Epithelial Membrane Protein 1 Inhibits Human Spinal Chondrocyte Differentiation.код для вставкиСкачать
THE ANATOMICAL RECORD 294:1015–1024 (2011) Epithelial Membrane Protein 1 Inhibits Human Spinal Chondrocyte Differentiation ZHI-YONG LI,1 SHAO-HU XIONG,1 MING HU,2 AND CHUAN-SEN ZHANG1* Department of Human Anatomy, Second Military Medical University, Shanghai, China 2 Department of Orthopedics, No. 309 Hospital of the Chinese People’s Liberation Army, Beijing, China 1 ABSTRACT 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). C 2011 WILEY-LISS, INC. V 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: firstname.lastname@example.org Received 5 April 2010; Accepted 9 March 2011 DOI 10.1002/ar.21395 Published online 29 April 2011 in Wiley Online Library (wileyonlinelibrary.com). 1016 LI ET AL. (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 immunohistochemistry. MATERIALS AND METHODS 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 EMP1 INHIBITS CHONDROCYTE DIFFERENTIATION TABLE 1. Primer sequences Gene Collagen I Collagen II Collagen X Sox9 Aggrecan Runx2 GAPDH Analysis of Apoptosis and Cell Cycle Primer 0 1017 0 For: 5 -CACACGTCTCGGTCATGGTA-3 Rev: 50 -AAGAGGAAGGCCAAGTCGAG-30 For: 50 -CGGCTTCCACACATCCTTAT-30 Rev: 50 -CTGTCCTTCGGTGTCAGGG-30 For: 50 -GTGGACCAGGAGTACCTTGC-30 Rev: 50 -CATAAAAGGCCCACTACCCA-30 For: 50 -GTAATCCGGGTGGTCCTTCT-30 Rev: 50 -GACGCTGGGCAAGCTCT-30 For: 50 -GCGAGTTGTCATGGTCTGAA-30 Rev: 50 -TTCTTGGAGAAGGGAGTCCA-30 For: 50 -ATACTGGGATGAGGAATGCG-30 Rev: 50 -ACAGTAGATGGACCTCGGGA-30 For: 50 -AAGGTGAAGGTCGGAGTCAAC-30 Rev: 50 -GGGGTCATTGATGGCAACAATA-30 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. 1018 LI ET AL. Statistical Analysis Differences between groups were analyzed using the Student t test. P values of <0.05 were considered statistically signiﬁcant. RESULTS EMP1 Promotes the Proliferation and Viability of Cultured Human Fetal Nucleus Pulposus Cells 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). EMP1 INHIBITS CHONDROCYTE DIFFERENTIATION 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 1019 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. 1020 LI ET AL. 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- EMP1 INHIBITS CHONDROCYTE DIFFERENTIATION 1021 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). DISCUSSION 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 1022 LI ET AL. 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. EMP1 INHIBITS CHONDROCYTE DIFFERENTIATION 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 1023 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. ACKNOWLEDGEMENTS This work was supported by fund from the National Natural Science Foundation of China for Distinguished Young Scholars (30400453). LITERATURE CITED Archer CW, Rooney P, Wolpert L. 1982. Cell shape and cartilage differentiation of early chick limb bud cells in culture. Cell Differ 11:245–251. 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. 1024 LI ET AL. 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 16:5316–5333. 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 157:72–80. 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 270:28824–28833. Urban JPG, Roberts S, Ralphs JR. 2000. The nucleus of the intervertebral disc from development to degeneration. Am Zool 40:53– 61. 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– 118. 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.