Connective Tissue Growth Factor and IGF-I Are Produced by Human Renal Fibroblasts and Cooperate in the Induction of Collagen Production by High Glucose

The human renal fibroblast cell line TK173 (28) was maintained in Dulbecco’s modified Eagle’s medium (DMEM) supplemented with 10% heat-inactivated FCS (ΔFCS), 100 IU/ml penicillin, and 100 μg/ml streptomycin (all purchased from Gibco/Invitrogen, Paisley, Scotland). TK173 cells expressing a truncated form of the TGF-β type II receptor (TK173-ΔTβRII) were previously described (8) and were maintained under the same conditions with the addition of hygromycin B (0.1 mg/ml; Boehringer Mannheim, Mannheim, Germany).

For experimental purposes, cells were seeded at a density of 50,000 cells/well in 48-well plates (Costar, Corning, NY), grown until confluence, and starved in DMEM with 0.5% ΔFCS. After 24 h, cells were cultured in 0.3 ml of DMEM containing 5.5 mmol/l d-glucose supplemented with 0.5% ΔFCS, 50 μg/ml ascorbic acid, and 100 μg/ml β-aminopropionitrile to promote collagen synthesis and prevent cross-links, respectively (both purchased at Sigma Chemical Co., St. Louis, MO). Cells were exposed to 25 mmol/l d-glucose (Merck, Darmstadt, Germany), 25 mmol/l l-glucose (Sigma), recombinant human TGF-β1 (R&D Systems, Abingdon, U.K.), recombinant human IGF-I (Roche Diagnostics, Mannheim, Germany), or recombinant human CTGF (FibroGen, South San Francisco, CA) at concentrations indicated. Blocking antibody studies were performed by the addition of a human neutralizing anti-CTGF monoclonal antibody (mAb; FibroGen) (29), a neutralizing pan-specific anti–TGF-β mAb (30), or a control IgG at the start of the culture. After 4 days, culture supernatants were harvested and measured for secreted proteins. Cells were trypsinized, and viable cells were counted using trypan blue exclusion.

CTGF was measured using a sandwich enzyme-linked immunosorbent assay (ELISA) specific for the NH₂-terminal part of CTGF (FibroGen). Microtiter plates were coated overnight at 4°C with capture antibody (10 μg/ml) in 100 μl of coating buffer (0.05 mol/l sodium bicarbonate, pH 9.6). After blocking the plates with 1% BSA in PBS for 1 h at room temperature and washing with PBS/0.05% Tween, a 50-μl sample was added to each well together with 50 μl of biotinylated monoclonal anti-human CTGF detection antibody diluted in assay buffer (0.05 mol/l Tris [pH 7.8], 0.1% BSA, 4 mmol/l MgCl₂, 0.2 mol/l ZnCl₂, 0.1% Na Azide, 50 mg/l sodium heparin, and 0.1% Triton X-100). After 2 h of incubation at 37°C, plates were washed and incubated with 100 μl of Streptavidin-conjugated alkaline phosphatase for 1 h at room temperature. The ELISA was developed with p-nitrophenyl phosphate (1.5 mg/ml; Sigma) in diethanolamine buffer (1 mol/l diethanolamine, 0.5 mmol/l MgCl₂, 0.02% Na azide) and read at an optical density (OD) of 405 nm. Purified recombinant human CTGF (FibroGen) was used as a standard.

Collagen type I and III accumulation in supernatants was measured using a specific inhibition ELISA as previously described (8). Briefly, 96-well Nunc Maxisorb microtiter plates (Gibco/Invitrogen) were coated with 0.5 μg/ml human collagen type I or III (both purchased from Sigma) in PBS (100 μl/well) overnight at room temperature. Collagen type I or III standard or samples were preincubated overnight at 4°C with goat–anti-human collagen type I or goat–anti-human collagen type III, respectively (both purchased from Immunologicals Direct, Oxfordshire, U.K.) in PBS/0.05% Tween/2% casein (PTC). After a blocking step with PTC for 1 h at 37°C, 100 μl of preincubated standard or sample was added onto the plates and incubated at 37°C for 1 h. After washing the plates, a peroxidase-conjugated secondary rabbit–anti-goat IgG (Nordic Immunology, Tilburg, the Netherlands) was added and incubated for 1 h at 37°C. The ELISA was developed with 2,2′-amino-bis-3-ethylbenzthiazoline-6-sulfonic acid/H₂O₂ and read at OD 415 nm.

Total cellular RNA was extracted using RNAzolB (Campro Scientific, Veenendaal, Netherlands) according to the manufacturer’s description. The quantity and the purity of the isolated RNA were measured at OD₂₆₀ and OD₂₈₀ and analyzed on a 0.5× TBE (0.045 mol/l Tris borate, 0.001 mol/l EDTA) 1% agarose gel to check the integrity of the RNA.

For RT-PCR, 1 μg of total RNA was reverse-transcribed into cDNA by oligo dT priming using Moloney Murine Leukemia Virus reverse transcriptase (all purchased from Invitrogen). The cDNA was amplified by PCR using primers as described in Table 1. Each PCR reaction was performed in a total volume of 40 μl, containing 10 mmol/l Tris.Cl (pH 8.3), 50 mmol/l KCl, MgCl₂ (see Table 1 for concentrations), 0.06 mg/ml BSA, 0.25 mmol/l dNTPs, 25 pmol forward primer, 25 pmol reverse primer (all primers were purchased from Invitrogen), and 0.8 units AmpliTaq (Perkin Elmer, Foster City, CA). Amplification of cDNA started with 5 min of denaturation at 95°C, followed by PCR cycles of 95°C for 1 min, subsequent annealing for 1 min at a primer-specific temperature, and 72°C for 1 min (the number of cycles and the annealing temperatures are summarized in Table 1). The final extension was performed at 72°C for 5 min. All PCR reactions were performed in a PTC-200 DNA engine (MJ Research, Waltham, MA) thermal cycler. PCR products were analyzed on a 0.5× TBE 1% agarose gel. The intensity of bands was determined by densitometry, using EagleSight software (Stratagene, La Jolla, CA).

Real-time PCR was performed in a Roche LightCycler using 1 μl of undiluted cDNA as template in a total reaction volume of 10 μl, containing 0.5 μmol/l of each forward and reverse primer (Table 1), 3 mmol/l MgCl₂, and 1 μl of FastStart DNA Master SYBR Green I reagent (all purchased from Roche Diagnostics). The temperature profile was as follows: an initial denaturation step for 6 min at 95°C, followed by 40 amplification cycles, each consisting of 15 s at 95°C, annealing for 5 s at 64°C, and elongation for 10 s at 72°C, each of these steps with ramp rates of 20°C/s. SybrGreen fluorescence was measured at the end of each elongation phase. Immediately after the last amplification cycle, a melting curve was recorded as follows: after denaturation at 95°C and rehybridization during 15 s at 65°C, the samples were slowly heated to 95°C with a ramp rate of 0.1°C/s with continuous acquisition of SybrGreen fluorescence. One sample was used in each PCR experiment as a standard, of which the cDNA was used at 2-, 4-, 8-, and 16-fold dilutions. The mRNA levels of TBP, encoding for TATA-box binding protein, were measured as a reference to correct for variable input.

Anti-CTGF mAb-coated Sepharose beads were incubated with CTGF for 1 h at 37°C. After three washing steps with PBS, IGF-I was added for 2 h at 37°C. As controls, either CTGF or IGF-I was incubated with anti-CTGF mAB-coated Sepharose beads for 1 h at 37°C only, without the secondary incubation step. After the last incubation step, Sepharose beads were washed again three times with PBS. The beads were resuspended in 20 μl of PBS, and bound proteins were denatured in 3× SDS sample buffer (New England Biolabs, Beverly, MA) by boiling and separated under reducing conditions by SDS-PAGE. The proteins were semidry electroblotted on polyvinylidene fluoride membranes (Immobilon-p; Millipore, Bedford, MA). Membranes were blocked with PTC and then probed with a biotinylated anti-human IGF-I antibody (R&D Systems). After incubation with peroxidase-conjugated streptavidin (Zymed Laboratories, San Francisco, CA), detection was performed with Supersignal (Pierce) and the blots were exposed to Hyperfilm films (Amersham Pharmacia Biotech, Buckinghamshire, U.K.).

The data are presented as means ± SD of triplicate cultures and are representative of at least three independent experiments. Differences between various culture conditions were evaluated by ANOVA with a Bonferroni correction for multiple comparison. In some experiments, relative differences were tested with a one-sample t test. A value of P < 0.05 was considered to represent a significant difference.

Exposure of human renal fibroblasts to hyperglycemic conditions resulted in the upregulation of the mRNA expression of both the α1 subunit of collagen type I (COL1A1) and type III (COL3A1; Fig. 1A). This increase was not due to the higher osmolarity to which the cells were exposed because there were no effects on the COL1A1 and COL3A1 mRNA expression when cells were exposed to 25 mmol/l l-glucose. Exposure to the profibrotic cytokine TGF-β1 also increased COL1A1 mRNA expression. In contrast, COL3A1 mRNA expression was not upregulated by TGF-β1.

The increased transcription of COL1A1 mRNA by high glucose and exogenous TGF-β1 was reflected by an increase in collagen type I protein secretion (Fig. 1B, left). Collagen type III protein secretion was increased by high glucose but not by TGF-β1 (Fig. 1B, right), thus following the same pattern as the induction of COL3A1 mRNA expression. Because collagen type III can be induced by glucose independent of TGF-β1, we also investigated whether TGF-β was involved in glucose-induced collagen type I secretion by renal fibroblasts. For this purpose, neutralizing anti–TGF-β antibodies were added to fibroblast cultures, which completely inhibited the stimulatory effect of TGF-β1 on collagen type I secretion. In contrast, glucose-induced collagen type I was not attenuated (Fig. 1C). In addition, TK173-ΔTβRII was exposed to either TGF-β1 or high glucose. We recently demonstrated that glucose-induced collagen type III was not affected in these transfected cells (8). Now we also show that collagen type I secretion induced by high glucose was not affected in TK173-ΔTβRII, whereas the effect of TGF-β1 was completely abolished (Fig. 1D). Thus, glucose-induced upregulation of collagens type I and III in human renal fibroblasts seems to be independent of endogenous TGF-β1. This observation prompts the question of which alternative growth factors might be involved.

CTGF has been demonstrated to be an important protein in mediating fibrotic processes (18,31,32). Because it has been demonstrated that CTGF is induced in mesangial cells under hyperglycemic conditions (14–16), we investigated the regulation of CTGF expression in renal fibroblasts. Stimulation with TGF-β1, a known inducer of CTGF, resulted in an increase of CTGF mRNA expression as shown by real-time PCR (Fig. 2A, left). Also, exposure to high glucose resulted in an increased CTGF expression, however, with altered kinetics. Although TGF-β1 showed an early response at 24 h, glucose-induced CTGF expression became apparent only after 72 h of stimulation (Fig. 2A, right).

We next investigated the regulation of CTGF at the protein level. Stimulation with either TGF-β1 or high glucose resulted in increased secretion of CTGF protein (Fig. 2B). Furthermore, when fibroblasts were incubated with TGF-β1 in medium containing high glucose, an additive increase in CTGF protein secretion was observed. This suggestion that TGF-β1 and glucose are independent stimulators of CTGF expression was confirmed by the induction of CTGF in TK173-ΔTβRII upon exposure to glucose (Fig. 2C).

Because CTGF was increased in renal fibroblasts exposed to high glucose, the role of endogenous CTGF in glucose-induced collagen secretion was investigated by blocking the effect of CTGF. Addition of a neutralizing anti-CTGF mAb to fibroblast cultures had no effect on the basal levels of collagen secretion. However, when this neutralizing antibody was added to high-glucose cultures, the effect of high ambient glucose on collagen type I secretion was partially inhibited (49.6 ± 23.0%; Fig. 3, left). In addition, glucose-induced collagen type III secretion by renal fibroblasts was partially blocked (53.5 ± 1.8% inhibition; Fig. 3, right). These data demonstrate that endogenous CTGF partially mediates glucose-induced collagen type I and III secretion in human renal fibroblasts.

It is interesting that addition of exogenous recombinant CTGF at a concentration of 50 ng/ml to normoglycemic cultures had no effect on collagen type I and III levels (Fig. 3). Moreover, exposure of renal fibroblasts to CTGF at concentrations ranging from 12.5 to 200 ng/ml had no effect on either collagen (Fig. 4). These results suggest that CTGF might exert its effects on collagen type I and III accumulation only in combination with another factor that seems to be absent in normoglycemic cultures of fibroblasts.

IGF-I seems to be an interesting candidate because it can be induced by hyperglycemia (33), it can function independent of TGF-β1 (34), and a functional collaboration with CTGF has been shown in rat fibroblasts (17). In nonstimulated fibroblasts, there were no detectable levels of IGF-I mRNA. However, IGF-I mRNA expression was induced in fibroblasts upon stimulation with either high glucose or TGF-β1 (Fig. 5A). In TK173-ΔTβRII cells, the TGF-β1–induced upregulation of IGF-I mRNA expression was completely inhibited, whereas the glucose-induced increase in IGF-I was still intact. Therefore, glucose-induced IGF-I seemed to be independent of TGF-β signaling.

Next, fibroblasts were stimulated with recombinant IGF-I to investigate the effect on collagen type I and III secretion. Collagen type I secretion was not affected by IGF-I (Fig. 5B). In contrast, collagen type III was already significantly and strongly upregulated by IGF-I at a concentration of 6.25 ng/ml, reaching a maximum at 25 ng/ml (Fig. 5C).

Next, fibroblasts were stimulated with a combination of CTGF and IGF-I. Although both CTGF and IGF-I individually had no effects on collagen type I accumulation, the combination of both factors resulted in a synergistic induction (Fig. 6A, left). Moreover, collagen type III secretion was further enhanced by the addition of CTGF and IGF-I (Fig. 6B, left).

Because CTGF and IGF-I have synergistic effects on collagen secretion in normoglycemic fibroblast cultures and both growth factors are induced upon glucose stimulation, the effects of CTGF and IGF-I were also investigated under hyperglycemic conditions. Stimulation with CTGF in high-glucose medium did not result in a further increase in collagen type I and III secretion compared with incubations with high glucose alone (Fig. 6A and B, right). In contrast, exposure to combinations of IGF-I and high glucose resulted in enhanced collagen type I and III secretion when compared with the individual stimuli. The synergistic effects of CTGF and IGF-I on collagen secretion were not observed under hyperglycemic conditions. This was probably because plateau levels were reached in these conditions. When lower concentrations of growth factor were used, which have only marginal effects under normoglycemic conditions, there was indeed an enhanced secretion of collagens type I and III under hyperglycemic conditions (data not shown).

To try to elucidate the mechanism of the synergy between CTGF and IGF-I, we examined the effect of these growth factors on each other’s expression. CTGF mRNA expression by renal fibroblasts was decreased by IGF-I, even in the presence of exogenous CTGF (Fig. 7A). The downregulation in CTGF mRNA was reflected by a decreased secretion of CTGF protein by renal fibroblasts (Fig. 7B). Although CTGF protein secretion was slightly decreased by exogenous CTGF (Fig. 7B), this was not reflected by a negative feedback on its own mRNA expression (Fig. 7A). In addition, IGF-I mRNA expression was measured in renal fibroblasts after CTGF and IGF-I stimulation. Stimulation with either growth factor did not have any clear effects on IGF-I mRNA expression (data not shown). These results suggest that the synergistic effects of CTGF and IGF-I are not mediated by the increase in endogenous CTGF and IGF-I.

Another possible mechanism of the synergy between CTGF and IGF-I might be the direct binding between the two growth factors. With the use of anti-CTGF mAb-coated Sepharose beads, the captured CTGF was able to bind IGF-I, which was demonstrated in an IGF-I Western blot (Fig. 7C). When only CTGF or IGF-I was incubated with the Sepharose beads, no IGF-I bands could be detected. These results confirm that CTGF has the ability to bind IGF-I.

In the present study, we investigated the role of CTGF and IGF-I in the induction of collagen type I and III secretion by high glucose in human renal fibroblasts. We have demonstrated that both CTGF and IGF-I expression are increased in human renal fibroblasts cultured under hyperglycemic conditions, also in the absence of TGF-β signaling. Exogenous IGF-I had a pronounced effect on the secretion of collagen type III but not on collagen type I. Although CTGF alone had no effect on the expression of either collagen type I or type III, when combined with IGF-I, an enhanced induction of both matrix components was observed. Furthermore, glucose-induced collagen accumulation was partially blocked by neutralizing anti-CTGF antibodies and further enhanced by exogenous IGF-I. These results indicate a role for both CTGF and IGF-I in glucose-induced matrix accumulation by human renal fibroblasts, which might involve the cooperation between CTGF and IGF-I, a mechanism that might contribute to the pathogenesis of diabetic nephropathy.

One of the most potent mediators of fibrosis is undoubtedly TGF-β, which has been studied extensively. It has been shown that TGF-β plays a central role in the development of diabetic nephropathy (7). In the present study, we have shown that glucose-induced collagens type I and III can occur independent of endogenous TGF-β1. Our finding seems to be in contrast to the findings by Han et al. (35), who demonstrated that glucose-induced collagen type I was TGF-β–mediated in murine fibroblasts. These results were obtained using fibroblasts from mice, whereas we used fibroblasts from human origin, which might contribute to the observed differences. More recently, other growth factors, such as CTGF and IGF-I, have also been implicated to contribute to the pathogenesis of diabetic nephropathy (9–11). Previous studies have shown that CTGF expression was increased in various models of experimental diabetic kidney disease (15,–17), as well as in human diabetic nephropathy (18). Concerning IGF-I, a correlation between increased renal IGF-I levels and diabetic nephropathy has been found (23–26).

Recently, we demonstrated a TGF-β1–independent mechanism of fibronectin and collagen type III upregulation in human renal fibroblasts by high glucose, in a process that might involve alternative growth factors (8). In the present study, we have shown that high glucose increased CTGF expression in renal fibroblasts. Moreover, using fibroblasts expressing a dominant negative TGF-β type II receptor, we could demonstrate TGF-β–independent CTGF induction. Several groups have reported the involvement of CTGF in glucose-induced ECM expression in mesangial cells (14–16). We now demonstrate for the first time that endogenous CTGF is also involved in glucose-induced collagen accumulation by human renal fibroblasts because this effect can be partially blocked by neutralizing anti-CTGF antibodies. However, surprisingly, we could not observe any effects of recombinant CTGF on collagen production by renal fibroblasts. This finding is in agreement with a previous study that showed no effect of recombinant CTGF on collagen type III mRNA expression in NRK cells (17), although others have found direct effects of CTGF on ECM accumulation by fibroblasts from other origins (31,36,37). Next to CTGF, we have identified IGF-I as a growth factor involved in the regulation of matrix accumulation by renal fibroblasts. We have demonstrated that IGF-I mRNA expression was increased in renal fibroblasts by high glucose, independent of endogenous TGF-β. In addition, IGF-I is a potent inducer of collagen type III secretion, whereas collagen type I was not affected. The production of fibronectin and collagen type IV was also not affected by IGF-I (data not shown). In addition, in these cells, TGF-β1 could not induce collagen type III under conditions in which collagens type I and IV and fibronectin were increased. This suggests that the requirement of growth factors for the induction of collagen type III is different from other ECM components. Whether there is a relation between the increase in collagen type III in response to IGF-I and the absence of a stimulatory effect of TGF-β is at present unknown.

Recently, it was demonstrated that collagen type III mRNA expression was synergistically upregulated by CTGF and IGF-I in NRK fibroblasts (17). We now demonstrate that CTGF and IGF-I functionally cooperate in the secretion of both collagens type I and III in human renal fibroblasts, although the mechanism is at present unknown. One possible mechanism is via the induction of autocrine growth factors. However, we have found that neither CTGF nor IGF-I was increased in renal fibroblasts upon stimulation with CTGF and/or IGF-I. On the contrary, IGF-I stimulation resulted in a decreased secretion of CTGF, even in the presence of exogenous CTGF. Therefore, we conclude that it is unlikely that the induction of either growth factor is involved in the enhanced effects of CTGF and IGF-I. Nevertheless, at present, we cannot exclude the involvement of alternative growth factors. Another possible mechanism for the functional collaboration between CTGF and IGF-I might be via a direct binding between both growth factors. CTGF has also been designated as IGF-binding protein (IGFBP)-related protein 2 (38) or IGFBP-8 (39) because it has the capacity to bind IGF-I via its IGF-binding domain, albeit with relatively low affinity compared with classical IGFBPs (39,40). In the present study, we were able to demonstrate this interaction between IGF-I and CTGF by immunoprecipitation followed by Western blotting. Although the potential of IGF-I to bind to CTGF has been discovered for some years now, there is still little information about the biological consequence of this interaction. It is known that the bioavailability of IGF-I is modulated by classical IGFBPs, which can either potentiate or inhibit IGF-I (41,42). Therefore, it is possible that the underlying mechanism of the functional cooperation between the two growth factors might be through increased bioavailability of IGF-I when it is bound to CTGF. The modulating action of CTGF on growth factors is not limited to IGF-I because it has been demonstrated recently that it can also promote TGF-β1 signaling but inhibits the action of BMP4 (43). In addition, CTGF can inhibit vascular endothelial growth factor–induced angiogenesis (44). Taken together, CTGF might function as a cofactor for other growth factors, thereby modulating the biological activity of these growth factors. However, more study on the molecular and biological level is needed to unravel the precise pathways involved in the functional collaboration between CTGF and other growth factors.

Apart from the effect of hyperglycemic conditions on CTGF and IGF-I production and the cooperation between CTGF and IGF-I in collagen type I and III accumulation, we have also investigated the effect of these growth factors under hyperglycemic conditions. We have demonstrated that IGF-I and not CTGF was able to further enhance glucose-induced collagen secretion. These data suggest that induction of IGF-I, not CTGF, is the rate-limiting step in the glucose-induced matrix accumulation. Therefore, it will be important to delineate further the molecular mechanism of the CTGF-IGF-I interactions.

In conclusion, this study demonstrates that CTGF and IGF-I cooperate in their upregulation of collagen type I and III expression in human renal fibroblasts. The synergy between CTGF and IGF-I might be involved in glucose-induced matrix accumulation, because both factors are induced by hyperglycemia.

Part of this work was performed in the framework of the EU project “Chronic Kidney Disease” (EU-QLG1-CT2002-01215). This study was supported by the Dutch Diabetes Research Foundation (DFN 98.130).

We thank Lotte Wieten (Department of Pathology, University Medical Center Utrecht, Utrecht, the Netherlands) and Nike Claessen (Department of Pathology, Academic Medical Center, Amsterdam, the Netherlands) for technical assistance and Dr. Frank Strutz (Department of Nephrology and Rheumatology, Georg-August-University Göttingen, Göttingen, Germany) for providing the TK173 cell line.