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J Am Soc Nephrol 13:2437-2445, 2002
© 2002 American Society of Nephrology

Connective Tissue Growth Factor and Regulation of the Mesangial Cell Cycle: Role in Cellular Hypertrophy

Nadia Abdel Wahab, Benjamin S. Weston, Terry Roberts and Roger M. Mason

Cell and Molecular Biology Section, Division of Biomedical Sciences, Faculty of Medicine, Sir Alexander Fleming Building, Imperial College, South Kensington, London, United Kingdom.

Correspondence to Dr. Roger M. Mason, Cell and Molecular Biology Section, Division of Biomedical Sciences, Faculty of Medicine, Sir Alexander Fleming Building, Imperial College, South Kensington, London, SW7 2AZ, UK. Phone: 0044-20-7594-3019; Fax: 0044-20-7594-3015;


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
ABSTRACT. Connective tissue growth factor (CTGF) is now considered to be one of the important driver molecules for the pathogenesis of diabetic nephropathy (DN) and possibly many other fibrotic disorders. However, the molecular mechanisms by which CTGF functions remain to be established. In an attempt to define these mechanisms, this study was designed to investigate whether CTGF has any effect on the cell cycle of human mesangial cells (HMC), which are known to undergo hypertrophy in DN. This report provides the first evidence that CTGF is a hypertrophic factor for HMC. CTGF stimulates HMC to actively enter the G1 phase from G0, but they do not then progress further through the cell cycle. The molecular mechanisms underlying this G1 phase arrest appear to be due to the induction of the cyclin-dependent kinase inhibitors (CDKI) p15INK4, p21Cip1, and p27Kip1, which are known to bind and inactivate cyclinD/CDK4/6 and the cyclin E/CDK2 kinase complexes. This could account for the maintenance of pRb protein in a non- or very low-phosphorylated state, preventing cell cycle progression. Using CTGF antisense oligonucleotides, the results also indicate that the previously identified transforming growth factor–{beta} (TGF-{beta})–induced hypertrophy in mesangial cells is CTGF-dependent. Mesangial cell hypertrophy is one of the earliest abnormalities of diabetic nephropathy; therefore, therapeutic strategies targeting CTGF may be beneficial in controlling DN. Email: roger.mason@ic.ac.uk


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
Diabetic Nephropathy (DN) is associated with mesangial cell hypertrophy rather than with cell proliferation (1,2). Cellular hypertrophy is characterized by a halt in the cell cycle at the G1 phase, with the continued production of cellular proteins, which subsequently leads to an increase in the overall size of the cell (3). Previously, cell cycle analysis revealed that prolonged exposure to high concentrations of glucose arrests mesangial cells in the G1 phase with the concomitant induction of hypertrophy. The effect was found to be mediated by the autocrine synthesis and activation of transforming growth factor–{beta} (TGF-{beta}) (4,5). Elevated expression levels of cyclin-dependent kinase inhibitors (CDKI), such as p15, p16, p21, and p27 (6–13), are the main cause of this cell cycle arrest at G1. Glomerular hypertrophy in DN could be considered an early marker for the development of irreversible glomerulosclerosis (3,14,15). Therefore, it is important to understand the pathways leading to mesangial cell growth arrest and hypertrophy. We hypothesized that connective tissue growth factor (CTGF) (CCN2), an immediate early gene encoding a 38-kD cysteine-rich protein and a member of the CCN family of cytokines, may act downstream of TGF-{beta} to mediate mesangial cell hypertrophy.

CTGF appears to be involved in diverse autocrine or paracrine actions in many different cell types (16). It has been reported to have mitogenic activity (17) and to mediate cell adhesion (18), angiogenesis (19), increased cell migration (20,21) and cell survival (22), induction of apoptosis (23), and regulation of gene expression (17,24–27). Its expression in mesangial cells is induced by exposure to high glucose or to TGF-{beta}, and it mediates some of the biologic effects of the latter (27).

There is strong evidence that CTGF plays a key role in the pathogenesis of many fibrotic disorders, including DN (25–28). However, the molecular mechanisms by which CTGF exerts these effects remain to be established. In an attempt to define these mechanisms, we investigated whether CTGF plays a role in mesangial cell hypertrophy.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Cell Cultures, Antibodies, and Reagents
Primary normal adult human mesangial cells (CC-2259, lot 3F1510) were purchased from Biowhittaker (Wokingham, Berkshire, UK) and maintained in culture as described previously (29) and used at passage 9 to 10. Antibodies against Phospho-Rb (Ser 780) and Phospho-Rb (Ser 795), Phospho-Rb (Ser-807/Ser811), Phospho-p53 (Ser 15) 16G8, Phospho-MAPK (Erk1/2) (Thr202/Tyr204), p44/42 MAP kinase were from New England Biolabs (Hitchin, Hertfordshire, UK). The p21 and Rb antibodies were from TCS Biosciences Ltd (Buckingham, UK). Antibodies against p27 (C19) and cyclin D1 (R124) were from Santa Cruz (Santa Cruz, CA). The antibody against p15 was from Serotec (Kidlington, Oxford, UK). rCTGF was expressed in transformed HMC and purified from the medium using Talon metal affinity resin, as reported previously (27). TGF-{beta} was purchased from R & D Systems (Abingdon, Oxfordshire, UK). Phosphothioate antisense (TGG GCA GAC GAA CG) and control oligonucleotides (ACC GAC CGA CGT GT) directed to CTGF were designed and manufactured by Biognostik GmbH (Göttingen, Germany), who own the intellectual property rights to the sequences (27).

Cell Cycle Analysis by Fluorescence-Activated Cell Sorter (FACS)
Cells were trypsinized, washed, collected, and fixed with 70% ethanol for 24 h after treatments. Fixed samples were centrifuged, treated with 100 µg/ml RNase A, and resuspended in 50 µg/ml propidium iodide. Stained cells (20,000 events) were analyzed on a FACS flow cytometer (Becton Dickinson, Franklin Lakes, NJ). The percentage of cells within the G1/S and G2/M phases of the cell cycle were determined by analysis with the CellQuest software provided by Becton-Dickinson.

Determining the Hypertrophy Index
Cells were made quiescent for 2 d in serum-free medium containing 4 mM glucose. The cultures were then treated with growth factors for 24 h, after which the cells were trypsinized, washed twice with phosphate-buffered saline (PBS) and counted using an improved Neubauer hemocytometer. Equal numbers of cells were lysed in RIPA buffer (0.1% [wt/vol] sodium dodecyl sulfate [SDS], 0.5% (wt/vol) sodium deoxycholate, 1.0% (wt/vol) Nonidet P-40, in PBS), and the total protein content measured using the BCA-200 protein assay kit (Pierce, Rockford, IL). Total protein was expressed as micrograms of protein per 104 cells. Experiments were performed independently three times.

Western Blotting
Cells were lysed in reducing SDS-PAGE loading buffer and immediately scraped off the plate. Cell lysates were sonicated for 10 s to shear DNA. Samples were boiled for 5 min and resolved on 4 to 12% gradient gels by SDS-PAGE. Proteins were transferred onto a polyvinylidene difluoride membrane filter (Immobilin-P, Millipore, Bedford, UK) using a BioRad (Hercules, CA) transfer apparatus. Blots were incubated in blocking buffer containing 1x Tris-buffered saline (TBS), 0.1% Tween-20 with 5% (wt/vol) nonfat dry milk for 1 h. Immunodetection was performed by incubating the blots in primary antibody at the appropriate dilution in antibody dilution buffer (1x TBS, 0.1% Tween-20 with 5% bovine serum albumin [BSA]) overnight at 4°C. Blots were then washed three times with washing buffer (1x TBS, 0.1% Tween-20) and incubated with secondary horseradish peroxidase (HRP)–conjugated antibodies for 1 h at room temperature. Bound antibodies were visualized using the enhanced chemiluminescence reagent Luminol (Autogen Bioclear UK Ltd, Wiltshire, UK). Prestained molecular weight standards (Amersham International PLC, Amersham, UK) were used to monitor protein migration.

Immunofluorescence Staining
Cells were fixed with 3.7% paraformaldehyde and permeabilized with 0.5% Triton X-100 in PBS for 10 min at room temperature. Coverslips were then incubated overnight at 4°C with serum (5% in PBS) from the same species as that in which the secondary antibody was raised. They were then incubated with primary antibodies (at optimum dilution in PBS containing 3% BSA) for 1 h at 37°C. Coverslips were then washed and incubated in the dark for 1 h with fluorescein-conjugated secondary antibody (Sigma Aldrich, Dorset, U.K.). After staining, the coverslips were mounted on glass slides with anti-fade mounting media (Vector Labs, Peterborough, UK) and examined using a fluorescence microscope.

Use of Antisense-CTGF Oligonucleotides
A CTGF mRNA antisense oligonucleotide (2 µM, as recommended by the manufacturer) or a CG-matched randomized sequence oligonucleotide (negative control) was added directly to cultures.

Transient Transfection and Reporter Gene Assay
p21 promoter-luciferase reporter gene constructs (30) or the pGL-3 basic vector were transfected by electroporation at a concentration of 15 µg with 5 µg of the pSV-{beta}-Galactosidase Control Vector (Promega, Southampton, UK) into 25 x 106 transformed human mesangial cells (THMC) using conditions described previously (27). Forty-eight hours after transfection, cells were lysed in the reporter lysis buffer (RLB), which permits both luciferase and {beta}-galactosidase assays, using Promega Kits. Luciferase activity was normalized to {beta}-galactosidase activity to correct for any difference in transfection efficiency.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Effect of CTGF on Cell Cycle Distribution in HMC
To investigate whether CTGF has an effect on the cell cycle of HMC, cells were seeded in dishes in equal numbers and left to grow to 50 to 60% confluence in medium containing 10% fetal calf serum (FCS). Cells were then washed twice with PBS buffer and maintained in medium containing 0.2% FCS. After 48 h, the growth-arrested cells were washed and incubated in serum-free medium for 2 h before adding CTGF (80 ng/ml), TGF-{beta} (5 ng/ml), or FCS (10%), after which they were incubated for an additional 24 h. The cell cycle distribution of the HMC was then analyzed by FACS after staining DNA with propidium iodide. Figure 1A demonstrates that after growth arrest by serum deprivation, an average of 91% of HMC was arrested in the G0/G1 phase, 2% in the S phase, and 7% in G2/M phase. Treatment with CTGF for 24 h (Figure 1B) did not alter the cycle distribution (91% of cells was arrested in G0/G1 phase, 3% in S phase, and 6% in G2/M phase), indicating that, in contrast to some other cell types, it is not a mitogen for HMC. This result is very similar to that in Figure 1C, which was obtained when HMC were treated with 5 ng/ml TGF-{beta} (91% of cells was arrested in G0/G1 phase, 3% in S phase, and 6% in G2/M phase). TGF-{beta} has been shown by others to inhibit MC DNA synthesis in response to mitogens (31). In contrast, after 10% serum stimulation for 24 h, the percentage of cells in G0/G1 phase decreased to 58%, whereas 12% were in S phase and 30% in G2/M phase, confirming HMC responsiveness to mitogens



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  		Figure 1. Cell cycle distribution in human mesangial cells (HMC). Growth-arrested HMC (A) were treated with 80 ng/ml connective tissue growth factor (CTGF) (B), 5 ng/ml transforming growth factor–{beta} (TGF-{beta}) (C), or 10% fetal calf serum (FCS) (D). Cells were harvested after 24 h, and their cell cycle distribution was analyzed by FACS after staining DNA with propidium iodide. 2 x 104 HMC were analyzed for each condition. The percentage of cells present in G0/G1, S, and G2/M phases are shown.

 
There was no apparent change in the cell number after 24 and 48 h of treatment with CTGF. However, when the CTGF-treated cells were washed with PBS and then incubated in medium containing 10% FCS, they reverted to the proliferative phenotype, reaching confluence within 3 d. Thus the cells were not irreversibly arrested or senescent after treatment with CTGF.

CTGF Induces Cellular Hypertrophy of HMC
To investigate whether the G0/G1 phase-arrested HMC undergo cellular hypertrophy in response to CTGF, we determined the hypertrophy index (ratio of total protein content to cell number) after 24-h incubation (5). As shown in Figure 2, 80 ng/ml CTGF induced cellular hypertrophy of HMC by 30%. TGF-{beta} (5 ng/ml) induced cellular hypertrophy by 42%, which is in agreement with the findings of Wolf et al. (5). Interestingly, treatment with CTGF antisense oligonucleotide (2 µM) completely abolished TGF-{beta}–dependent hypertrophy, whereas the control CTGF antisense oligonucleotide (2 µM) had no effect. We have shown previously that the markedly increased CTGF mRNA pool in either TGF-{beta}- or high glucose–stimulated HMC is reduced to below basal levels when cultures are treated with CTGF antisense oligonucleotide and that changes in CTGF mRNA levels correlate closely with changes in CTGF protein levels (27).



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  		Figure 2. CTGF induces cellular hypertrophy of HMC. Growth-arrested HMC were treated with CTGF, TGF-{beta}, TGF-{beta} in the presence of CTGF antisense oligonucleotide, or control CTGF antisense for 24 h. The hypertrophy index was determined as described in the Materials and Methods section. The figure shows the results from three independent experiments, which were pooled and expressed as the mean ± SEM. Triplicate or quadruplicate cultures were assessed for each condition in each experiment (total n = 11). Statistical analyses were performed using a two-sample unpaired t test. *P < 0.001.

 
CTGF Regulates Protein Expression of the G1 Phase Cyclins and CDKI in HMC
To understand the molecular mechanisms underlying CTGF-mediated hypertrophy, we carried out experiments in which serum-starved cells were treated with the growth factor for different periods of time and then lysed. The cell lysates were used for Western blotting to assess the expression levels of several key regulatory G1 phase proteins. Figure 3 shows the results obtained for quiescent HMC that were stimulated with 80 ng/ml CTGF for periods up to 24 h. CTGF stimulated the cells to actively enter the cell cycle as is clearly indicated by the increased expression level of cyclin D protein within 30 min of the addition of CTGF (Figure 3A). The Western blot was quantified by densitometry. Cyclin D1 levels were increased by twofold and threefold after 30 min and 2 h, respectively, and remained elevated until 24 h. In contrast, cyclin E was hardly detectable at any time during the experiment (data not shown). CTGF also stimulated increased expression of the cell cycle negative regulators, the CDKI p15INK4, p21Cip1,and p27Kip1 (Figure 3, B, C, and D, respectively).



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  		Figure 3. CTGF regulates protein expression of the G1 phase cyclins and cyclin-dependent kinase inhibitors (CDKI) in HMC. Growth-arrested HMC were treated with CTGF. At the time points indicated, cell extracts were prepared and equal volumes were analyzed by Western blotting, as described in the Materials and Methods section, using anti-cyclin D1(A), anti-p15INK4(B), anti-p21Cip1(C), and anti-p27Kip1 (D) antibodies. Expression of {beta}-actin protein was examined as a quantity loading control (E).

 
Mesangial cells were also synchronized by serum withdrawal for 24 h and then treated with 80 ng/ml rCTGF in the presence of 10% FCS for 24 h. The expression of CDKI was examined after 2 h and at 24 h. Under these conditions, there was no induction of p15INK4, but p27Kip1 was increased at both 2 h and 24 h and p21Cip1 at 24 h, though not so markedly as in quiescent cells (data not shown). Thus, even in the presence of a very strong unphysiologic mitogen such as 10% FCS, rCTGF is still able to induce key regulatory CDKI.

CTGF Does Not Stimulate Phosphorylation of Retinoblastoma Protein (pRb)
The phosphorylation status of the negative cell cycle regulator protein, pRb, can determine whether cells proliferate or hypertrophy in response to growth factors (31–33). Thus we examined whether changes in pRb protein phosphorylation occur on treatment of mesangial cells with CTGF. We used a specific anti-phospho-Rb (Ser 807/Ser 811) antibody to probe Western blots (Figure 4). The results indicate that in serum-deprived quiescent cells phosphorylated pRb is barely detectable. Neither the level of total pRb nor of phospho-pRb changed significantly over the experimental period after the addition of CTGF. In contrast, 10% FCS markedly induced pRb phosphorylation within 12 h of stimulation. Similar results were obtained when we used phospho-Rb (Ser780) and phospho-Rb (Ser795) antibodies (data not shown). This indicates that hyperphosphorylation of pRb, which is required for the S phase transition (32,34) does not occur after treatment of HMC with CTGF.



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  		Figure 4. CTGF does not stimulate phosphorylation of retinoblastoma protein (pRb). Growth-arrested HMC were treated with CTGF or 10% FCS. At the time points indicated, cell extracts were prepared and equal volumes were analyzed by Western blotting, as described in the Materials and Methods section using a specific anti-phospho-Rb (Ser 807/Ser 811) antibody. Expression of {beta}-actin protein was examined as a quantity loading control.

 
When mesangial cells were deprived of serum for 24 h and then treated simultaneously with 10% FCS and 80 ng/ml rCTGF, the latter was not able to suppress phosphorylation of pRb (data not shown). This finding is consistent with the observation that p15INK4 was not induced under these conditions (see above), this CDK1 being an inhibitor of the cyclin D1/cdk4/6 complex, which hypophosphorylates pRb (see Discussion).

CTGF Induces MAP Kinase (ERK)
The classical MAP kinase (ERK) has been reported to be a critical mediator of the cell-cycle machinery, particularly at the G1 phase. Transcription of the cyclin D1 gene as well as the cyclin D1 protein level has been shown to be regulated by the MAPK cascade (35,36). Direct phosphorylation of the CDKI p27Kip1 by MAP kinase increases the stability of the protein (37). Activation of the ERK MAP kinase pathway has also been implicated with the increased transcription of p21Cip1 (38,39). We therefore investigated whether CTGF activates the MAPK pathway. The results show that CTGF indeed activates the MAPK pathway in HMC within 5 min of exposure to the growth factor and that this activity is sustained for 24 h (Figure 5A). Activation and nuclear translocation of the MAP kinase was also confirmed by immunofluorescence (Figure 5B).



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  		Figure 5. Activation of the MAP kinase (ERK) pathway by CTGF. Growth-arrested HMC were treated with CTGF. (A) At the times indicated, cell extracts were prepared and equal volumes were analyzed by Western blotting, as described in the Materials and Methods section, using a specific anti-phospho-MAP kinase (Thr202/Tyr 204) antibody and an antibody against total MAP kinase. Expression of {beta}-actin was examined as a quantity loading control. (B) In parallel experiments, cells were fixed, permeabilized, and immunostained using the same anti-phospho-MAP kinase antibody, as described in the Materials and Methods section. (a) Growth-arrested cells; (b, c, and d) cells stimulated with 80 ng/ml CTGF for 5 min, 30 min, and 2 h, respectively.

 
Modification and Stabilization of p53 by CTGF in HMC
The p53 protein is a stress-responsive transcription factor that is induced by a variety of stimuli, which act through mechanisms that stabilize the protein posttranscriptionally, including phosphorylation (40). It can initiate cell cycle arrest at the G1/S transition or apoptosis. Induction of a G1/S block by p53 is due partly to its ability to transactivate the CDKI p21Cip1 gene (41). Phosphorylation of p53 at Ser15 is critical for the activation of this protein (42,43), its stabilization and nuclear accumulation (44,45), and its interaction with the transcriptional coactivators CBP/p300 and PCAF (46). Therefore, we examined the phosphorylation status of p53 at Ser15 in HMC after exposure of quiescent cells to CTGF for different periods of time. Figure 6A reveals that CTGF induces the phosphorylation of p53 at Ser15 with maximum induction after 2 h of exposure. This time appears to be similar to that required to bring about the maximum accumulation of p21 (Figure 3). Nuclear accumulation of phosphorylated p53 at Ser15 was also detected by immunofluorescence (Figure 6B). The presence of 10% FCS abolished the CTGF-induced phosphorylation of p53 (data not shown). However, serum supplemented cultures contained only very low levels of total p53 (data not shown), presumably because the nonphosphorylated protein is unstable and rapidly removed (40).



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  		Figure 6. CTGF induces phosphorylation of p53 protein in HMC. Growth-arrested HMC were treated with CTGF. (A) At the times indicated, cell extracts were prepared and analyzed by Western blotting, as described in the Materials and Methods section, using a specific anti-phospho p53 (Ser-15) antibody. (B) In parallel experiments, cells were fixed, permeabilized, and immunostained using the same anti-phospho p53 (Ser-15) antibody, as described in the Materials and Methods section. (a) Growth-arrested cells; (b and c) cells treated with 80 ng/ml CTGF for 30 min and 24 h, respectively.

 
To determine whether CTGF induces transcription of the p21Cip1 gene in quiescent HMC in a p53-dependent manner, we used the murine p21 promoter-luciferase reporter gene. Constructs prepared by Xiao et al. (30) were used. The human, rat, and mouse p21 promoters all have two p53 binding site motifs (47). The constructs used in this study consisted of the wild-type construct, which contains two p53 binding sites situated near nt-2800 and nt-1900 relative to the TATA box; mutant 1 where the p53 binding site near nt-2800 was mutated; and mutant 2 where the p53 binding site near nt-1900 was mutated. The relative activity of the p21 promoters in driving luciferase transcription in the absence or presence of 80 ng/ml CTGF was tested. Figure 7 summarizes the results of three independent experiments. As shown in the figure, CTGF led to a twofold enhancement of basal activity of the wild-type promoter. Mutating the first p53 binding site near nt-2800 resulted in a 60% reduction of the basal activity of the wild-type p21 promoter. CTGF-enhanced activity was also abolished. In contrast, mutating the second p53 binding site near nt-1900 had no significant effect on either the basal activity of the promoter or on the enhanced activity induced by CTGF.



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  		Figure 7. Effect of CTGF on p21 promoter activity. A p21 promoter-reporter construct (wild-type), promoterless vector (pGL-3), or p21 promoter-reporters in which the p53 binding sites had been deleted (mutant 1, mutant 2), were transiently cotransfected with the pSV-{beta}-galactosidase control vector into a mesangial cell line by electroporation, as described in the Materials and Methods section. The cultures were incubated in medium with ({blacksquare}) or without ({image}) 80 ng/ml CTGF. Luciferase activity was assayed 48 h after transfection and 18 h after the addition of CTGF and was normalized to the {beta}-galactosidase activity to correct for any difference in transfection efficiency. The results represent the mean ± SEM of three independent experiments (three replicates for each experiment). Statistical analyses were used to compare samples incubated with or without CTGF. The analyses were performed using a two-sample unpaired t test. *P < 0.001.

 
CTGF Mediates TGF-{beta}–Induced Expression of CDKI in HMC
Finally, we investigated whether CTGF mediates TGF-{beta}–dependent hypertrophy by regulating the expression of the CDKI of the G1 phase. We carried out experiments in which cells were treated with 5 ng/ml TGF-{beta} in the presence or absence of CTGF antisense or control antisense oligonucleotides (2 µM). Cells were then lysed, and cell lysates were used for Western blotting to assess the expression levels of p15INK4, p21Cip1, and p27Kip1 proteins. Figure 8 shows that TGF-{beta} increased the expression level of all three CDKI after 24 h. This increased expression was abrogated in the presence of CTGF antisense oligonucleotide but not in the presence of control CTGF antisense oligonucleotide.



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  		Figure 8. CTGF mediates TGF-{beta}–induced expression of CDKI in HMC. Growth-arrested HMC were treated with TGF-{beta} or with TGF-{beta} in the presence of CTGF antisense oligonucleotide or control CTGF antisense for 24 h. Cell extracts were prepared, and equal volumes were analyzed by Western blotting, as described in the Materials and Methods section, using anti-p15INK4 (A), anti-p21Cip1 (B), and anti-p27Kip1 (C) antibodies. Expression of {beta}-actin protein was examined as a quantity loading control (D).

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
CTGF is now considered to be an important driver molecule for the pathogenesis of DN, and our previous work showed its likely involvement as one of the pathways stimulating the deposition of extracellular matrix around mesangial cells exposed to high-glucose conditions (27). Our major goal is to understand the molecular mechanisms through which this growth factor functions. This will undoubtedly help in designing specific drugs to control fibrosis in the diabetic kidney and possibly in many other fibrotic disorders. In the present study, we investigated whether CTGF has an effect on the HMC cell cycle, as cell cycle analysis can be informative as to whether growth factors have proliferative or hypertrophic effects on specific cell types.

When stimulated with growth factors, quiescent cells (G0) enter the cell cycle (48,49) at the G1 phase, where they increase their size and exhibit enhanced RNA and protein synthesis. After passing a restriction point in late G1, they become committed to enter the S phase and are no longer responsive to growth factors. DNA replication occurs in the S phase. Cells then progress through G2 and enter mitosis, where they divide. Cell proliferation requires normal progression through the cell cycle while cell hypertrophy occurs when the cell cycle is arrested in G1 (50,51). In contrast, apoptosis occurs when cells exit from the cell cycle, typically in G1 (52).

The cell cycle is controlled by positive and negative regulators. The positive regulators are cyclin/cyclin-dependent kinase (CDK) complexes (53–56). Cell cycle entry in mammalian cells requires the synthesis of the D-type cyclins (D1–3) that associate with and activate CDK4/6 (54). This in turn hypophosphorylates pRb, the retinoblastoma protein, a key regulator of events in G1 (see below). The negative regulators are CDK inhibitors (CDKI), which are grouped into two families (57). The INK4 family (p15, p16, p18, p19, and p20) only bind to the cyclin D/CDK4/6 complex and inhibit its activity, whereas the CIP/KIP family (p21Cip1, p27Kip1, and p57Kip2) bind to and inhibit cyclin-CDK complexes containing either CDK4/6 or CDK2. Our results show that CTGF induces cyclin D1 levels, but also simultaneously p15INK4, p21Cip1, and p27Kip1. p21 transcription was induced by CTGF via a phospho p53-dependent mechanism.

It seems likely that mesangial cells exposed to CTGF arrest in the early G1 phase of the cell cycle. pRb exists in three different states of phosphorylation—a non-phosphorylated state, which is inactive and associated with G0 cells, a hypophosphorylated state, which acts as a transcriptional repressor of E2F-regulated genes in early G1, and a hyperphosphorylated state which dissociates from E2F and allows transcription of genes required for late G1 and S phase of the cell cycle (58,59). Hyperphosphorylation of pRb is driven by cyclinE-CDK2, which is activated during transition across the restriction point in the late G1 phase (58). Cyclin E was barely detected in CTGF-treated HMC, and there was no evidence for increased phosphorylation of pRb in these cells, whereas phosphorylation occurred in FCS-stimulated cells. Thus CTGF-treated HMC do not enter late G1 phase. However, exposure to CTGF does lead to a rapid increase in cyclin D1 levels, indicating entry into the G1 phase. This rapid increase is likely to be due to protein stabilization after phosphorylation by MAPK.

Cyclin D1/CDK4/6 complex is active throughout the G1 phase, and one of its roles is to hypophosphorylate pRb in early G1 (59). However, we observed no measurable increase in phosphorylation of pRb in CTGF-treated cells compared with that in serum-deprived HMC. This may be due to the very early induction in CTGF-treated cells of p15INK4, a specific inhibitor of cyclin D activation (57). It is noteworthy that protein transduction of the related p16INK inhibitor of CDK4/6 into mitogen-stimulated lymphocytes completely blocked phosphorylation of pRb in early G1 phase, whereas control cells contained hypophosphorylated pRb (59). Thus our data are consistent with arrest in early G1 phase when HMC are exposed to CTGF.

Cyclin D1 does not appear to be essential for the development of most tissues, as demonstrated by the phenotype of knockout mice (60). However, it has been proposed that it has a role in promoting cell growth. Studies on Drosophila provide convincing evidence for this in that organism (61), but it has also been proposed that renal epithelial cell hypertrophy depends on cyclin D kinase activation to stimulate protein synthesis as cells enter the cycle without subsequent cyclin E kinase activation (62). The connecting pathways to protein synthesis are not understood, but if a similar mechanism is operative in bringing about hypertrophy of CTGF-arrested cells, it must involve some kinase activity that is subtly different from that responsible for hypophosphorylation of pRb which, as discussed above, appears to be inhibited.

Prolonged exposure to high glucose or to TGF-{beta} have been found to arrest mesangial cells in the G1 phase with concomitant cellular hypertrophy (3,4,63,64). Indeed, high glucose-induced hypertrophy was found to be mediated by TGF-{beta} because it could be prevented by neutralizing antibodies to the molecule (4). Treatment with TGF-{beta}–neutralizing antibodies was also shown to attenuate mesangial cell hypertrophy in diabetic mice (65). Both in vivo and in vitro studies indicate that the molecular mechanisms underlying this G1-phase arrest involve the overexpression of p21Cip1 and p27Kip1 proteins and absence of phosphorylation of pRb (Table 1). CTGF antisense, but not missense, oligonucleotides abolished TGF-{beta}–induced hypertrophy and abrogated TGF-{beta}–dependent expression of p15INK4, p21Cip1, and p27Kip1; this suggests that TGF-{beta}–dependent G1 phase arrest and induced hypertrophy is mediated through CTGF. Although CTGF has been reported to be pro-apoptotic for MC (66) we did not observe this in our experiments that used a concentration of 80 ng/ml. Moreover, it has been shown that induction of p27, a downstream effector of CTGF, limits apoptosis in mesangial cells and fibroblasts in vitro (67).


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  		Table 1. Hypertrophic factors for HMCa
 
Transient transfection experiments with a p21 promoter-luciferase-reporter gene indicated that CTGF enhances the expression of p21 in THMC cells. This occurs in a p53-dependant manner via the first p53 binding site of the promoter. This site, unlike the second p53 binding site, has two copies of the consensus decamer motif 5'-RRRCWWGYYY-3', where R = G or A, W = T or A, and Y = C or T (68). In agreement with this result, CTGF also induces the phosphorylation of the transcription factor p53 at Ser15 and its nuclear accumulation in HMC. This mechanism may be also significant in mediating the regulation of other genes by CTGF such as TSP-1 (69), MMP-2 (70), and PAI-1 (27), which have two or more p53 binding sites in their promoters (71).

In summary, our results strongly indicate that CTGF is a hypertrophic factor for HMC. CTGF stimulates these cells to actively enter the G1 phase from G0 but not to progress further through the cell cycle. The molecular mechanisms underlying this G1 phase arrest appear to be due to the induction of the CDKI, p15INK4, p21Cip1, and p27Kip1, which subsequently bind to and inactivate cyclinD/CDK4/6 and cyclin E/CDK2 kinase complexes. The inhibition of these G1-phase cyclin/CDK complexes results in very low or no phosphorylation of the pRb protein leading to cell cycle arrest. Finally, the results of experiments using CTGF antisense oligonucleotides indicate that CTGF functions as a downstream mediator of TGF-{beta} hypertrophic activity in mesangial cells. Interestingly, CTGF has also been shown to function as a downstream mediator of TGF-{beta} mitogenic activity in NRK fibroblasts in suspension cultures (72) by controlling cell cycle progression through the G1/S phase. Under these conditions, CTGF induces the level of cyclin A activity via the reduction of p27Kip1 levels, which results in the hyperphosphorylation of pRb and release of E2F.


   Acknowledgments
 
We thank the Medical Research Council (UK) for the financial support. We are grateful to Dr. Hengyi Xiao (Anagahora, Shimoshidami, Moriyamaku, Japan) for making the p21Cip1 reporter constructs available to us.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Received for publication February 27, 2002. Accepted for publication June 16, 2002.




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