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Epithelial Membrane Protein 1 Inhibits Human Spinal Chondrocyte Differentiation.

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THE ANATOMICAL RECORD 294:1015–1024 (2011)
Epithelial Membrane Protein 1 Inhibits
Human Spinal Chondrocyte
Department of Human Anatomy, Second Military Medical University, Shanghai, China
Department of Orthopedics, No. 309 Hospital of the Chinese People’s Liberation Army,
Beijing, China
The molecular mechanisms underlying human spinal chondrocyte differentiation remain unclear. We recently demonstrated that epithelial
membrane protein 1 (EMP1) is highly expressed in degenerative intervertebral discs. EMP1 is involved in the differentiation of multiple cell types,
including progenitor/pre-B cells, neurons, and podocytes. Therefore, we
hypothesize that EMP1 may participate in the differentiation of spinal
chondrocytes. We cultured chondrocytes from human nucleus pulposus.
Through lentivirus-mediated knockdown and overexpression of EMP1, we
find that EMP1 promotes cell proliferation and survival, alters cell morphology and cell cycle, reduces cell condensation, and inhibits cell hypertrophy and the expression of chondrocyte maturation markers such as
collagen X, aggrecan, sex-determining region Y (SRY)-box 9, and runtrelated transcription factor 2. We also show that EMP1 is not expressed
in the ossification center of vertebrae but is highly expressed in the nucleus pulposus and growth plate, where chondrocytes are immature and
endochondral ossification has not occurred. These results suggest that
EMP1 inhibits human spinal chondrocyte differentiation. Anat Rec,
C 2011 Wiley-Liss, Inc.
294:1015–1024, 2011. V
Key words: epithelial membrane protein 1; chondrocytes;
differentiation; intervertebral disc development
Lower back pain caused by disc degeneration is one of
the most common clinical conditions worldwide; yet,
there is no completely effective treatment because the
molecular mechanisms of chondrocyte differentiation,
and disc growth and maintenance after birth, are not
well understood. Chondrocyte differentiation is a multistep process that begins with mesenchymal cell condensation and commitment (Widelitz et al., 1993;
Oberlender and Tuan, 1994; DeLise and Tuan, 2002), followed by morphological changes (Archer et al., 1982;
Glowacki et al., 1983; Loty et al., 2000), cell proliferation, and secretion of the chondrogenic matrix (Muir,
1995). Joint cartilage and nucleus pulposus are formed
by chondrocytes that differentiate at this stage. During
terminal differentiation, chondrocytes become hypertrophic and exit the cell cycle (Gerstenfeld and Shapiro,
1996; Hirsch et al., 1996; Shum and Nuckolls, 2002).
Finally, chondrocytes undergo apoptosis, mineralization
occurs, and cartilage is replaced with bone (Urban et al.,
2000; Karsenty and Wagner, 2002; Kronenberg, 2003).
In our recent screen for genes that are differentially
expressed between degenerative and normal discs, we
discovered that epithelial membrane protein 1 (EMP1)
was highly expressed in human degenerative nucleus
pulposus (Hu et al., 2004). EMP1 (Lobsiger et al., 1996),
also known as CL-40 (Taylor et al., 1995), tumor-associated membrane protein (TMP) (Ben-Porath et al., 1999),
B4B (Ruegg et al., 1996), and pancreatitis-associated
protein (PAP) (Schiemann et al., 1997), is a putative tetraspan transmembrane protein that participates in
numerous cellular processes, including proliferation
*Correspondence to: Chuan-Sen Zhang, Department of Anatomy, Second Military Medical University, Shanghai 200433,
China. Fax: þ86-21-81870949. E-mail:
Received 5 April 2010; Accepted 9 March 2011
DOI 10.1002/ar.21395
Published online 29 April 2011 in Wiley Online Library
(Ben-Porath et al., 1999), apoptosis, membrane trafficking, and cell adhesion (Wilson et al., 2002).
EMP1 is expressed by a number of immature cell
types, including embryonic stem cells, adult liver stem
cells, immune pre-B cells, and neuronal progenitors (Lee
et al., 2005). In fact, EMP1 is highly expressed in these
cell types transitionally but is not expressed after terminal differentiation. Therefore, EMP1 could participate in
the dedifferentiation processes associated with intervertebral disc degeneration.
To clarify the role of EMP1 in the differentiation of
intervertebral disc chondrocytes, we cultured spinal
chondrocytes derived from fetal nucleus pulposus, which
can undergo terminal differentiation in culture media
supplemented with fetal bovine serum (FBS). Furthermore, following lentivirus-mediated knockdown and
overexpression of EMP1, we observed changes in proliferation, apoptosis, cell cycle length, cellular morphology,
cell condensation, and expression of chondrocyte maturation markers such as sex-determining factor (SRY)-box 9
(Sox9), collagen I, collagen II, collagen X, aggrecan,
and runt-related transcription factor 2 (Runx2). We also
analyzed EMP1 expression in the fetal spine via
Tissue Collection and Culture of Human Fetal
Nucleus Pulposus Cells
This study was examined and approved by the bioethics committee of the Second Military Medical University (Shanghai, China). Tissue was collected from
naturally aborted fetuses donated for research purposes
to the Department of Human Anatomy of the Second
Military Medical University. After obtaining informed
consent from the donors, three fetuses aged 14–18
weeks, not clinically found to have any hereditary or infectious diseases, were used for this study.
The nucleus pulposus derived from donor fetuses was
sheared into 1-mm3 chips and digested with 0.2% collagenase Type II (Roche Molecular Biochemicals, Germany) in Dulbecco’s Modified Eagle’s Medium/Ham’s
F-12 (DMEM/F12, GIBCO BRL, Paisley, Scotland), with
agitation for 6 hr. After two washes in DMEM/F12, the
cells were cultured in DMEM/F12 with 10% FBS
(Hyclone Laboratories, UT) at a density of 2 105 per
T25 flask (Corning Glass Works, Corning, NY). Cells
were used between passages 3 and 11.
Construction of Recombinant Lentiviruses
The lentivirus encoding small hairpin RNA (shRNA)
directed against human EMP1 and enhanced green fluorescent protein (EGFP) was constructed by Genechem
(Shanghai, China). The targeting sequence of the shRNA
was GGACTTAGAAGTAGTATGT, which was confirmed
by sequencing. Both the recombinant lentivirus encoding
shRNA targeting EMP1 (EMP1-shRNA-Lentivirus) and
the lentivirus encoding scrambled shRNA (scrambleshRNA-lentivirus) were prepared and titered to 108
transfection units (TU)/mL.
The recombinant lentivirus expressing EMP1 was constructed by Innovation Biotechnology (Shanghai, China).
The coding sequence of EMP1 was obtained from the
previously constructed plasmid pcDNA3.1 (þ)-EMP1 via
PCR. The forward and reverse primers used were as
follows: 50 -CGTTGACATTGATTATTGACTAG-30 and 50 GAGTTATTTCTTTCTCAGGACC-30 , respectively. The
PCR product was purified by gel extraction, double
digested, and inserted into linearized lentivirus shuttle
plasmid. The shuttle plasmid sequence was confirmed by
sequencing, and the lentivirus containing EMP1 was
constructed and titered to 108 TU/mL.
Cell Transfection
Cell transfection was achieved by culturing cells in a
culture flask at a concentration of 3 105 per T25 flask
for 48 hr. When cells reached the logarithmic proliferation stage, at about 60% confluence, the culture medium
was changed and lentivirus stock solution was added to
these cells at a multiplicity of infection of 10 with 8 lg/
mL polybrene. The medium was changed 10 hr later.
Semiquantitative and Quantitative RT-PCR
To detect EMP1 transcript expression following lentiviral infection, total RNA was extracted from cultured cells
3 days after transfection using TRIzol reagent (Invitrogen
Life technology, Carlsbad, CA) according to the manufacturer’s instructions. A total of 1.5 mg of RNA was used for
transcription, and 1/20 of the RT product was used for
PCR amplification. The primers used to amplify EMP1
were as follows: forward 50 -ATCTTTGTGGTCCACATC
GCT-30 and reverse 50 -CTTCTCCATGGTGAAGAGCT-30 .
Glyceraldehyde 3-phosphate dehydrogenase (GAPDH)
was used as an internal control, and the following primers were used: forward 50 -TTCAGCTCAGGGATGACC
TT-30 and reverse 50 -GGCATGGACTGTGGTCATGAG-30 .
PCR conditions used were as follows: 95 C for 4 min; 95 C
for 45 sec, 54 C for 45 sec, 72 C for 30 sec, 27 cycles; 72 C
for 10 min, 4 C for 5 min. PCR products were tested via
1.2% agarose gel electrophoresis. Three replicates of each
independent experiment were performed.
To detect the expression of chondrocyte maturation
markers, total RNA from cultured cells was extracted 6
weeks after transfection and converted to cDNA using
reverse transcriptase (RT). Quantitative real-time PCR
was performed using an ABI 7500 Real-Time PCR System (Applied Biosystems, Foster City, CA). RT-PCR was
carried out using the SYBR Green PCR master mix
(Applied Biosystems, Foster City, CA) with 1 mL cDNA
template in a 20-mL final reaction mixture (95 C for 15
min; 95 C for 15 sec, 60 C for 60 sec, 40 cycles). The average threshold cycle (Ct) for each gene was determined
from triplicate reactions, and target gene expression was
normalized to GAPDH. This was then calibrated to the
control sample in each experiment to give the DDCt
value, where the control had a DDCt value of 0. The fold
target gene expression, when compared with the control
value, was given by the formula 2DDCt. Primer sequences are shown in Table 1.
Western Blotting
One week after lentivirus-mediated transfection, total
protein was extracted using radioimmunoprecipitation
assay lysis buffer [50 mM Tris-HCl (pH 7.4), 150 mM
NaCl, 1% Triton X-100, 1% Nonidet P-40, 1 mM EDTA,
1 mM phenylmethylsulfonyl fluoride, and protease
TABLE 1. Primer sequences
Collagen I
Collagen II
Collagen X
Analysis of Apoptosis and Cell Cycle
inhibitor mixtures (1, Roche Molecular Biochemicals)].
Equal amounts of protein were loaded and separated via
12% sodium dodecyl sulfate polyacrylamide gel electrophoresis. Separated proteins were transferred to 0.2-mm
polyvinylidene difluoride membranes, and the membranes were blocked in 5% skim milk for 3 hr before
incubating with mouse anti-human EMP1 antibody
(1:1,000, Abnova, China) or mouse anti-human GAPDH
antibody (1:5,000, Biosynthesis Biotechnology, China)
diluted in blocking solution overnight. After washing
three times with Tris-buffered saline (TBS) with Tween
20 (TBST) (50 mM Tris-HCl (pH 7.5), 150 mM NaCl,
and 0.05% Tween 20), horseradish peroxidase-conjugated secondary antibodies were incubated with the
membranes for 1 hr at room temperature (RT). Membranes were washed twice with TBST and once in TBS
and then soaked in enhanced chemiluminescence (ECL)
reagent. Protein bands were visualized using Western
Blotting Systems ECL kit (Amersham, USA). Data
obtained from the Western blot experiments were analyzed using Bio-Rad Quantity One 1-D Analysis software (Bio-Rad, USA).
Cell Viability and Proliferation Assays
The proliferation and viability of transfected, cultured
human disc cells were measured by generating a growth
curve and by using the Cell Counting Kit-8 (CCK-8)
assay (Dojindo, Kumamoto, Japan). CCK-8 contains
WST-8, water-soluble tetrazolium salt-8, which is
reduced by dehydrogenase activity of a viable cell to produce a yellow color formazan dye. Cells were inoculated
into a 96-well plate at an initial density of 3 103 per
well (each group consisting of five wells). After incubation for 24 hr, 10 lL CCK-8 was added to each well containing 200 lL culture medium and incubated for 3 hr
at 37 C. Viable cells were counted by absorbance measurements at 450 nm using an automicroplate reader
(Infinite M200, Tecan, Austria).
To generate a growth curve, cells were inoculated into
a 24-well plate at a density of 104 per well. Cells in each
well were digested and counted at 24-hr intervals, using
a hemocytometer, for 5 consecutive days. Each independent experiment was repeated two times.
Cells were treated with the protein synthesis inhibitor
cycloheximide (CHX), the protein kinase C inhibitor staurosporine (ST), or in suspension culture for a period of time
before analysis. The suspension culture consisted of 1.5%
(w/v) agarose solution in phosphate-buffered saline (PBS)
that had been autoclaved and cooled to 70 C and was
poured into 25-cm2 polystyrene tissue culture flasks. When
the solution turned to a gel, 5 mL of DMEM/F12 medium
was added to the flask to replace the PBS in the gel. The
antiattachment flask was ready for use 2 hr later. The fluid
on top of the gel was removed before seeding cells.
Treated cells, including those floating in the medium,
were collected by digestion, washed with PBS twice,
fixed with 75% ethanol, stained with propidium iodide,
and detected by flow cytometry.
Cells were cultured in suspension for 16 hr, centrifuged to remove the medium, and cultured for 30 min
after addition of a binding buffer composed of annexin
v-PE and 7-aminoactinomycin D (7-AAD, BD Biosciences
Pharmingen, USA). Annexin V could detect phosphatidylserine exposure on the cell surface as a marker of apoptosis, and 7-AAD could stain dead cells. Early
apoptotic cells are Annexin V-FITC positive but 7-AAD
negative. Whole cells were detected by flow cytometry.
Cells were cultured in suspension for 16 hr before treatment with 1 mL Red-DEVD-FMK (Biovision, USA), which
can bind activated caspase 3. Cells were then cultured for
additional 30 min, washed with wash buffer (Biovision,
USA) three times, and detected by flow cytometry.
Detection of Cell Condensation and
Morphology After EMP1 Overexpression
Cell condensation was observed in flask or suspension
cultured cells. Suspension culture conditions were identical to those mentioned in the ‘‘Analysis of apoptosis
and cell cycle’’ subsection. Cell morphology was observed
every other day after passaging.
Detection of EMP1 Expression in Human Spine
To detect EMP1 expression via immunohistochemistry,
isolated vertebral tissue was fixed in 4% paraformaldehyde overnight, decalcified in 10% formic acid for 24 hr,
dehydrated, hyalinized, paraffin embedded, and sectioned
at 6-lm thickness. The tissue slices were hydrated, and
antigen retrieval was performed by treating with proteinase K [20 mg/mL in buffer containing 20 mM Tris-HCl
(pH 8.0)] for 10 min. The slides were then incubated in
Triton X-100 (0.3%) for 10 min. Endogenous peroxidases
were inactivated by treating the tissue with 1% H2O2 for
20 min. The sections were then blocked with 3% bovine
serum albumin in PBS (PBSB) for 1 hr and incubated
with primary antibody (1:100 in PBSB) at 4 C overnight,
sealed with antibody dilution (1:100), incubated with avidin-labeled secondary antibody at RT for 1 hr, reacted
with streptavidin–biotin complex for 30 min, and the
color reaction was developed using 3,30 diaminobenzidine.
For detection of EMP1 in spinal tissue via semiquantitative PCR, the nucleus pulposus and decorticated
vertebral tissue were separated, cut into chips, and homogenized completely in TRIzol reagent. The subsequent
RT-PCR procedures were the same as those used for cultured cells.
Statistical Analysis
Differences between groups were analyzed using the
Student t test. P values of <0.05 were considered statistically significant.
EMP1 Promotes the Proliferation and Viability
of Cultured Human Fetal Nucleus Pulposus
We derived human fetal nucleus pulposus cells
(hfNPCs) from 14 to 18 weeks human fetal nucleus pulposus, which is a jelly-like substance in the middle of
the spinal disc. Forty-eight hours after transfection,
transfection efficiency of lentivirus was examined
through assessing the percentage of cells positive for
EGFP under a fluorescent microscope, and the transfection efficiency was more than 80%. To test the effects of
EMP1 on chondrocyte proliferation and viability, we generated a growth curve and performed a CCK-8 assay 4 weeks
after cultured chondrocytes were transfected with EMP1
(Fig. 1A). The growth curve obtained suggested that
EMP1-transfected cells proliferated more rapidly than control cells (transfected with a virus encoding EGFP alone).
Furthermore, cells transfected with EMP1-shRNA-lentivirus proliferated more slowly than those transfected with
scrambled-shRNA-lentivirus (Fig. 1B). Although all cultures were started at the same cell density, cells transfected with EMP1 were completely confluent after 3 days,
whereas cells in the control group achieved 75% confluence
during the same period (Fig. 1D). The CCK-8 assay results
showed enhanced cell viability in cells transfected with
EMP1, whereas cells expressing EMP1-shRNA exhibited
reduced viability (Fig. 1C).
EMP1 Protects hfNPCs From Apoptosis
Immature chondrocytes are resistant to apoptosis but
are known to undergo programed cell death during the
terminal differentiation process. Therefore, we examined
the effects of EMP1 expression on apoptosis by analyzing
the numbers of sub-G1 and annexin V(þ), 7-AAD()
cells. The sub-G1 peak was detected via flow cytometry
in cells treated with ST or CHX and transfected with
EMP1 or shRNA directed against EMP1. Knockdown of
EMP1 expression increased the ratio of the sub-G1 apoptosis peak (from 1.25% 0.14% to 1.51% 0.13% with
20 lM CHX for 8 hr; 3.26% 0.31% to 4.11% 0.51%
with 0.01 lM ST for 8 hr; and 1.58% 0.16% to 5.83%
0.64% in suspension conditions for 12 hr) (Fig. 2A).
Likewise, overexpression of EMP1 reduced the ratio of
the apoptosis peak, when compared with that of controltreated cells under similar conditions (from 0.74% 0.08% to 0% 0% with 20 lM CHX for 8 hr; 4.12% 0.45% to 0.13% 0.02% with 0.01 lM ST for 8 hr; and
32.87% 4.2% to 11.35% 1.3% in suspension culture
for 24 hr) (Fig. 2B). Our results also suggest that EMP1
expression protected cells from apoptosis in the presence
of different concentrations of ST or CHX (from 1.31% 0.12% to 0.23% 0.02% with 10 lM CHX for 16 hr;
7.68% 0.83% to 0.85% 0.09% with 40 lM CHX for
16 hr; 20.03% 2.8% to 1.34% 0.14% with 100 lM
CHX for 6 hr; 7.94% 0.81% to 2.78% 0.29% with 0.1
lM ST for 16 hr; 1.85% 0.19% to 0.89% 0.07% with
0.2 lM ST for 6 hr; and 2.54% 0.22% to 2.05% 0.16% with 1 lM ST for 6 hr) (Fig. 2E).
However, the sub-G1 peak is considered a nonspecific
marker of apoptosis. Therefore, we used both annexin-VPE and 7-AAD to detect apoptotic changes after EMP1
overexpression. The results showed that the percentage
of apoptotic cells decreased from 9.79% in controls to
2.06% in EMP1-overexpressing cells when EGFP-positive cells were analyzed (Fig. 2D). We also found that
cells transfected with EMP1-shRNA-lentivirus exhibited
increased caspase-3 activity (Fig. 2C).
EMP1 Expression Accelerates the Cell Cycle
in Cultured hfNPCs
EMP1 overexpression increased cell viability and
enhanced cell proliferation, implying that, given a fixed
cell cycle length, more S/G2/M-phase cells should be
present in this condition. However, the results of our cell
cycle detection experiments showed that EMP1 knockdown resulted in an increase in S/G2/M-phase cells (from
65.10% 5.87% to 76.50% 4.27%). In particular, the
proportion of S-phase cells was increased from 43.12% 6.12% to 54.17% 2.98% (Fig. 3A). After EMP1 overexpression, the number of proliferating cells decreased from
34.83% 6.64% to 18.36% 8.49%, and the percentage
of S-phase cells was decreased from 22.62% 4.89% to
9.72% 3.96% (Fig. 3B). These results indicate that the
cell cycle length might be different depending on the level
of EMP1 expression. For example, S-phase may be
shorter in cells overexpressing EMP1.
EMP1 Inhibits the Expression of Chondrocyte
Maturation Markers
To test the influence of EMP1 on the expression of the
chondrocyte maturation markers, quantitative RT-PCR
was performed on EMP1 knockdown and overexpressing
cells. Collagen I was considered a marker of undifferentiated chondrocytes, whereas collagen II, collagen X,
Sox9, aggrecan, and Runx2 were considered to be cell
maturation markers. Knockdown of EMP1 resulted in a
fourfold decrease in collagen I expression and a 1.3-fold
decrease in collagen II expression, as well as increases
in collagen X (3-fold), Sox9 (1.5-fold), aggrecan (7.8fold), and Runx2 (5.9-fold) expression (Fig. 4A). Overexpression of EMP1 resulted in an 8.2-fold increase in
collagen I expression and decreases in collagen X, Sox9,
aggrecan, and Runx2 expression (by about 67%, 50%,
75%, and 85%, respectively) (Fig. 4B).
Overexpression of EMP1 Inhibits
Condensation of Cultured hfNPCs
Because the chondrocyte differentiation process begins
with cell condensation, cell adhesion changes following
EMP1 overexpression were evaluated. In control cultures, we found overlapping or multilayered cells, which
formed cell clusters, especially at or near confluence.
However, no overlapping cells or cell clusters were
observed after EMP1 overexpression. Under suspension
culture conditions, cells overexpressing EMP1 rarely
condensed and seldomly formed large cell clusters when
compared with control cells (Fig. 5A).
Fig. 1. Analysis of EMP1 expression after transfection and the role
of EMP1 in chondrocyte proliferation. A: Western blotting and semiquantitative RT-PCR confirmed lentivirus-mediated EMP1 overexpression and RNAi silencing. Cell extracts were prepared 3 days after
transfection, and GAPDH was used as an internal control. B: EMP1
enhanced chondrocyte proliferation. Growth curves were generated 4
weeks after RNAi-mediated silencing or overexpression of EMP1. Independent experiments were performed in duplicate. C: EMP1
enhanced chondrocyte viability. Cell viability was quantified by CCK-8
assay after 24 hr of culture. Values are expressed as means SE (N
¼ 5) *P < 0.05; **P < 0.01. D: Photos were taken daily after seeding 3
105 EGFPþ or EMP1/EGFPþ cells. Scale bar ¼ 20 mm.
Fig. 2. The role of EMP1 in chondrocyte survival. A: Apoptosis levels increased after EMP1 knockdown. B: Apoptosis levels decreased
after EMP1 overexpression. Sub-G1 peak was assayed by flow
cytometry. C: Cells in the EMP1 shRNA and scrambled shRNA groups
were analyzed by flow cytometry after suspension culture for 16 hr
and incubation with RED-DEVD-FMK caspase 3 reagent for 30 min.
D: Apoptosis was detected in 16-hr suspension cultures via flow
cytometry in EGFPþ- and EGFP/EMP1þ-transfected cells after preincubation with annexin V-PE and 7-AAD for 30 min. E: Apoptosis (SubG1 peak) at different concentrations of cycloheximide (CHX) and staurosporine (ST) in EGFPþ and EGFP/EMP1þ groups.
Overexpression of EMP1 Induces
Morphological Changes and Inhibits
Hypertrophy of Cultured hfNPCs
trophy. Therefore, we examined the morphology of cultured chondrocytes after EMP1 overexpression. Two
weeks after transfection with the EMP1-encoding virus,
cells grew out multiple, tiny cytoplasmic processes and
appeared to flatten. Six weeks after transfection, cells
had an oval shape or spindly, fibroblast-like shape,
Morphological changes that occur during chondrocyte
terminal differentiation typically include cellular hyper-
Fig. 3. The role of EMP1 in cell cycle progression. A: The number of cells in G0/G1 decreased, and
the number in S-phase (only light gray shading indicated) increased, after EMP1 knockdown (N ¼ 8). B:
The number of cells in G0/G1 increased, and the number of cells in S-phase decreased, after overexpression of EMP1 (N ¼ 12). *P < 0.05; **P < 0.01 vs. control.
cells transfected with EMP1 did not display these differentiation-related changes (Fig. 5C).
EMP1 Expression Is Restricted to Nucleus
Pulposus and the Vertebral Growth Plate
To examine EMP1 expression in different parts of the
spinal column, we performed immunohistochemistry and
found that EMP1 was highly expressed in nucleus pulposus cells (NPCs). No significant EMP1 expression was
detected in the annulus fibrosus. We also found that
EMP1 expression was highest in the vertebral growth
plate (Fig. 6A). Furthermore, EMP1 expression was lowest in the middle of the vertebral ossification center, and
its expression in prechondrocytes and chondrocytes was
higher than that in prehypertrophic and hypertrophic
chondrocytes (Fig. 6B).
As the density of hypertrophic cells was relatively low,
and large cell size may affect antigen density, we examined EMP1 transcript expression in the nucleus pulposus
and decorticated vertebra via semiquantitative RT-PCR.
EMP1 expression in vertebral cells was decreased by twofold that observed in the nucleus pulposus (Fig. 6C).
Fig. 4. Effects of EMP1 on the expression of chondrocyte maturation markers. A: EMP1 knockdown decreased collagen I and II expression and increased collagen X, Sox9, aggrecan, and Runx2 expression
(N ¼ 3). B: Overexpression of EMP1 increased collagen I expression
and decreased collagen X, aggrecan, Sox9, and Runx2 expression (N ¼
3). *P < 0.05; **P < 0.01 vs. control. COL, collagen; AGG, aggrecan.
rather than the typical rounded appearance (Fig. 5B).
With increased passage number, numerous hypertrophic
and dead cells were seen in the control group, whereas
In this study, we investigated the role of EMP1 in fetal intervertebral nucleus cell differentiation. Through
lentivirus-mediated overexpression and knockdown of
EMP1, we showed that EMP1 expression could promote
cell survival and proliferation, accelerate the cell cycle,
decrease the expression of chondrocyte maturation
markers, alter cell morphology, reduce cell condensation,
and inhibit hypertrophy of cultured fetal NPCs. Immunohistochemistry experiments revealed that EMP1 was
expressed in the nucleus pulposus and vertebral growth
Fig. 5. Effects of EMP1 overexpression on chondrocyte condensation and morphology. A: Reduction of cell cluster formation after
EMP1 overexpression in the presence or absence of adhesive medium. Images were taken 3 weeks after transfection. B: Representative images depict an abundance of tiny processes on cultured
chondrocytes 2 weeks after lentivirus-mediated overexpression of
EMP1; representative images of morphological changes observed 6
weeks after lentivirus-mediated overexpression of EMP1. C: Few hypertrophic cells appeared after overexpression of EMP1. Photos were
taken 8 weeks after transfection. Scale bar ¼ 20 mm.
plates. These results suggest that EMP1 can promote
chondrocyte proliferation and survival, and inhibit terminal differentiation of human spinal chondrocytes.
The human spine develops from a combination of mesenchymal and notochord cells, which form the vertebral
bodies and intervertebral discs, respectively. During the
process of chondrification, both tissues form a gelatinous
cartilage (Walmsley, 1953; Urban et al., 2000). To form
cartilage, mesenchymal cells express N-cadherin and NCAM, resulting in increased cell–cell adhesion, changes
Fig. 6. Local expression of EMP1 in developing human spine. A:
EMP1 was expressed in the growth plate (A1) and nucleus pulposus
(A3), but was absent from the annulus fibrosis (A2) and the ossification
center of the vertebral body (A4). B: Cells nearer to the growth plate
express higher levels of EMP1. Photos (N ¼ 5) were taken and divided
into four sections from growth plate to the ossification center of the
vertebrae. The number of positive cells was calculated. C: Semiquantitative RT-PCR was used to analyze the expression of EMP1 between
the nucleus pulposus and the ossification center of the vertebral body
(N ¼ 3). Values are expressed as mean SE. Scale bars ¼ 20 mm. *P
< 0.05; **P < 0.01. GP, growth plate; NP, nucleus pulposus; AF, annulus fibrosis; VB, vertebral body; OC, ossification center.
in cell morphology, and altered ECM component expression, such as increased collagen II and aggrecan expression and reduced collagen I expression. These processes
are regulated by transcription factors such as Sox9 and
Runx2 (Castagnola et al., 1988; Fukada et al., 1999).
Further differentiation occurs in the vertebrae, though
not in the intervertebral discs. Chondrocytes undergo
hypertrophy, express collagen X, withdraw from the cell
cycle, and undergo apoptosis (Gibson et al., 1997), resulting in a replacement of cartilage with bone (Kronenberg,
2003; Mackie et al., 2008). However, many of the molecular mechanisms of this process remain unclear.
EMP1 expression is restricted to the nucleus pulposus
and growth plate, where chondrocytes reside at the early
stage of differentiation. In other words, the nearer to the
growth plate, the more immature the chondrocyte, and
the highest expression of EMP1 can be observed in the
immature cells (Walmsley, 1953; Kronenberg, 2003).
Similarly, p53 apoptosis effector related to PMP22
(PERP), a member of the same gene family as EMP1, is
also expressed specifically in the intervertebral disc of
fetal mouse (Ihrie et al., 2005).
Various studies have reported that cultured chondrocytes differentiate to varying degrees in culture (Descalzi Cancedda et al., 1992; Garimella et al., 2004).
Recently, NPCs have been shown to migrate from the
end plate, suggesting that NPCs have the potential to
differentiate like chondrocytes in the end plate (Coverley
et al., 2002; Kim et al., 2003). In this study, our cultured
fetal NPCs became hypertrophic, exited the cell cycle,
and underwent apoptosis, suggesting that these cells
underwent terminal differentiation. Our morphological
analysis revealed that overexpression of EMP1 resulted
in a flattening and elongation of cultured chondrocytes.
Furthermore, cell proliferation was enhanced and cell
condensation reduced when EMP1-expressing cells were
cultured in either flasks or suspension cultures. In addition, EMP1 increased the expression of the immature
chondrocyte marker collagen I and decreased the expression of chondrocyte maturation markers such as collagen
X, aggrecan, Sox9, and Runx2, although EMP1 showed
little regulated function on the expression of collagen II.
All of these phenotypes occur during chondrocyte dedifferentiation. It is believed that cyclin A can accelerate the Sphase progression (Coverley et al., 2002). Therefore, we
propose that EMP1 may play a role in regulating cyclin A.
Interestingly, cyclin A can also form complexes with
cyclin-dependent kinase 2 (CDK2/E2F4/P130) and inhibit
the differentiation of osteoblasts (Sunters et al., 2004).
EMP1 is a member of the peripheral myelin protein
(PMP22) family, which also includes epithelial membrane proteins 2–3, PERP, and the lens membrane protein 20 (MP20). All family members are putative
tetraspan transmembrane proteins, which localize to the
plasma membrane and are involved in the secretory
pathway. Besides EMP1, other members of this gene
family seem to play an important role in regulating the
differentiation of chondrocytes. For example, PERP is
expressed specifically in the intervertebral disc of the
mouse, and loss of PERP function increases apoptosis
rates in the zebrafish notochord (Kronenberg, 2003;
Nowak et al., 2005). Furthermore, PMP22 expression
increases during redifferentiation of human articular
chondrocytes (Tallheden et al., 2004) and is frequently
highly expressed in high-grade osteosarcoma (van Dartel
and Hulsebos, 2004a,b). EMP2 is highly expressed during chondrocyte differentiation (James et al., 2005).
Therefore, the PMP22/EMP family could be important
players in chondrocyte differentiation.
We acknowledge that there are some limitations to
this study. For instance, we did not detect a change in
EMP1 expression during the terminal differentiation of
mesenchymal cells into chondrocytes. Furthermore, this
study would be enhanced by testing cultured cells for
the expression of additional chondrocyte differentiation
markers, such as Sox5, Sox6, matrix metalloproteinase
13, alkaline phosphatase, and osteocalcin. In addition,
our work would be greatly enhanced by the addition of
EMP1 knockout mouse data. However, although further
work will be needed to confirm the molecular mechanisms involved and to determine the in vivo role of
EMP1, our data suggest that EMP1 plays a role in chondrocyte proliferation and differentiation.
This work was supported by fund from the National
Natural Science Foundation of China for Distinguished
Young Scholars (30400453).
Archer CW, Rooney P, Wolpert L. 1982. Cell shape and cartilage differentiation of early chick limb bud cells in culture. Cell Differ
Ben-Porath I, Yanuka O, Benvenisty N. 1999. The tmp gene, encoding a membrane protein, is a c-Myc target with a tumorigenic activity. Mol Cell Biol 19:3529–3539.
Castagnola P, Dozin B, Moro G, Cancedda R. 1988. Changes in the
expression of collagen genes show two stages in chondrocyte differentiation in vitro. J Cell Biol 106:461–467.
Coverley D, Laman H, Laskey RA. 2002. Distinct roles for cyclins E
and A during DNA replication complex assembly and activation.
Nat Cell Biol 4:523–528.
DeLise AM, Tuan RS. 2002. Alterations in the spatiotemporal
expression pattern and function of N-cadherin inhibit cellular
condensation and chondrogenesis of limb mesenchymal cells in
vitro. J Cell Biochem 87:342–359.
Descalzi Cancedda F, Gentili C, Manduca P, Cancedda R. 1992.
Hypertrophic chondrocytes undergo further differentiation in culture. J Cell Biol 117:427–435.
Fukada K, Shibata S, Suzuki S, Ohya K, Kuroda T. 1999. In situ
hybridisation study of type I, II, X collagens and aggrecan mRNas
in the developing condylar cartilage of fetal mouse mandible.
J Anat 195 (Part 3):321–329.
Garimella R, Bi X, Camacho N, Sipe JB, Anderson HC. 2004. Primary culture of rat growth plate chondrocytes: an in vitro model
of growth plate histotype, matrix vesicle biogenesis and mineralization. Bone 34:961–970.
Gerstenfeld LC, Shapiro FD. 1996. Expression of bone-specific genes
by hypertrophic chondrocytes: implication of the complex functions of the hypertrophic chondrocyte during endochondral bone
development. J Cell Biochem 62:1–9.
Gibson G, Lin DL, Roque M. 1997. Apoptosis of terminally differentiated chondrocytes in culture. Exp Cell Res 233:372–382.
Glowacki J, Trepman E, Folkman J. 1983. Cell shape and phenotypic
expression in chondrocytes. Proc Soc Exp Biol Med 172:93–98.
Hirsch MS, Cook SC, Killiany R, Hartford Svoboda KK. 1996.
Increased cell diameter precedes chondrocyte terminal differentiation, whereas cell-matrix attachment complex proteins appear
constant. Anat Rec 244:284–296.
Hu M, Zhang CS, Chen DY. 2004. Analysis of Gene expression pattern of lumbar intervertebral disc degeneration in human. Chin J
Anat 27:348–351.
Ihrie RA, Marques MR, Nguyen BT, Horner JS, Papazoglu C, Bronson RT, Mills AA, Attardi LD. 2005. Perp is a p63-regulated gene
essential for epithelial integrity. Cell 120:843–856.
James CG, Appleton CT, Ulici V, Underhill TM, Beier F. 2005.
Microarray analyses of gene expression during chondrocyte differentiation identifies novel regulators of hypertrophy. Mol Biol Cell
Karsenty G, Wagner EF. 2002. Reaching a genetic and molecular
understanding of skeletal development. Dev Cell 2:389–406.
Kim KW, Lim TH, Kim JG, Jeong ST, Masuda K, An HS. 2003. The
origin of chondrocytes in the nucleus pulposus and histologic findings associated with the transition of a notochordal nucleus pulposus to a fibrocartilaginous nucleus pulposus in intact rabbit
intervertebral discs. Spine (Phila Pa 1976) 28:982–990.
Kronenberg HM. 2003. Developmental regulation of the growth
plate. Nature 423:332–336.
Lee HS, Sherley JL, Chen JJ, Chiu CC, Chiou LL, Liang JD, Yang
PC, Huang GT, Sheu JC. 2005. EMP-1 is a junctional protein in a
liver stem cell line and in the liver. Biochem Biophys Res Commun 334:996–1003.
Lobsiger CS, Magyar JP, Taylor V, Wulf P, Welcher AA, Program
AE, Suter U. 1996. Identification and characterization of a cDNA
and the structural gene encoding the mouse epithelial membrane
protein-1. Genomics 36:379–387.
Loty S, Sautier JM, Forest N. 2000. Phenotypic modulation of nasal septal
chondrocytes by cytoskeleton modification. Biorheology 37:117–125.
Mackie EJ, Ahmed YA, Tatarczuch L, Chen KS, Mirams M. 2008.
Endochondral ossification: how cartilage is converted into bone in
the developing skeleton. Int J Biochem Cell Biol 40:46–62.
Muir H. 1995. The chondrocyte, architect of cartilage. Biomechanics, structure, function and molecular biology of cartilage matrix macromolecules. Bioessays 17:1039–1048.
Nowak M, Koster C, Hammerschmidt M. 2005. Perp is required for
tissue-specific cell survival during zebrafish development. Cell
Death Differ 12:52–64.
Oberlender SA, Tuan RS. 1994. Expression and functional involvement of N-cadherin in embryonic limb chondrogenesis. Development 120:177–187.
Ruegg CL, Wu HY, Fagnoni FF, Engleman EG, Laus R. 1996. B4B, a
novel growth-arrest gene, is expressed by a subset of progenitor/
pre-B lymphocytes negative for cytoplasmic mu-chain. J Immunol
Schiemann S, Valentine M, Weidle UH. 1997. Assignment of the
human progression associated protein (PAP) to chromosome
12p12.3. Anticancer Res 17:4281–4285.
Shum L, Nuckolls G. 2002. The life cycle of chondrocytes in the
developing skeleton. Arthritis Res 4:94–106.
Sunters A, Thomas DP, Yeudall WA, Grigoriadis AE. 2004. Accelerated cell cycle progression in osteoblasts overexpressing the c-fos
proto-oncogene: induction of cyclin A and enhanced CDK2 activity. J Biol Chem 279:9882–9891.
Tallheden T, Karlsson C, Brunner A, Van Der Lee J, Hagg R, Tommasini R, Lindahl A. 2004. Gene expression during redifferentiation of
human articular chondrocytes. Osteoarthritis Cartilage 12:525–535.
Taylor V, Welcher AA, Program AE, Suter U. 1995. Epithelial membrane protein-1, peripheral myelin protein 22, and lens membrane protein 20 define a novel gene family. J Biol Chem
Urban JPG, Roberts S, Ralphs JR. 2000. The nucleus of the intervertebral disc from development to degeneration. Am Zool 40:53–
van Dartel M, Hulsebos TJ. 2004a. Amplification and overexpression of genes in 17p11.2~p12 in osteosarcoma. Cancer Genet Cytogenet 153:77–80.
van Dartel M, Hulsebos TJ. 2004b. Characterization of PMP22
expression in osteosarcoma. Cancer Genet Cytogenet 152:113–
Walmsley R. 1953. The development and growth of the intervertebral disc. Edinb Med J 60:341–364.
Widelitz RB, Jiang TX, Murray BA, Chuong CM. 1993. Adhesion
molecules in skeletogenesis. II. Neural cell adhesion molecules
mediate precartilaginous mesenchymal condensations and
enhance chondrogenesis. J Cell Physiol 156:399–411.
Wilson HL, Wilson SA, Surprenant A, North RA. 2002. Epithelial
membrane proteins induce membrane blebbing and interact with
the P2X7 receptor C terminus. J Biol Chem 277:34017–34023.
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