918-5 [pii]. doi: 10.1016/j.biomaterials.2006.10.
027 PMID: 17118444
12. Munoz-Pinto DJ, Jimenez-Vergara AC, Hou Y, Hayenga HN, Rivas A, Grunlan M, et al. Osteogenic
potential of poly(ethylene glycol)-poly(dimethylsiloxane) hybrid hydrogels. Tissue Eng Part A. 2012; 18:
1710–1719. doi: 10.1089/ten.TEA.2011.0348 PMID: 22519299
13. Wang JR, Ahmed SF, Gadegaard N, Meek RM, Dalby MJ, Yarwood SJ. Nanotopology potentiates
growth hormone signalling and osteogenesis of mesenchymal stem cells. Growth Horm IGF Res. 2014;
24: 245–250. doi: 10.1016/j.ghir.2014.10.003 PMID: 25466909
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 15 / 18
14. Reilly GC, Engler AJ. Intrinsic extracellular matrix properties regulate stem cell differentiation. J Bio-
mech. 2010; 43: 55–62. doi: 10.1016/j.jbiomech.2009.09.009 PMID: 19800626
15. Viswanathan P, Ondeck MG, Chirasatitsin S, Ngamkham K, Reilly GC, Engler AJ, et al. 3D surface
topology guides stem cell adhesion and differentiation. Biomaterials. 2015; 52: 140–147. doi: 10.1016/j.
biomaterials.2015.01.034 PMID: 25818420
16. Keselowsky BG, Wang L, Schwartz Z, Garcia AJ, Boyan BD. Integrin α5 controls osteoblastic prolifera-
tion and differentiation responses to titanium substrates presenting different roughness characteristics
in a roughness independent manner. 2007; 80A: 700–710.
17. Olivares-Navarrete R, Raz P, Zhao G, Chen J, Wieland M, Cochran DL, et al. Integrin alpha2beta1
plays a critical role in osteoblast response to micron-scale surface structure and surface energy of tita-
nium substrates. 2008; 105: 15767–72. 0805420105 [pii]. doi: 10.1073/pnas.0805420105 PMID:
18843104
18. Du J, Chen X, Liang X, Zhang G, Xu J, He L, et al. Integrin activation and internalization on soft ECM as
a mechanism of induction of stem cell differentiation by ECM elasticity. Proc Natl Acad Sci U S A. 2011;
108: 9466–9471. doi: 10.1073/pnas.1106467108 PMID: 21593411
19. Steward AJ, Wagner DR, Kelly DJ. The pericellular environment regulates cytoskeletal development
and the differentiation of mesenchymal stem cells and determines their response to hydrostatic pres-
sure. Eur Cell Mater. 2013; 25: 167–178. vol025a12 [pii]. PMID: 23389751
20. Enomoto M, Leboy PS, Menko AS, Boettiger D. Beta 1 integrins mediate chondrocyte interaction with
type I collagen, type II collagen, and fibronectin. Exp Cell Res. 1993; 205: 276–285. S0014-4827(83)
71087-6 [pii]. doi: 10.1006/excr.1993.1087 PMID: 8387015
21. Loeser RF, Carlson CS, McGee MP. Expression of beta 1 integrins by cultured articular chondrocytes
and in osteoarthritic cartilage. Exp Cell Res. 1995; 217: 248–257. S0014-4827(85)71084-1 [pii]. doi: 10.
1006/excr.1995.1084 PMID: 7535235
22. Nava MM, Raimondi MT, Pietrabissa R. Controlling self-renewal and differentiation of stem cells via
mechanical cues. J Biomed Biotechnol. 2012; 2012: 797410. doi: 10.1155/2012/797410 PMID:
23091358
23. Smith KE, Hyzy SL, Sunwoo M, Gall KA, Schwartz Z, Boyan BD. The dependence of MG63 osteoblast
responses to (meth)acrylate-based networks on chemical structure and stiffness. 2010; 31: 6131–41.
S0142-9612(10)00549-1 [pii]. doi: 10.1016/j.biomaterials.2010.04.033 PMID: 20510445
24. Smith KE, Trusty P, Wan B, Gall K. Long-term toughness of photopolymerizable (meth)acrylate net-
works in aqueous environments. Acta Biomater. 2011; 7: 558–567. doi: 10.1016/j.actbio.2010.09.001
PMID: 20828638
25. Engel FE, Khare AG, Boyan BD. Phenotypic changes of rabbit mandibular condylar cartilage cells in
culture. 1990; 69: 1753–8. PMID: 2229613
26. Elisseeff J. Injectable cartilage tissue engineering. Expert Opin Biol Ther. 2004; 4: 1849–1859.
EBT041201 [pii]. doi: 10.1517/14712598.4.12.1849 PMID: 15571448
27. Xu JW, Zaporojan V, Peretti GM, Roses RE, Morse KB, Roy AK, et al. Injectable tissue-engineered car-
tilage with different chondrocyte sources. Plast Reconstr Surg. 2004; 113: 1361–1371. 00006534-
200404150-00007 [pii]. PMID: 15060348
28. Chung C, Erickson IE, Mauck RL, Burdick JA. Differential behavior of auricular and articular chondro-
cytes in hyaluronic acid hydrogels. Tissue Eng Part A. 2008; 14: 1121–1131. doi: 10.1089/tea.2007.
0291 PMID: 18407752
29. Lohan A, Marzahn U, El Sayed K, Haisch A, Kohl B, Muller RD, et al. In vitro and in vivo neo-cartilage
formation by heterotopic chondrocytes seeded on PGA scaffolds. Histochem CellBiol. 2011; 136: 57–
69. doi: 10.1007/s00418-011-0822-2 PMID: 21656225
30. Lohan A, Marzahn U, El Sayed K, Haisch A, Muller RD, Kohl B, et al. Osteochondral articular defect
repair using auricle-derived autologous chondrocytes in a rabbit model. Ann Anat. 2014; 196: 317–326.
doi: 10.1016/j.aanat.2014.03.002 PMID: 24812031
31. Olivares-Navarrete R, Raines AL, Hyzy SL, Park JH, Hutton DL, Cochran DL, et al. Osteoblast matura-
tion and new bone formation in response to titanium implant surface features are reduced with age. J
Bone Miner Res. 2012; 27: 1773–1783. doi: 10.1002/jbmr.1628 PMID: 22492532
32. Kitaoka E, Satomura K, Hayashi E, Yamanouchi K, Tobiume S, Kume K, et al. Establishment and char-
acterization of chondrocyte cell lines from the costal cartilage of SV40 large T antigen transgenic mice.
J Cell Biochem. 2001; 81: 571–582. PMID: 11329612
33. Park JS, Chu JS, Tsou AD, Diop R, Tang Z, Wang A, et al. The effect of matrix stiffness on the differenti-
ation of mesenchymal stem cells in response to TGF-beta. Biomaterials. 2011; 32: 3921–3930. doi: 10.
1016/j.biomaterials.2011.02.019 PMID: 21397942
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 16 / 18
34. Bayati V, Altomare L, Tanzi MC, Fare S. Adipose-derived stem cells could sense the nano-scale cues
as myogenic-differentiating factors. J Mater Sci Mater Med. 2013; 24: 2439–2447. doi: 10.1007/
s10856-013-4983-5 PMID: 23793565
35. Li X, Huang Y, Zheng L, Liu H, Niu X, Huang J, et al. Effect of substrate stiffness on the functions of rat
bone marrow and adipose tissue derived mesenchymal stem cells in vitro. J Biomed Mater Res A.
2014; 102: 1092–1101. doi: 10.1002/jbm.a.34774 PMID: 23630099
36. Genes NG, Rowley JA, Mooney DJ, Bonassar LJ. Effect of substrate mechanics on chondrocyte adhe-
sion to modified alginate surfaces. Arch Biochem Biophys. 2004; 422: 161–167. doi: 10.1016/j.abb.
2003.11.023 PMID: 14759603
37. Huebsch N, Arany PR, Mao AS, Shvartsman D, Ali OA, Bencherif SA, et al. Harnessing traction-medi-
ated manipulation of the cell/matrix interface to control stem-cell fate. Nat Mater. 2010; 9: 518–526. doi:
10.1038/nmat2732 PMID: 20418863
38. Sun J, Zhou H, Deng Y, Zhang Y, Gu P, Ge S, et al. Conditioned medium from bone marrow mesenchy-
mal stem cells transiently retards osteoblast differentiation by downregulating runx2. Cells Tissues
Organs. 2012; 196: 510–522. doi: 10.1159/000339245 PMID: 22906827
39. Lee KS, Kim HJ, Li QL, Chi XZ, Ueta C, Komori T, et al. Runx2 is a common target of transforming
growth factor beta1 and bone morphogenetic protein 2, and cooperation between Runx2 and Smad5
induces osteoblast-specific gene expression in the pluripotent mesenchymal precursor cell line C2C12.
Mol Cell Biol. 2000; 20: 8783–8792. PMID: 11073979
40. Dieudonne FX, Severe N, Biosse-Duplan M, Weng JJ, Su Y, Marie PJ. Promotion of osteoblast differen-
tiation in mesenchymal cells through Cbl-mediated control of STAT5 activity. Stem Cells. 2013; 31:
1340–1349. doi: 10.1002/stem.1380 PMID: 23533197
41. Alexopoulos LG, Williams GM, Upton ML, Setton LA, Guilak F. Osteoarthritic changes in the biphasic
mechanical properties of the chondrocyte pericellular matrix in articular cartilage. J Biomech. 2005; 38:
509–517. S0021929004002039 [pii]. doi: 10.1016/j.jbiomech.2004.04.012 PMID: 15652549
42. Willems NM, Langenbach GE, Stoop R, den Toonder JM, Mulder L, Zentner A, et al. Higher number of
pentosidine cross-links induced by ribose does not alter tissue stiffness of cancellous bone. Mater Sci
Eng C Mater Biol Appl. 2014; 42: 15–21. doi: 10.1016/j.msec.2014.05.006 PMID: 25063086
43. Adhami MD, Rashid H, Chen H, Javed A. Runx2 activity in committed osteoblasts is not essential for
embryonic skeletogenesis. Connect Tissue Res. 2014; 55 Suppl 1: 102–106.
44. Bruderer M, Richards RG, Alini M, Stoddart MJ. Role and regulation of RUNX2 in osteogenesis. Eur
Cell Mater. 2014; 28: 269–286. vol028a19 [pii]. PMID: 25340806
45. Schwartz Z, Schlader DL, Swain LD, Boyan BD. Direct effects of 1,25-dihydroxyvitamin D3 and 24,25-
dihydroxyvitamin D3 on growth zone and resting zone chondrocyte membrane alkaline phosphatase
and phospholipase-A2 specific activities. 1988; 123: 2878–84. doi: 10.1210/endo-123-6-2878 PMID:
3264240
46. Rho JY, Tsui TY, Pharr GM. Elastic properties of human cortical and trabecular lamellar bone measured
by nanoindentation. Biomaterials. 1997; 18: 1325–1330. S0142961297000732 [pii]. PMID: 9363331
47. Guillemot F. Recent advances in the design of titanium alloys for orthopedic applications. Expert Rev
Med Devices. 2005; 2: 741–748. doi: 10.1586/17434440.2.6.741 PMID: 16293101
48. Thompson ID, Hench LL. Mechanical properties of bioactive glasses, glass-ceramics and composites.
Proc Inst Mech Eng H. 1998; 212: 127–136. PMID: 9612004
49. Keogh MB, O’Brien FJ, Daly JS. Substrate stiffness and contractile behaviour modulate the functional
maturation of osteoblasts on a collagen-GAG scaffold. Acta Biomater. 2010; 6: 4305–4313. doi: 10.
1016/j.actbio.2010.06.001 PMID: 20570642
50. Chandler EM, Berglund CM, Lee JS, Polacheck WJ, Gleghorn JP, Kirby BJ, et al. Stiffness of photo-
crosslinked RGD-alginate gels regulates adipose progenitor cell behavior. Biotechnol Bioeng. 2011;
108: 1683–1692. doi: 10.1002/bit.23079 PMID: 21328324
51. Murphy CM, Matsiko A, Haugh MG, Gleeson JP, O’Brien FJ. Mesenchymal stem cell fate is regulated
by the composition and mechanical properties of collagen-glycosaminoglycan scaffolds. J Mech Behav
Biomed Mater. 2012; 11: 53–62. doi: 10.1016/j.jmbbm.2011.11.009 PMID: 22658154
52. Witkowska-Zimny M, Walenko K, Walkiewicz AE, Pojda Z, Przybylski J, Lewandowska-Szumiel M.
Effect of substrate stiffness on differentiation of umbilical cord stem cells. Acta Biochim Pol. 2012; 59:
261–264. 2011_204 [pii]. PMID: 22577624
53. Darling EM, Wilusz RE, Bolognesi MP, Zauscher S, Guilak F. Spatial mapping of the biomechanical
properties of the pericellular matrix of articular cartilage measured in situ via atomic force microscopy.
Biophys J. 2010; 98: 2848–2856. doi: 10.1016/j.bpj.2010.03.037 PMID: 20550897
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 17 / 18
54. Yeung T, Georges PC, Flanagan LA, Marg B, Ortiz M, Funaki M, et al. Effects of substrate stiffness on
cell morphology, cytoskeletal structure, and adhesion. Cell Motil Cytoskeleton. 2005; 60: 24–34. doi:
10.1002/cm.20041 PMID: 15573414
55. Shih YR, Tseng KF, Lai HY, Lin CH, Lee OK. Matrix stiffness regulation of integrin-mediated mechano-
transduction during osteogenic differentiation of human mesenchymal stem cells. J Bone Miner Res.
2011; 26: 730–738. doi: 10.1002/jbmr.278 PMID: 20939067
56. Sanz-Ramos P, Mora G, Ripalda P, Vicente-Pascual M, Izal-Azcarate I. Identification of signalling path-
ways triggered by changes in the mechanical environment in rat chondrocytes. Osteoarthritis Cartilage.
2012; 20: 931–939. doi: 10.1016/j.joca.2012.04.022 PMID: 22609478
57. Frith JE, Mills RJ, Hudson JE, Cooper-White JJ. Tailored integrin-extracellular matrix interactions to
direct human mesenchymal stem cell differentiation. Stem Cells Dev. 2012; 21: 2442–2456. doi: 10.
1089/scd.2011.0615 PMID: 22455378
58. Bougault C, Cueru L, Bariller J, Malbouyres M, Paumier A, Aszodi A, et al. Alteration of cartilage
mechanical properties in absence of beta1 integrins revealed by rheometry and FRAP analyses. J Bio-
mech. 2013; 46: 1633–1640. doi: 10.1016/j.jbiomech.2013.04.013 PMID: 23692868
Substrate Stiffness Controls Osteoblastic and Chondrocytic Differentiation of Mesenchymal Stem Cells
PLOS ONE | DOI:10.1371/journal.pone.0170312 January 17, 2017 18 / 18
... It is not only CM containing secreted factors such as cytokines that can influence MSC osteoblast differentiation and osteoblast function. Upon culturing MSC and osteoblasts on stiff surfaces in comparison to limper surfaces, increased MSC osteogenic differentiation can be seen upon analysing RUNX2, Col1a1, OCN gene expression and calcium deposition [544][545][546][547]. Increased MSC and osteoblast mineral deposition on stiffer matrices than softer matrices is observed whilst the maintenance of spindle-shaped morphology remains and in vitro pro-bone formation characteristics could be observed [545,546,548,549]. ...
... This gene in particular encodes a nuclear protein with a Runt DNA-Binding domain which is key for osteoblast differentiation [664]. [541,543,670,671,544,547,548,587,[666][667][668][669]. However, despite the wide number of genes available for this evaluation, the most common ones used are RUNX2, Col1a1, OCN and OPN. ...
... However, the reducing capacity of each sample is altered by cell number, therefore in this study, PrestoBlue would not be applicable without the prior knowledge that CM would not affect cell number. The MTT based assay works using a similar method and is also limited by the probability of CM affecting cell number, which the literature has shown, does occur more often than not [541,543,544,547,550,552,587,669,670]. Through the use of the Trypan Blue method, not only was cell viability assessed, but also cell number could be derived. ...
... Receptor Activated of Nuclear Factor Kappa-B (RANK) is an osteoclast receptor and its ligand, It is not only CM containing secreted factors such as cytokines that can influence MSC osteoblast differentiation and osteoblast function. Upon culturing MSC and osteoblasts on stiff surfaces in comparison to limper surfaces, increased MSC osteogenic differentiation can be seen upon analysing RUNX2, Col1a1, OCN gene expression and calcium deposition [544][545][546][547]. Increased MSC and osteoblast mineral deposition on stiffer matrices than softer matrices is observed whilst the maintenance of spindle-shaped morphology remains and in vitro pro-bone formation characteristics could be observed [545,546,548,549]. ...
... (osteoprotegerin) can be used to evaluate MSC differentiation down osteogenic lineages [541,543,670,671,544,547,548,587,[666][667][668][669]. However, despite the wide number of genes available for this evaluation, the most common ones used are RUNX2, Col1a1, OCN and OPN. ...
... However, the reducing capacity of each sample is altered by cell number, therefore in this study, PrestoBlue would not be applicable without the prior knowledge that CM would not affect cell number. The MTT based assay works using a similar method and is also limited by the probability of CM affecting cell number, which the literature has shown, does occur more often than not [541,543,544,547,550,552,587,669,670]. Through the use of the Trypan Blue method, not only was cell viability assessed, but also cell number could be derived. ...
Thesis
One of the greatest requirements of modern medicine is the ability to treat patients suffering from osteoarthritis (OA) and bone fractures. Currently, there is no long-term therapy for OA; symptoms can be managed with anti-inflammatories and analgesics until they worsen to the extent that the damage becomes debilitating, and joint arthroplasty, is necessitated. However, these replacements are not perfect; firstly, there is the need for surgery and secondly, if the patient is young, the prosthetic can deteriorate, engendering further surgery. Bone fractures are regularly seen in orthopaedic clinics and are commonly repaired using fixation techniques or biomaterials. After any intervention, the fracture site can remain compromised, potentially engendering re-fracture and/or further surgical involvement. Regenerative strategies for both OA and bone fracture aim to alleviate pain, whilst maintaining or restoring damaged tissues to healthy states. Mesenchymal stem/stromal cells (MSC) are thought to facilitate tissue repair via either progenitor or secreaome functions. BM-MSC have, in previous work, been investigated as a therapy for OA via either their direct application or through their secreted Extracellular Vesicles (EV). In this study, MSC have been successfully isolated from bone marrow, and from these isolated cells, EV have been captured and characterised. The isolated EV have been shown to be readily internalised by chondrocytes and, in order to determine the method of EV internalisation by chondrocytes, in vitro drug inhibition studies were performed on labelled EV. Via inhibition of the caveolin dependent endocytosis pathway, EV uptake was prevented, thus indicating that this method of endocytosis is the method of EV internalisation. In regenerative medicine for knee OA, it is likely that MSC and EV would be injected into the knee. In order to determine if the MSC and EV would reside in the joint, both were labelled with gold nanostars and Supra Magnetic Iron Oxide Nanoparticles (SPION). These labelled cells and EV were then injected into a sheep stifle 1 week post creation of an OA model (meniscal transection model). These labelled cells and EV could then be seen within the knee for up to 4 weeks post injection, as ascertained via Magnetic Resonance Imaging (MRI) and MultiSpectral Optoacoustic Tomography (MSOT). Upon evaluating the regenerative effects of the MSC and EV, no difference in cartilage damage could be seen. During bone fracture, MSC and osteoblasts are recruited to the site of injury. Bioglasses have been used previously as a material to improve bone repair through the release of ions and conditioning the local environment. Our work has shown that conditioned media from bioglasses can influence both MSC and osteoblasts to augment the bone repair process. Through screening bioglasses on MSC and osteoblasts, the potential for bioglasses to alter MSC derived EV to promote osteogenesis has been shown. As a conclusion, this study has shown that the BM-MSC are a source of EV, and that both the MSC and EV can potentially be used in a musculoskeletal scenario of regenerative medicine.
... Recent studies have shown that directing stem cell fate could be achieved by finely tuning the mechanical environment. [18][19][20][21][22][23][24] In particular, stiffness, defined as the extent to which a material resists deformation in response to an applied force, has been shown to influence cell phenotype and to play a pivotal role in tissue development, homoeostasis and disease. [25][26][27][28][29][30] On stiff substrates, MSCs have been shown to acquire an elongated spindle shape triggering the activation of extracellular signal-regulated kinase 1 (ERK1) and ERK2 thus promoting osteogenic differentiation. ...
Article
Full-text available
Mesenchymal stem cells (MSCs) hold great promise for the treatment of cartilage related injuries. However, selectively promoting stem cell differentiation in vivo is still challenging. Chondrogenic differentiation of MSCs usually requires the use of growth factors that lead to the overexpression of hypertrophic markers. In this study, for the first time the effect of stiffness on MSC differentiation has been tested without the use of growth factors. Three-dimensional collagen and alginate scaffolds were developed and characterised. Stiffness significantly affected gene expression and ECM deposition. While, all hydrogels supported chondrogenic differentiation and allowed deposition of collagen type II and aggrecan, the 5.75 kPa hydrogel showed limited production of collagen type I compared to the other two formulations. These findings demonstrated for the first time that stiffness can guide MSCs differentiation without the use of growth factors within a tissue engineering scaffold suitable for the treatment of cartilage defects.
... Alternatively, some of them play critical roles in stem cell recruitment, proliferation, and differentiation, which enhances the repair process and osseointegration of the implant with the native tissue [48]. Thirdly, mechanical properties such as stiffness affect cell migration, proliferation, and differentiation [49]. This study demonstrated that even though different surfaces did not directly bring about the significant changes in cell attachment and proliferation, they had different protein adsorptions and induced pro-inflammatory TNF-α and IL-6 gene expressions at different levels. ...
Article
Full-text available
The surfaces of 3D printed titanium prostheses have major impacts on the clinical performance of the prostheses. To investigate the surface effects of the products generated by 3D printed titanium on osseointegration, six surface types of titanium discs produced by the direct metal laser sintering (DMLS) and electron beam melting (EBM) methods, with two sizes of titanium particles and post-printing acid etching, were used to examine the surface topography and to explore the protein adsorption, pro-inflammatory cytokine gene expressions, and MC3T3-E1 cell adhesion, proliferation, and differentiation. The EBM-printed disc showed a stripy and smooth surface without evidence of the particles used, while the DMLS surface contained many particles. After acid etching, small particles on the DMLS surface were removed, whereas the large particles were left. Moreover, distinct proteins with low molecular weights were attached to the 3D printed titanium discs but not to the pre-printing titanium particles. The small titanium particles stimulated the highest TNF-α and IL-6 gene expressions at 24 h. The alizarin red content and osteocalcin gene expression at day 21 were the highest in the groups of acid-etched discs printed by DMLS with the small particles and by EBM. Therefore, the acid-treated surfaces without particles favor osteogenic differentiation. The surface design of 3D printed titanium prostheses should be based on their clinical applications.
... In most cases, different soluble factors, especially members of transforming growth factor beta (TGF-β) superfamily, were added to assist chondrogenesis throughout the whole culturing cycle with different matrix stiffness (Olivares-Navarrete et al. 2017;Srinivasan et al. 2018;Wu et al. 2017). In fact, it is pointed out that there exists a biphasic expression of TGF-β in chondrogenic differentiation of MSCs, which means TGF-β is firstly upregulated then downregulated (Ruiz et al. 2019). ...
Article
Full-text available
Articular cartilage is one of the most important weight-bearing components in human body, thus the chondrogenesis of stem cells is reactive to many intracellular and extracellular mechanical signals. As a unique physical cue, matrix stiffness plays an integral role in commitment of stem cell fate. However, when examining the downstream effects of matrix stiffness, most studies used different soluble factors to assist physical inducing process, which may mask the chondrogenic effects of matrix stiffness. Here we fabricated polyacrylamide (PAAm) hydrogels with gradient stiffness to unravel the role of matrix stiffness in chondrogenic process of mesenchymal stem cells (MSCs), with or without TGF-β3 as induction factor. The results showed that with micromass culture mimicking relatively high cell density in vivo, the chondrogenic differentiation of MSCs can be promoted by soft substrates (about 0.5 kPa) independently with assembled cytoskeleton. Further analysis indicated that addition of TGF-β3 generally increased expression level of cartilage-related markers and masked the stiffness-derived expression pattern of hypertrophic markers. These results demonstrate how mechanical cues experienced in developmental context regulate commitment of stem cell fate and have significant impact on the design of tissue regeneration materials.
... Features of the analyzed scaffolds are given in Table 1. In previous investigations, expression levels in human osteoblasts increased only on the stiffer surfaces (310 MPa) [53]. Higher matrix moduli in 2D and 3D structures have generally been found to promote osteogenesis. ...
Article
Full-text available
Autologous cell therapy uses patients' own cells to deliver precise and ideal treatment through a personalized medicine approach. Isolation of patients' cells from residual tissue extracted during surgery involves specific planning and lab steps. In the present manuscript, a path from isolation to in vitro research with human mesenchymal stem cells (MSCs) obtained from residual bone tissues is described as performed by a medical unit in collaboration with a research center. Ethical issues have been addressed by formulating appropriate harvesting protocols according to European regulations. Samples were collected from 19 patients; 10 of them were viable and after processing resulted in MSCs. MSCs were further differentiated in osteoblasts to investigate the bi-ocompatibility of several 3D scaffolds produced by electrospinning and 3D printing technologies; traditional orthopedic titanium and nanostructured titanium substrates were also tested. 3D printed scaffolds proved superior compared to other substrates, enabling significantly improved response in osteoblast cells, indicating that their biomimetic structure and properties make them suitable for synthetic tissue engineering. The present research is a proof of concept that describes the process of primary stem cells isolation for in vitro research and opens avenues for the development of person-alized cell platforms in the case of patients with orthopedic trauma. The demonstration model has promising perspectives in personalized medicine practices.
Chapter
Bone implants play a pivotal role in orthopedics as they represent one of the overarching treatments for patients with structural damage as well as other orthopedic diseases. Here, we reviewed the important aspects of bone implants and their differential roles in various orthopedic applications. In particular, we focused on implants developed and utilized in bone regeneration and bone cancer treatments in order to provide an overview for broader audience of this chapter. For bone regeneration, we mainly concentrated on various strategies in biomaterials design and applications as bone implants. For bone cancer therapy, we reviewed both bone cancer therapies in clinical and research phases. Finally, we also provide our perspectives regarding the current limitations and future directions for the interest of the audiences.
Article
Increased life expectancy has led to a rise in age-related disorders including neurological diseases such as Alzheimer's disease and Parkinson's disease. Limited progress has been made in the development of clinically translatable therapies for these central nervous system (CNS) diseases. Challenges including the blood−brain barrier, brain complexity, and comorbidities in the elderly population are some of the contributing factors toward lower success rates. Various invasive and noninvasive ways are being employed to deliver small and large molecules across the brain. Biodegradable, implantable drug-delivery systems have gained lot of interest due to advantages such as sustained and targeted delivery, lower side effects, and higher patient compliance. 3D printing is a novel additive manufacturing technique where various materials and printing techniques can be used to fabricate implants with the desired complexity in terms of mechanical properties, shapes, or release profiles. This review discusses an overview of various types of 3D-printing techniques and illustrative examples of the existing literature on 3D-printed systems for CNS drug delivery. Currently, there are various technical and regulatory impediments that need to be addressed for successful translation from the bench to the clinical stage. Overall, 3D printing is a transformative technology with great potential in advancing customizable drug treatment in a high-throughput manner.
Article
The combined effect of surface topography and substrate rigidity in stem cell cultures is still under‐investigated, especially when biodegradable polymers are used. Herein, we assessed human bone marrow stem cell response on aliphatic polyester substrates as a function of anisotropic grooved topography and rigidity (7 and 12 kPa). Planar tissue culture plastic (TCP, 3 GPa) and aliphatic polyester substrates were used as controls. Cell morphology analysis revealed that grooved substrates caused nuclei orientation/alignment in the direction of the grooves. After 21 days in osteogenic and chondrogenic media, the 3 GPa TCP and the grooved 12 kPa substrate induced significantly higher calcium deposition and alkaline phosphatase (ALP) activity and glycosaminoglycan (GAG) deposition, respectively, than the other groups. After 14 days in tenogenic media, the 3 GPa TCP upregulated four and downregulated four genes; the planar 7 kPa substrate upregulated seven genes and downregulated one gene; and the grooved 12 kPa substrate upregulated seven genes and downregulated one gene. After 21 days in adipogenic media, the softest (7 kPa) substrates induced significantly higher oil droplet deposition than the other substrates and the grooved substrate induced significantly higher droplet deposition than the planar. Our data pave the way for more rational design of bioinspired constructs.
Article
The stiffness of most biomaterials used in bone tissue engineering is static at present, and does not provide an ideal biomimetic dynamical mechanical microenvironment for bone regeneration. To simulate the dynamic stiffness better during bone repair, the preparation of dynamic materials, especially hydrogels, has aroused researchers' interest. However, there are still many problems limiting the development of hydrogels such as small-scale stiffness changes and unstable mechanical properties. Here, magnetic liquid metal (MLM) was introduced into bone tissue engineering for the first time. A MLM scaffold was obtained by adding magnetic silicon dioxide particles ([email protected]2) into galinstan. Furthermore, a porous MLM (PMLM) scaffold was obtained by adding polyethylene glycol as a template to the MLM scaffold. Both scaffolds can respond to external magnetic fields, so changing the magnetic field intensity can achieve a large-scale of dynamic stiffness change. The results showed that the MLM scaffold has good biocompatibility and can promote the osteogenic differentiation of mesenchymal stem cells (MSCs). The PMLM scaffold with dynamic stiffness can promote new bone regeneration and osseointegration in vivo. Our research will open up a new field for the application of liquid metal and bring new ideas for the development of bone tissue engineering materials.
Article
Full-text available
Objective The objective of this study was to investigate the impact of two different commercially available dental implants on osseointegration. The surfaces were sandblasting and acid etching (Group 1) and sandblasting and acid etching, then maintained in an isotonic solution of 0.9% sodium chloride (Group 2). Material and Methods X-ray photoelectron spectroscopy (XPS) was employed for surface chemistry analysis. Surface morphology and topography was investigated by scanning electron microscopy (SEM) and confocal microscopy (CM), respectively. Contact angle analysis (CAA) was employed for wetting evaluation. Bone-implant-contact (BIC) and bone area fraction occupied (BAFO) analysis were performed on thin sections (30 μm) 14 and 28 days after the installation of 10 implants from each group (n=20) in rabbits' tibias. Statistical analysis was performed by ANOVA at the 95% level of significance considering implantation time and implant surface as independent variables. Results Group 2 showed 3-fold less carbon on the surface and a markedly enhanced hydrophilicity compared to Group 1 but a similar surface roughness (p>0.05). BIC and BAFO levels in Group 2 at 14 days were similar to those in Group 1 at 28 days. After 28 days of installation, BIC and BAFO measurements of Group 2 were approximately 1.5-fold greater than in Group 1 (p<0.05). Conclusion The surface chemistry and wettability implants of Group 2 accelerate osseointegration and increase the area of the bone-to-implant interface when compared to those of Group 1.
Article
Full-text available
The aim of this study was to investigate the combined effects of physical and chemical surface factors on in vivo bone responses by comparing chemically modified hydrophilic sandblasted, large-grit, acid-etched (modSLA) and anodically oxidized hydrophobic implant surfaces. Five modSLA implants and five anodized implants were inserted into the tibiae of five New Zealand white rabbits (one implant for each tibia). The characteristics of each surface were determined using field emission scanning electron microscopy, energy dispersive spectroscopy, and confocal laser scanning microscopy before the installation. The experimental animals were sacrificed after 1 week of healing and histologic slides were prepared from the implant-tibial bone blocks removed from the animals. Histomorphometric analyses were performed on the light microscopic images, and bone-to-implant contact (BIC) and bone area (BA) ratios were measured. Nonparametric comparison tests were applied to find any significant differences (P<0.05) between the modSLA and anodized surfaces. The roughness of the anodized surface was 1.22 ± 0.17 µm in Sa, which was within the optimal range of 1.0-2.0 µm for a bone response. The modSLA surface was significantly rougher at 2.53 ± 0.07 µm in Sa. However, the modSLA implant had significantly higher BIC than the anodized implant (P=0.02). Furthermore, BA ratios did not significantly differ between the two implants, although the anodized implant had a higher mean value of BA (P>0.05). Within the limitations of this study, the hydrophilicity of the modSLA surface may have a stronger effect on in vivo bone healing than optimal surface roughness and surface chemistry of the anodized surface.
Article
Full-text available
Runt-related transcription factor 2 (RUNX2) is a transcription factor closely associated with the osteoblast phenotype. While frequently referred to, the complexity of its regulation and its interactions within the osteoblast differentiation pathway are often overlooked. This review aims to summarise the knowledge of its regulation at the transcriptional, translational and post-translational level. In addition, the regulation of RUNX2 by factors commonly used during osteogenic studies will be discussed.
Article
Abstract Runx2 transcription factor is essential for the development of mineralized tissue, and is required for osteoblast commitment and chondrocyte maturation. Mice with global deletion of Runx2 exhibit complete failure of bone tissue formation, while chondrocyte-specific Runx2-deficient mice lack endochondral ossification. However, the function of Runx2 after commitment of mesenchymal cells to the osteoblast lineage remains unknown. Here, we elucidate the osteoblast-specific requirements of Runx2 during development of the tissue. Runx2 was deleted in committed osteoblasts using Cre-recombinase driven by the 2.3kbCol1a1 promoter. Surprisingly, Runx2(ΔE8/ΔE8) mice were born alive and were essentially indistinguishable from wild-type littermates. At birth, we failed to detect any alterations in skeletal patterning or extent of bone development in homozygous mutants. However, by 4 weeks of age, mutant mice showed obvious growth deficiencies, and weighed 20-25% less than sex-matched wild-type littermates. Micro-CT analysis of the hindlimb revealed a dramatic decrease of 50% in both cortical and trabecular bone volume compared with wild-type mice. Consistent with this observation, trabecular number and thickness were decreased by 51% and 21%, respectively, and trabecular space was increased by 2-fold in limbs of Runx2(ΔE8/ΔE8) mice. In addition to poor acquisition of bone mass, the average density of hydroxyapatite was markedly decreased in bone of Runx2(ΔE8/ΔE8) mice. Together, these findings demonstrate that loss of Runx2 activity in committed osteoblasts impairs osteoblast function, and that Runx2 is critical for postnatal, but not embryonic endochondral ossification.
Article
Objectives: This study evaluated the effect of implant macrodesign and position, related to the bone crest, on bone-to-implant contact (BIC) and crestal bone (CB) in immediate implants. Material and methods: The study comprised of six foxhound dogs in which 48 immediate implants were placed. Three types of implants from the same manufacturer with similar surface characteristics but different macrodesigns were randomly placed: Group A (external hex with no collar microthreads), Group B (internal hex and collar microrings), and Group C (internal conical connection and collar microrings). Half of the implants were placed leveled with the bone crest (control) and the remaining, 2 mm subcrestally (test). Block sections were obtained after 12 weeks and processed for mineralized ground sectioning. Statistical analysis consisted of nonparametric Friedman and Wilcoxon test. Results: All implants were clinically stable and histologically osseointegrated. Mean BIC percentage within the control group was as follows: A: 42.52 ± 8.67, B: 35.19 ± 18.12, and C: 47.46 ± 11.50. Within the test group: A: 47.33 ± 5.23, B: 48.38 ± 11.63, and C: 54.88 ± 11.73. Differences between each subgroup in the test and the control groups were statistically significant. BIC was statistically significantly higher in the test (50.588 ± 8.663) than in the control (43.317 ± 9.851) group. Within both groups, differences between group C and the other 2 were statistically significant. Distance from the implant shoulder to the buccal CB was statistically significantly larger in the control than in the test group and between subgroups B and C in the control and test groups. Within the test groups, relative bone gain was noticed. Conclusions: Subcrestal immediate implant positioning may lead to a relatively reduced CB resorption and increased BIC. Implants macrodesign with crestal microrings may enhance BIC in post-extraction implants.
Article
The role of mature collagen cross-links, pentosidine (Pen) cross-links in particular, in the micromechanical properties of cancellous bone is unknown. The aim of this study was to examine nonenzymatic glycation effects on tissue stiffness of demineralized and non-demineralized cancellous bone. A total of 60 bone samples were derived from mandibular condyles of six pigs, and assigned to either control or experimental groups. Experimental handling included incubation in phosphate buffered saline alone or with 0.2 M ribose at 37 °C for 15 days and, in some of the samples, subsequent complete demineralization of the sample surface using 8% EDTA. Before and after experimental handling, bone microarchitecture and tissue mineral density were examined by means of microcomputed tomography. After experimental handling, the collagen content and the number of Pen, hydroxylysylpyridinoline (HP), and lysylpyridinoline (LP) cross-links were estimated using HPLC, and tissue stiffness was assessed by means of nanoindentation. Ribose treatment caused an up to 300-fold increase in the number of Pen cross-links compared to nonribose-incubated controls, but did not affect the number of HP and LP cross-links. This increase in the number of Pen cross-links had no influence on tissue stiffness of both demineralized and nondemineralized bone samples. These findings suggest that Pen cross-links do not play a significant role in bone tissue stiffness.
Article
Hypothesizing that the implantation of non-articular (heterotopic) chondrocytes might be an alternative approach to support articular cartilage repair, we analysed joint cartilage defect healing in the rabbit model after implantation of autologous auricle-derived (auricular) chondrocytes. Autologous lapine articular and auricular chondrocytes were cultured for three weeks in polyglycolic acid (PGA) scaffolds before being implanted into critical sized osteochondral defects of the rabbit knee femoropatellar groove. Cell-free PGA scaffolds and empty defects served as controls. Construct quality was determined before implantation and defect healing was monitored after 6 and 12 weeks using vitality assays, macroscopical and histological score systems. Neo-cartilage was formed in the PGA constructs seeded with both articular and auricular chondrocytes in vitro and in vivo. At the histological level, cartilage repair was slightly improved when using autologous articular chondrocyte seeded constructs compared to empty defects and was significantly superior compared to defects treated with auricular chondrocytes 6 weeks after implantation. Although only the immunohistological differences were significant, auricular chondrocyte implantation induced an inferior healing response compared with the empty defects. Elastic auricular chondrocytes might maintain some tissue-specific characteristics when implanted into joint cartilage defects which limit its repair capacity.