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. 2012 Mar 15;26(5):833–845. doi: 10.1210/me.2011-1261

Low-Density Lipoprotein Induced Expression of Connective Tissue Growth Factor via Transactivation of Sphingosine 1-Phosphate Receptors in Mesangial Cells

Hesham M El-Shewy 1, Mimi Sohn 1, Parker Wilson 1, Mi Hye Lee 1, Samar M Hammad 1, Louis M Luttrell 1, Ayad A Jaffa 1,✉
PMCID: PMC3355554  PMID: 22422617

Abstract

The pro-fibrotic connective tissue growth factor (CTGF) has been linked to the development and progression of diabetic vascular and renal disease. We recently reported that low-density lipoproteins (LDL) induced expression of CTGF in aortic endothelial cells. However, the molecular mechanisms are not fully defined. Here, we have studied the mechanism by which LDL regulates CTGF expression in renal mesangial cells. In these cells, treatment with pertussis toxin abolished LDL-stimulated activation of ERK1/2 and c-Jun N-terminal kinase (JNK), indicating the involvement of heterotrimeric G proteins in LDL signaling. Treatment with LDL promoted activation and translocation of endogenous sphingosine kinase 1 (SK1) from the cytosol to the plasma membrane concomitant with production of sphingosine-1-phosphate (S1P). Pretreating cells with SK inhibitor, dimethylsphinogsine or down-regulation of SK1 and SK2 revealed that LDL-dependent activation of ERK1/2 and JNK is mediated by SK1. Using a green fluorescent protein-tagged S1P1 receptor as a biological sensor for the generation of physiologically relevant S1P levels, we found that LDL induced S1P receptor activation. Pretreating cells with S1P1/S1P3 receptor antagonist VPC23019 significantly inhibited activation of ERK1/2 and JNK by LDL, suggesting that LDL elicits G protein-dependent activation of ERK1/2 and JNK by stimulating SK1-dependent transactivation of S1P receptors. Furthermore, S1P stimulation induced expression of CTGF in a dose-dependent manner that was markedly inhibited by blocking the ERK1/2 and JNK signaling pathways. LDL-induced CTGF expression was pertussis toxin sensitive and inhibited by dimethylsphinogsine down-regulation of SK1 and VPC23019 treatment. Our data suggest that SK1-dependent S1P receptor transactivation is upstream of ERK1/2 and JNK and that all three steps are required for LDL-regulated expression of CTGF in mesangial cells.

Diabetic nephropathy is a major cause of morbidity and mortality in diabetes. It is the single most common cause of end-stage renal failure (1). A very characteristic early event of the development of diabetic nephropathy is glomerulosclerosis, which is characterized by thickening of the glomerular basement membrane and widening of the mesangium with accumulation of extracellular matrix (ECM). The degree of mesangial expansion is strongly related to the clinical manifestations of diabetic nephropathy, such as albuminuria and decreased glomerular filtration rate (2). Inflammatory mediators and growth factors are increasingly recognized as playing important roles in the development of glomerular injury and remodeling (3).

Connective tissue growth factor (CTGF) is a 36- to 38-kDa secreted protein belonging to the CNN family of matricellular proteins (4). It regulates ECM synthesis, including types of collagens, fibronectin, and laminin (5–7). CTGF expression is markedly elevated in pathological conditions characterized by ECM deposition and fibrosis (8). It is also up-regulated in the kidney of experimental animal models of proliferative glomerulonephritis and experimental diabetic glomerulosclerosis (9–11). Enhanced CTGF mRNA expression in response to TGF-β has been reported in cultured mesangial cells, podocytes, proximal tubular cells, and renal fibroblasts (12–15). These findings, together with recent human studies suggesting that elevated plasma and urinary CTGF is a marker for the progression of diabetic nephropathy (10, 16–18), strongly support a role for CTGF in the pathogenesis of renal fibrosis. In addition to TGF-β, several other factors have been shown to induce the expression of CTGF, including high glucose (10, 11, 18), advanced glycated end products (19), reactive oxygen species (20), angiotensin II (9), sphingosine-1-phosphate (S1P) (21, 22), and lysophosphatidic acid (13). Although CTGF and its role in development and progression of fibrotic diseases, particularly diabetic nephropathy, have been extensively studied, the molecular mechanisms by which these factors regulate CTGF expression are not fully elucidated. We and other independent investigators recently reported that activation of MAPK is an essential step for CTGF expression (23–25). We also demonstrated that low-density lipoprotein (LDL)-induced CTGF expression involves ERK1/2 and c-Jun N-terminal kinase (JNK), but not p38 MAPK in mesangial cells and human aortic endothelial cells (23, 25). We further demonstrated that LDL responses were mediated by binding to LDL receptors expressed on the surface of mesangial cells (26). However, the exact mechanism by which LDL induces MAPK-dependent CTGF expression has yet to be determined.

Sphingosine kinase 1 (SK1), a key enzyme catalyzing the formation of S1P, has been implicated in signaling by myriad effectors including growth factors, cytokines, and agonists of various G protein-coupled receptors. S1P is a bioactive sphingolipid that has attracted considerable attention during the last few years because of its ability to regulate diverse cellular responses, including cell growth, motility, proliferation, differentiation, migration, and apoptosis (27). S1P has been reported to function as both an intracellular second messenger and an extracellular autocrine factor the effects of which are mediated by binding to a family of five high-affinity G protein-coupled receptors, S1P1–5 (28). A growing body of evidence suggests the involvement of S1P in regulation of CTGF expression. cDNA microarray analysis revealed up-regulation of CTGF expression upon S1P stimulation of cultured rat mesangial cells (29). S1P-mediated CTGF expression is reported to be pertussis toxin (PTX) sensitive in mesangial calls (21), and Rho A-dependent in smooth muscle cells (30). Stimulation with S1P increases CTGF mRNA and protein expression in a concentration and time-dependent manner in human umbilical vein endothelial cells (22) and in a Wilm's tumor cell line (WiT49) (24), in which the response was blocked by inhibition of S1P2 receptor and JNK signaling. Moreover, stimulation with FTY720, a synthetic substrate for SK1 and phospho-FTY720, an S1P analog that binds S1P receptors, led to marked up-regulation of CTGF and collagen type IV protein expression in mesangial cells (31). Most recently, using live cell imaging and membrane biotinylation of endothelial cells, plasma membrane S1P1 receptor was shown to be rapidly internalized and translocated to the nuclear compartment upon S1P stimulation, whereupon it was able to regulate transcription of CTGF (32).

We undertook this study to delineate the mechanism underlying LDL-promoted expression of CTGF in rat mesangial cells. We found that LDL stimulated activation and translocation of SK1, which in turn transactivates S1P receptors. Pharmacological inhibition of SK1 or competitive antagonism of S1P1/S1P3 receptor signaling inhibited LDL-dependent activation of ERK1/2 and JNK. S1P stimulated CTGF expression in a dose-dependent manner that was attenuated by ERK1/2 and JNK inhibitors. Moreover, LDL-promoted CTGF mRNA and protein expression was dependent upon SK activity, S1P1/S1P3 receptors, and PTX-sensitive G proteins. These data suggest that SK1-dependent S1P receptor transactivation mediates LDL-induced CTGF expression in mesangial cells.

Results

LDL activates ERK1/2 and JNK through a heterotrimeric G protein-dependent pathway

Several studies have implicated heterotrimeric G proteins in LDL signaling (33, 34). To test whether G proteins were involved in the LDL-dependent activation of MAPK in rat mesangial cells (RMC), we employed PTX to inactivate Gi/o family heterotrimeric G proteins before short-term stimulation with either LDL or S1P. As shown in Fig. 1, both LDL and S1P stimulation markedly increased phosphorylation of ERK1/2 and JNK (P < 0.001), and the response was abolished by PTX treatment.

Fig. 1.

Fig. 1.

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LDL activates ERK1/2 and JNK through a PTX-sensitive pathway. Quiescent RMC were serum deprived overnight in the presence or absence of 500 ng/ml PTX before stimulation for 5 min with 50 μg/ml LDL or 5 μm S1P. Basal (NS), LDL-, and S1P-stimulated phosphorylation of ERK1/2 and JNK in whole-cell lysate samples was determined as described. Representative phospho-ERK1/2 (A) and phospho-JNK (B) immunoblots are shown above bar graphs depicting mean ± sd for four independent experiments. The change in phosphorylation ERK1/2 and JNK are expressed as the percent above the basal level in unstimulated cells not exposed to PTX. *, P < 0.001 vs. unstimulated (NS); and #, P < 0.001 stimulated vs. PTX treated. NS, Not stimulated.

SK1 mediates LDL-dependent activation of ERK1/2 and JNK

The finding that PTX inhibited LDL-stimulated activation of MAPK suggested that Gi/o protein activation is required for the response. One mechanism that could account for this would be LDL-induced transactivation of an endogenous Gi/o-coupled G protein-coupled receptor (GPCR). We considered the family of S1P receptors as likely candidates for LDL-regulated GPCR. S1P receptors are ubiquitously expressed and are known to be involved in diverse cellular responses including cell growth, survival, and migration (35, 36). The ligand for S1P receptors, S1P, is a bioactive sphinoglipid metabolite formed through the action of two known isoforms of sphingosine kinase, SK1 and SK2 (37). To test whether the SK/S1P pathway plays a role in LDL signaling, we initially employed the pharmacological inhibitor N,N-dimethyl-sphingosine (DMS) that efficiently inhibit both isoforms of SK. As shown in Fig. 2, stimulation of RMC with LDL for 5 min induced a significant increase in ERK1/2 and JNK phosphorylation (6.6 ± 0.9 and 7.8 ± 0.2 fold over basal, respectively; P < 0.001). Both responses were significantly inhibited by preincubation with DMS (4.3 ± 0.8- and 3.5 ± 0.21-fold over basal, respectively; P < 0.05). To further confirm this finding and to determine which isoform is involved in this pathway we employed RNA interference specifically targeting hSK1 and hSK2 using human mesangial cells (HMC). Using 100 nm concentrations of small interfering RNA (siRNA) efficiently knock down endogenous SK1 and SK2 RNA levels by 82% and 84%, respectively; and protein levels by 52% and 63%, respectively (Fig. 3, A and B). Figure 3, C and D, depicts that down-regulation of SK1 significantly inhibited LDL-dependent activation of ERK1/2 and cJNK (∼63% and 50%, respectively; P < 0.01). Interestingly, down-regulation of SK2 showed no significant changes in activation of ERK1/2 and JNK, suggesting the isoenzyme specificity of SK1 involvement in this signaling pathway.

Fig. 2.

Fig. 2.

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Inhibition of SK1 inhibits activation of ERK1/2 and JNK by LDL. RMC were serum deprived overnight and incubated in the presence or absence of 20 μm DMS for 60 min before stimulation for 5 min with 50 μg/ml LDL. Basal (NS) and LDL-stimulated phosphorylation of ERK1/2 and JNK in whole cell lysate samples was determined as described. Representative phospho-ERK1/2 (A) and phospho-JNK (B) immunoblots are shown above bar graphs depicting mean ± sd for at least three independent experiments. The change in phosphorylation of ERK1/2 and JNK is expressed as the percent above the basal level in unstimulated cells not exposed to DMS. *, P < 0.001 vs. unstimulated (NS); and #, P < 0.05 stimulated vs. DMS treated. NS, Not stimulated.

Fig. 3.

Fig. 3.

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Down-regulation of SK1 by RNA interference inhibited LDL-stimulated activation of ERK1/2 and JNK. A, HMC were transfected with control scrambled siRNA (SCR) or siRNA targeting SK1 (siSK1) or SK2 (siSK2) for 48 h. RNA was isolated, and mRNA levels of SK1, SK2, and GAPDH were determined by quantitative real-time PCR. B, The levels of SK1, SK2, and GAPDH were determined by immunoblotting whole-cell lysates. A representative SK1, SK2, and basal GAPDH immunoblot are shown above a bar graph depicting mean ± sd for three independent experiments. C and D, Serum-deprived cells transfected with scrambled siRNA or siRNA targeting SK1 and SK2 (100 nm) were stimulated with 50 μg/ml LDL or 5 μm S1P for 5 min, after which whole-cell lysates were assayed for phosphorylation of ERK1/2 and JNK as described. Data shown represent the mean ± sd of three independent experiments. *, P < 0.01 vs. scrambled (SCR) treated. NS, Not stimulated.

LDL stimulation induces activation and membrane translocation of SK1

Previous work has demonstrated that external stimuli, such as phorbol ester-dependent protein kinase C and IGF receptor activation, can promote membrane translocation and activation of SK1 (38, 39), suggesting that regulated production of S1P might provide a mechanism for transactivating S1P receptors. To test whether LDL stimulation activated SK1, we first employed a membrane fractionation assay measuring the quantity of endogenous SK1 and phospho-SK1 present in the membrane fraction. Exposure of RMC to LDL for 15 or 30 min caused a significant increase in SK1 and phospho-SK1 in the membrane fraction compared with nonstimulated cells (P < 0.05). No significant change was detected in the cytosolic fraction, suggesting that despite a marked increase in plasma membrane-associated SK1, the amount translocated represents only a small fraction of the total cellular SK1 pool (Fig. 4A). Furthermore, time course stimulation of RMC with LDL induced phosphorylation of SK1 within 2 min, an effect that persisted for up to 30 min (Fig. 4B). To further confirm SK1 activation by LDL stimulation, we employed in situ C17-sphingosine labeling. C17-sphingosine lacks one carbon in their nonpolar moiety compared with the natural C18-sphingosine and has been used recently as an indicator for SK1 activation for mass spectrometry measurement of S1P production (40). As shown in Fig. 4C, stimulation of RMC with LDL or phorbol 12-myristate 13-acetate (PMA) for 15 min induced significant increases in C17-S1P production in cell lysates (picomoles/nmol phosphate) (3.414 ± 0.527, P > 0.02 and 3.642 ± 0.567, P > 0.012; respectively) compared with unstimulated cells (2.276 ± 0.173), and significant increases C17-S1P in the media (picomoles/ml) (8.813 ± 0.974, P > 0.0001 and 2.739 ± 0.109, P > 0.0001; respectively) compared with unstimulated cells (345 ± 0.208), indicating SK1 activation by LDL stimulation.

Fig. 4.

Fig. 4.

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LDL promotes activation and membrane translocation of endogenous SK1 in mesangial cells. A, Serum starved RMC were stimulated with 50 μg/ml LDL for 15 and 30 min. Fractionation of cytosolic and membrane fractions of basal (NS) and LDL-treated cells and detection of endogenous SK1 and phospho-SK1 protein were performed as described. SK1 immunoblots are shown above a bar graph depicting mean ± sd for three independent experiments. The changes in SK1 and phospho-SK1 protein levels are expressed as the percent above the basal level in unstimulated cells. *, P < 0.05 vs. unstimulated (NS). B, Serum-deprived RMC were stimulated with 50 μg/ml LDL for indicated times, and activation of SK1 in whole-cell lysate samples was determined by immunoblotting with phosphorylation state-specific IgG. SK1 phosphorylation is expressed as fold increase above basal levels in unstimulated cells. A representative phospho-SK1 and basal GAPDH immunoblots are shown above a bar graph presenting mean ± sd of three independent experiments. *, P < 0.05 vs. unstimulated (NS). C, Serum starved RMC were stimulated with 50 μg/ml LDL or 200 nm PMA for 15 min after which C17–S1P in media and whole-cell lysates were assayed as described. Data shown present the mean ± sd of three independent experiments. *, P < 0.05 vs. unstimulated (NS). NS, Not stimulated.

LDL induces activation and internalization of green fluorescent protein (GFP)-S1P1 receptors

SK1 catalyzes the phosphorylation of sphingosine to generate S1P. Because our data indicated that LDL-stimulated MAPK activation in mesangial cells was heterotrimeric G protein dependent and required SK1 activity, we tested the hypothesis that the S1P generated in response to LDL was eliciting G protein-dependent signals by transactivating endogenous S1P receptors. To determine whether LDL stimulation generated sufficient S1P to activate S1P receptors, we transfected RMC with a GFP-tagged S1P1 receptor (36) and employed it as a biological sensor for physiologically relevant extracellular quantities of S1P. A characteristic feature of most GPCR is ligand-dependent internalization (41). As shown in Fig. 5, GFP-S1P1 receptors expressed in RMC underwent a striking rearrangement from a uniform distribution on the plasma membrane to a punctate distribution within cytoplasm upon exposure to S1P, consistent with activation-dependent receptor internalization. Treatment with LDL produced a similar pattern, suggesting that LDL stimulation activates S1P receptors.

Fig. 5.

Fig. 5.

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LDL stimulates internalization of GFP-S1P1 receptors. Serum-deprived RMC transfected with GFP-tagged S1P1 receptors were stimulated with vehicle (NS), 50 μg/ml LDL, or 5 μm S1P for 10 min. After stimulation, cells were fixed and the distribution of S1P1R-GFP was determined by confocal fluorescence microscopy. Shown are representative confocal fields from one of three independent experiments that gave similar results. NS, Not stimulated.

Antagonizing S1P1/S1P3 receptors inhibits LDL-dependent activation of ERK1/2 and JNK

To further test our hypothesis that S1P generated in response to LDL stimulation acts via endogenous S1P receptors, we tested whether S1P receptor antagonism would affect LDL signaling. Nonselective S1P receptor antagonists that potently inhibit all five S1P receptor subtypes are not currently available. Because the S1P1 receptor has been implicated in S1P-regulated transcription of CTGF in endothelial cells (32), we employed VPC23019, a selective S1P1/S1P3 receptor antagonist (38), using S1P stimulated ERK1/2 phosphorylation in RMC as the control for the efficiency of endogenous S1P receptor blockade. Cells were pretreated with varying concentrations of the antagonist for 1 h before stimulation with S1P for 10 min. As shown in Fig. 6A, 50 μm VPC23019 reduced S1P-stimulated ERK1/2 activation by approximately 50%. We next determined the effect of VPC23019 on LDL-dependent MAPK activation. As shown in Fig. 6, B and C, stimulation with LDL for 5 min produced a significant increase in ERK1/2 and JNK phosphorylation (9.5 ± 0.3- and 9.7 ± 1.3-fold over basal, respectively), and the response was attenuated in the presence of the VPC23019 (5.9 ± 0.6- and 6.6 ± 0.9-fold over basal for ERK1/2 and JNK phosphorylation, respectively; P < 0.05). This was similar to the extent of inhibition observed after direct S1P stimulation (Fig. 6A), suggesting that endogenous S1P1/S1P3 receptors contribute to LDL-dependent activation of ERK1/2 and JNK in mesangial cells.

Fig. 6.

Fig. 6.

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The competitive S1P1/S1P3 receptor antagonist VPC23019 attenuates LDL-stimulated activation of ERK1/2 and JNK. A, Serum deprived RMC were pretreated with 50 μm VPC23019 (VPC) for 30 min before stimulation for 10 min 5 μm S1P. Basal (NS) and S1P-stimulated ERK1/2 phosphorylation in whole-cell lysates was determined as described. B and C, Serum-deprived RMC were pretreated with 50 μm VPC23019 (VPC) for 30 min before stimulation for 10 min with 50 μg/ml LDL. Basal (NS) and LDL-stimulated ERK1/2 and JNK phosphorylation in whole-cell lysate samples was determined as described. Representative phospho-ERK1/2 and JNK immunoblots are shown above bar graphs depicting mean ± sd for three independent experiments. The change in ERK1/2 phosphorylation is expressed as the percent increase above the basal level in unstimulated cells not exposed to VPC23019. *, P < 0.05 vs. unstimulated (NS); and #, P < 0.05 stimulated vs. VPC treated. NS, Not stimulated.

S1P-stimulated CTGF expression is sensitive to inhibition of ERK1/2 and JNK

Previous reports have suggested that S1P regulates expression of CTGF via a PTX-sensitive pathway (22, 23, 29). Because our data suggested that LDL stimulates MAPK activation via SK-dependent activation of S1P receptors, we next tested whether S1P stimulation is sufficient to promote CTGF expression in mesangial cells and whether this response is dependent on ERK1/2 and JNK activation. As shown in Fig. 7A, stimulation of RMC with S1P for 24 h resulted in a concentration-dependent increase in CTGF protein expression with the maximal response (3.5-fold) occurring at 10 μm S1P (P < 0.001). To test the involvement of ERK1/2 and JNK in the response, we measured S1P-stimulated CTGF expression in the presence and absence of MEK1 (PD98059) and JNK (SP600125) inhibitors. As shown in Fig. 7, B and C, the increase in CTGF expression induced by S1P was markedly suppressed in the presence of either inhibitor (1.5 ± 0.1- vs. 1.1 ± 0.1-fold over basal for PD98059, P < 0.05; 2.4 ± 0.5 vs. 0.8 ± 0.2 for SP600125, P < 0.05). These findings suggest that the S1P receptor-dependent MAPK activation is required for S1P-induced production of CTGF.

Fig. 7.

Fig. 7.

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ERK1/2 or JNK mediates S1P-induced increase of mesangial cell CTGF expression. A, Quiescent RMC were serum deprived overnight before stimulation with the indicated concentrations of S1P for 24 h. Basal (NS) and S1P-stimulated CTGF protein expression in whole-cell lysate samples was determined as described. A representative CTGF immunoblot is shown above a bar graph depicting mean ± sd for four independent experiments. The change in CTGF is expressed as the percent above the basal level in unstimulated cells. *, P < 0.05; and #, P < 0.005 vs. unstimulated (NS). B, Serum-deprived mesangial cells were pretreated with 40 μm PD98059 (panel B) or 30 μm SP600125 (panel C) for 60 min before stimulation with 5 μm S1P for 24 h. Basal (NS) and S1P-stimulated CTGF protein expression in whole-cell lysate samples was determined as described. Representative CTGF immunoblots are shown above bar graphs depicting mean ± sd for three independent experiments. The change in CTGF protein is expressed as the percent above the basal level in unstimulated cells. *, P < 0.05 vs. unstimulated (NS). NS, Not stimulated.

LDL-dependent up-regulation of CTGF is sensitive to PTX

We next tested whether the SK1/S1P pathway is involved in LDL-induced expression of CTGF in mesangial cells. As shown in Fig. 8, stimulation of RMC with LDL or S1P for 24 h resulted in significant elevation of CTGF protein expression (1.5 ± 0.2- and 2.0 ± 0.4-fold over basal, respectively; P < 0.05). The response was inhibited by PTX (0.9 ± 0.1- and 1.3 ± 0.2-fold over basal, respectively; P < 0.03) suggesting that Gi/o protein activation is required for LDL-stimulated production of CTGF in these cells.

Fig. 8.

Fig. 8.

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PTX inhibits LDL- and S1P-induced CTGF expression. RMC were serum deprived overnight in the presence or absence of 500 ng/ml PTX before stimulation with 50 μg/ml LDL or 5 μm S1P for 24 h. Basal (NS) and S1P-stimulated CTGF protein expression in whole-cell lysate samples was determined as described. Representative CTGF immunoblots are shown above a bar graph depicting mean ± sd for three independent experiments. The change in CTGF protein is expressed as the percent above the basal level in unstimulated cells. *, P < 0.05 vs. unstimulated (NS); and #, P < 0.05 stimulated vs. PTX treated. NS, Not stimulated.

LDL-induced up-regulation of CTGF mRNA requires both SK1 activity and S1P receptors

To further assess the involvement of SK1 and S1P receptors in LDL-induced expression of CTGF, we determined the effect of SK1 inhibition and S1P1/S1P3 receptor antagonism on CTGF mRNA transcription in RMC. As shown in Fig. 9, LDL stimulation for 4 h significantly increased CTGF mRNA abundance (7.5 ± 0.7-fold over basal; P < 0.001). The transcriptional response was markedly reduced when cells were pretreated with DMS (1.8 ± 0.1-fold over basal; P < 0.001) and VPC23019 (2.1 ± 0.4-fold over basal; P < 0.002) suggesting that SK1-dependent transactivation of endogenous S1P receptors is required for LDL-induced CTGF mRNA transcription. A similar increase in CTGF mRNA abundance was observed with direct S1P stimulation (6.6 ± 0.4-fold over basal; P < 0.001), confirming that S1P receptor activation is sufficient to increase CTGF mRNA transcription and translation in mesangial cells. Predictably, the response to exogenously supplied S1P, which would be expected to bypass the requirement of SK activity, was insensitive to DMS (7.1 ± 0.2-fold over basal), but inhibited by S1P1/S1P3 receptor antagonist, VPC23019 (2.5 ± 0.3-fold over basal; P < 0.002). Interestingly, VPC23019 was more effective at inhibiting LDL- and S1P-stimulated CTGF mRNA transcription than acute ERK1/2 activation (Fig. 6), suggesting that although other S1P1–5 receptors can contribute to LDL- and S1P-stimulated MAPK signaling, S1P1/S1P3 receptors are the principal regulators of mesangial cell CTGF expression. To further confirm that SK1 is required for LDL-regulated expression of CTGF, we used RNA interference to down-regulate hSK1 in HMC. As shown in Fig. 10, LDL stimulation for 4 h significantly increased CTGF mRNA abundance (4.4 ± 0.7-fold over basal; P < 0.001) and protein levels (2.1 ± 0.5-fold over basal; P < 0.001). Similar to the previous finding using DMS, efficient knock down of SK1 (data not shown) abolished LDL-stimulated CTGF mRNA abundance (0.966 ± 0.02-fold over basal; P < 0.001 vs. SCR treated) and protein expression (1.13 ± 0.12-fold over basal; P < 0.02 vs. SCR treated) (Fig. 10, A and B) indicting that LDL-induced CTGF expression in renal mesangial cells is mediated by SK1.

Fig. 9.

Fig. 9.

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Inhibition of SK or S1P1/S1P3 receptor signaling inhibits LDL-stimulated CTGF mRNA transcription. RMC were serum deprived overnight in the presence or absence of 20 μm DMS for 60 min before stimulation for 4 h with 50 μg/ml LDL. RNA was isolated and mRNA levels of CTGF and β-actin were determined by quantitative real-time PCR as described. Data are presented as mean ± sd for at least three independent experiments. Change in CTGF mRNA abundance is expressed as fold increase above the basal level in unstimulated cells. *, P < 0.001 vs. unstimulated (NS); and #, P < 0.001 stimulated vs. inhibitor. NS, Not stimulated.

Fig. 10.

Fig. 10.

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Down-regulation of SK1 inhibited LDL-stimulated CTGF protein expression. Serum-deprived HMC transfected with (100 nm) of scrambled siRNA (SCR) or siRNA targeting SK1 (siSK1) for 48 h were stimulated with 50 μg/ml LDL or 5 μm S1P for 4 h. RNA was isolated, and mRNA levels of CTGF and GAPDH were determined by quantitative real-time PCR (A), and CTGF protein expression in whole-cell lysate samples (B) were determined as described. Data shown represent the mean ± sd of three independent experiments. Change in CTGF mRNA abundance and CTGF protein is expressed as fold increase and the percent above the basal level in unstimulated cells. Representative CTGF and GAPDH immunoblots are shown above a bar graph depicting mean ± sd for three independent experiments. The change in. * P < 0.01 and 0.2 vs. SCR treated. NS, Not stimulated.

Discussion

Although the association of chronic hyperglycemia and dyslipidemia with diabetic microvascular disease is well recognized, the factors and cellular signaling mechanisms that link them with renal injury and remodeling are not fully defined. Metabolic imbalances associated with high tissue glucose and abnormal lipid levels in the diabetic state may directly influence signal transduction pathways that contribute to the pathogenesis of diabetic vascular and renal injury. Abnormalities in lipid and lipoprotein metabolism have been linked to a myriad of diabetic complications including micro- and macrovascular disease, altered gene expression in arterial endothelial and smooth muscle cells, in which cell proliferation and apoptosis coexist, stimulation of monocyte infiltration into the mesangial space, mesangial cell hypercellularity, and deposition of ECM (42–45). In this regard, we have previously shown that LDL, via binding to LDL receptor, activate ERK1/2 in mesangial cells through a mechanism involving increased intracellular calcium, Src family kinases, and protein kinase C (26, 46). Because of their central role in the regulation of cell proliferation, inflammation and stress responses (47–49), the MAPK are likely to be important regulators of the expression of factors, like CTGF, that mediate tissues responses to LDL.

It is recognized that renal injury develops through processes involving all cell types in the kidney, including synthesis of extracellular proteins; e.g. cytokines, growth factors and matrix proteins, and activation of both proliferative and antiproliferative pathways. There is growing appreciation of the role of sphingolipids in regulating these events. Sphingolipid metabolites are carried in circulating lipoprotein particles with more than 60% of the S1P in blood associated with very low density lipoprotein, LDL, and high-density lipoprotein (HDL), consistent with the hypothesis that extracellular delivery of S1P may be involved in the generation of intracellular signals that promote proliferation of vascular cells, favoring thickening and plaque stabilization (50–52).

Recent studies, however, have shown that HDL is the major carrier of plasma S1P (51, 53, 54). HDL3 and HDL2 carry 78.6% and 16.3% of lipoprotein-associated plasma S1P, respectively (54). In contrast, LDL-associated S1P comprises only 3.73% of plasma S1P (54), an amount equivalent to 0.65 pmol S1P per 50 μg LDL, the amount used in our assays. Thus, the quantity of S1P we added exogenously in the form of LDL was likely to be insufficient to produce direct activation of S1P receptors. Moreover, our data clearly demonstrate that LDL induces immediate activation and membrane translocation of SK1, increasing S1P production. Further, the S1P receptor-dependent responses we observe in response to LDL, including activation of the ERK1/2 and JNK cascades and up-regulation of CTGF expression, are blocked by inhibition and down-regulation of SK1, which would not occur if LDL-associated S1P was providing the stimulus. Collectively, these data strongly implicate S1P generated within the cell by SK1 in the LDL-mediated transactivation of S1P receptors.

Stimuli present in the diabetic milieu have been shown to regulate SK1 expression or activity, causing vascular or renal cells to generate an endogenous pool of S1P. SK1 expression is induced by the inflammatory cytokine TGF-β and is involved in the TGF-β signaling pathway leading to up-regulation of tissue inhibitor metalloproteinase-1 (55). Conversely, S1P transactivates the TGF-β receptor and triggers activation of Sma- and Mad-related protein signaling (56). Endogenously produced S1P acts both intracellularly as a second messenger and extracellularly as a ligand for the S1P1–5 family of G protein-coupled receptors, allowing it to activate processes downstream of PTX-sensitive G proteins (57). SK1 activation in response to extracellular stimuli leads to S1P receptor-dependent activation of MAPK, calcium mobilization, and mesangial cell proliferation (58, 59). Consistent with these findings, our data indicate that activation of endogenous SK1 contributes to increased CTGF expression in mesangial cells exposed to LDL.

Figure 11 depicts our proposed model for LDL-regulated expression of CTGF in mesangial cells. Exposure to LDL promotes activation and membrane translocation of SK1, generating sufficient extracellular S1P to activate endogenous G protein-coupled S1P receptors. Once activated, S1P receptors activate ERK1/2 and JNK via a PTX-sensitive G protein pathway, which in turn regulates CTGF expression. Using DMS and RNA interference to block SK1, VPC23019 to antagonize S1P1/S1P3 receptors, PTX to inhibit Gi/o proteins, and PD98059 and SP600125 to block ERK1/2 and JNK signaling, our data implicate all three steps in the process of LDL-regulated CTGF expression in mesangial cells. Although these data do not exclude the involvement of other S1P receptors in LDL-stimulated MAPK activation, they strongly suggest that transactivation of endogenous S1P receptors contributes to LDL-dependent MAPK activation in mesangial cells.

Fig. 11.

Fig. 11.

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Model depicting the mechanism of LDL-dependent up-regulation of CTGF by SK1-dependent transactivation of S1P receptors. LDL stimulation promotes activation and membrane translocation of SK1, leading to transactivation of endogenous S1P receptors that, in turn, induces G protein-dependent activation of the ERK1/2 and JNK pathways required for CTGF expression.

Recent studies have focused on the role of CTGF, an emerging determinant of progressive fibrotic diseases, in the pathogenesis of vascular and renal injury. Indirect evidence supporting this notion is provided by the fact that high levels of expression of CTGF mRNA and protein occurs in vascular smooth muscle cells and endothelial cells of advanced human atherosclerotic lesions, but not in normal patients (60). In addition, vascular smooth muscle cells expressing CTGF localize predominantly in areas with ECM production and especially in areas around the fibrous cap, thus indicating that CTGF may regulate the production of matrix proteins in these cells (60). We have recently reported that type 1 diabetic patients with macroalbuminuria have increased circulating and urinary levels of CTGF, and demonstrated a positive and significant association between plasma CTGF levels and lipoprotein levels, suggesting that lipoproteins may modulate CTGF expression in diabetic patients (17). Consistent with this, we found that LDL induces CTGF expression in renal mesangial and human aortic endothelial cells via activation of the ERK1/2 and JNK MAPK pathways (1, 25). Our present findings extend these observations, demonstrating that LDL induces the activation of SK1 and that both LDL and S1P stimulate CTGF expression in mesangial cells. Furthermore, our data demonstrate that inhibition of SK1 and or S1P receptors attenuates the effect of LDL on CTGF expression in mesangial cells. These findings point to a new mechanistic pathway through which lipoproteins stimulate CTGF to promote renal and vascular dysfunction in diabetes and identify novel targets for interventional strategies.

Materials and Methods

Materials

Tissue culture medium, fetal bovine serum, and penicillin/streptomycin were from Invitrogen (Carlsbad, CA). FuGENE 6 was from Roche Diagnostics (Indianapolis, IN). Double-stranded siRNA were purchased from Xeragon (Germantown, MD). GeneSilencer transfection reagent was from Gene Therapy Systems (San Diego, CA). Primers for real-time PCR were from Integrated DNA Technologies (Coralville, IA). RNeasy kits were from QIAGEN (Valencia, CA) and iScript cDNA synthesis kits and iQ SYBR Green Supermix kits were from Bio-Rad Laboratories, Inc. (Hercules, CA). PTX was from List Biological Laboratories (Campbell, CA). PMA, PD98059, and SP600125 were from Sigma Chemical Co. (St. Louis, MO). VPC23019 was from Avanti Polar Lipids (Alabaster, AL). S1P and DMS were provided by the Medical University of South Carolina Lipidomics Core (Department of Biochemistry, Medical University of South Carolina, Charleston, SC). Rabbit polyclonal anti-ERK1/2, JNK, phospho-ERK1/2, and phospho-JNK IgG were from Cell Signaling Technology (Beverly, MA). Rabbit polyclonal anti-CTGF IgG was from Abcam (Cambridge, MA). Rabbit polyclonal anti-SK1 and anti phosphor-SK1 (Ser225) IgG were from Exalpha Biologicals (Watertown, MA) and ECM Bioscience (Versailles, KY), respectively. Horseradish peroxidase-conjugated donkey antirabbit IgG was from Amersham Biosciences (Pittsburgh, PA).

cDNA constructs

The pEGFP/myc S1P1R construct was a generous gift from Dr. Timothy Palmer (Institute of Biomedical and Life Sciences, University of Glasgow, Glasgow, Scotland, UK) and was prepared as previously described (36).

Mesangial cell culture

Rat glomerular mesangial cells (RMC) were prepared by a modification of the method of Lovett et al. (61). Isolated glomeruli were incubated in PBS plus 0.1% gentamicin and 1% antibiotic/antimycotic, pH 7.4, containing collagenase (5 mg/ml), at 37 C for 30 min to remove epithelial cells, leaving the glomerular cores containing mesangial and endothelial cells. Glomeruli were vortexed every 10 min during the incubation. The cores were diluted in 1.5 ml RPMI 1640 medium with HEPES and l-glutamine (Invitrogen) per kidney, containing 0.1% gentamicin, 1% antibiotic /antimycotic, 0.5% insulin-transferrin solution, and 20% FBS, conditions that favor growth of mesangial cells. Cells were incubated at 37 C in a humidified atmosphere of 95% air-5%CO2. RMC isolated by this procedure were homogenous and used in all studies between passage 3 and 8. Human mesangial cells were maintained in a 1:1 mixture of DMEM and Ham's F12 supplemented with 17% fetal bovine serum, 26 μg/ml insulin, and penicillin/streptomycin.

LDL preparation and characterization

Human LDL was prepared as previously described (1). Briefly, blood was taken from fasting healthy nondiabetic volunteers into a lipoprotein preservative/antioxidant cocktail (LPPC) containing EDTA (0.1% wt/vol), chloramphenicol (20 μg/ml), gentamicin sulfate (50 μg/ml), and ϵ-amino-n-caproic acid (0.13% wt/vol). Phenylmethylsulfonylfluoride (20 μg/ml) was added to plasma to retard proteolysis. All samples were processed at low temperature and in the absence of white light to minimize oxidation. Density gradient solutions were supplemented with LPPC, degassed, and purged with N2. Plasma density was increased to d = 1.21 g/ml using dried KBr, and 11 ml were layered under d = 1.019 g/ml saline/LPPC. Ultracentrifugation was performed at 7 C in a Beckman (Brea, CA) VTi50 rotor for 2.5 h at 50,000 rpm, with slow acceleration and deceleration, after which the LDL band was harvested by piercing the tube and syringe aspiration. LDL isolated by this procedure was free from contamination with apolipoprotein A-I and albumin. Each LDL preparation was tested for purity by electrophoresis on 1% agarose gels (Paragon Gels, Beckman). LDL pools were tested for endotoxin contamination by the Limulus Amebocyte Lysate (Bioproducts; Walkersville, MD) according to the manufacturer's suggestion.

Cell fractionation

Cell fractionation was performed as previously described (39). Briefly, rat mesangial cells were cultured to 80% confluence in 100-mm dishes and then serum starved overnight. Cells were stimulated with LDL (50 μg/ml) for 15 or 30 min, and then washed twice with ice-cold PBS and scraped in 500 μl of lysis buffer [20 mm Tris-HCl (pH 7.5), 10 mm EDTA, 2 mm EGTA, 250 mm sucrose, 1 mm phenylmethylsulfonyl fluoride, 10 nm okadaic acid, 1 mm dithiothreitol, 5 mm NaF, 1 mm Na3VO4, 0.5 mm 4-deoxypyridoxine, and 10 μg/ml leupeptin/aprotinin/soybean trypsin inhibitor]. Cell suspensions were sonicated, followed by centrifugation at 1000 × g for 10 min at 4 C. Supernatants were separated into membrane and cytosolic fractions were separated by centrifugation at 100,000 × g for 60 min at 4 C. The membrane fractions were resuspended in 350 μl of lysis buffer containing 0.8% Triton X-100, sonicated, and left on ice for 45 min. Triton-insoluble membrane fractions were collected by centrifugation at 10,000 × g for 10 min at 4 C, and the pellet was resuspended in 60 μl of lysis buffer. The protein concentration of each fraction was measured using the BCA Protein Assay Kit (Pierce Chemical Co., Rockford, IL) with BSA as a standard protein.

siRNA down-regulation of expression of SK1 and SK2

SK1 expression was down-regulated using human SK1 sequence-specific siRNA (5′-GGCCCAGCUGCCUAUGUAATT-3′ and 5′-UUACAUAGGC AGCUGGCCCA-3′). SK2 expression was down-regulated using human SK1 sequence-specific siRNA (5′-GAGGGUAGUGCCUGAUCAATT-3′and 5′-UUGAUCAGGCACUACCCUCGG-3′). Scrambled siRNA sequences (5′-ACGUGACACGUUCGGAGAAAdTdT-3′ and 5′-UUCUCCGAACGUGUCACGU dTdT-3′) were used as negative controls. Human mesangial cells were seeded in 10-cm dishes 24 h before transfection. Cells were transfected using Gene Silencer siRNA Transfection Reagent according to the manufacturer's protocol. The efficiency of the knockdown was determined by quantitative real-time PCR for hSK1 and hSK2 mRNA and immunoblotting for their proteins 48 h after transfection.

Protein immunoblotting

Mesangial cells were grown in 12-well multiwell plates, serum deprived overnight, and preincubated in the presence or absence of inhibitors as described in the figure legends. Cells were stimulated with LDL (50 μg/ml) or S1P (5 μm) for indicated time, after which monolayers were washed once in 4 C PBS and lysed in 200 μl of Laemmli sample buffer. For the determination of CTGF levels and phosphorylation of ERK1/2 and JNK, samples containing 20 μg of cell protein were resolved by SDS-PAGE and transferred to nitrocellulose filters. Phospho-ERK1/2, phospho-JNK, and CTGF were detected by protein immunoblotting using rabbit polyclonal IgG, with horseradish peroxidase-conjugated polyclonal donkey antirabbit IgG as secondary antibody. Immune complexes were visualized by enzyme-linked chemiluminescence and quantified using a Fluor-S MultiImager. In each experiment, equal loading of protein was confirmed by probing parallel immunoblots using antisera of basal proteins. Data obtained were normalized to actin protein level in the same sample and presented as percent of basal (nonstimulated).

Quantitative real-time PCR

Total cellular RNA was isolated using the RNeasy kit according to manufacturer's instructions. cDNA was prepared from 1 μg of total RNA with A260/A280 more than 1.8 using the iScript cDNA synthesis kit per manufacturer's instructions. Quantitative real-time PCR was performed with an iCycler 1Q Real-Time Detection System using the iQ SYBR Green Supermix kit. Reactions were performed using hCTGF-specific primers: forward primer, 5′-ACTATGATTAGAGCCAACTG-3′; reverse primer, 5′-TGTTCTCTTCCAGGTCAG-3′; β-actin-specific primers: forward primer, 5′-ATT GGCAATGAGCGGTTCC-3′; reverse primer, 5′-GGTAGTTTCGTCGATGCCACA-3′; hSK1: 5′-ACTATGATTAGAGCCAACTG 3′; reverse primer, 5′-TGTTCTCTTCCAGGTCAG-3′; hSK2: 5′-GGAGGAAGCTGTGAAGATGC-3′; reverse primer, 5′-GCAACAGTGAGCAGTTGAGC-3′; and glyceraldehyde-3-phosphate dehydrogenase (GAPDH): forward primer, 5′-CTGAGTACGTCGTGGAGTC-3′; reverse primer, 5′-AATGAGCCCCAGCCTTC-3′. Real-time PCR results were analyzed using Softmax Pro software (Molecular Devices, Sunnyvale, CA). Protein expression data were normalized to expression of β-actin or GAPDH as endogenous controls.

C17-Sphingosine labeling

Cells were labeled with C17-sphingosine (1 μm, 13 min, Avanti), stimulated as indicated, after which 2 ml of media were collected and cells were washed three times with PBS and collected. The reaction was stopped by the addition of 1 ml of extraction solvent containing ethyl acetate/2-propanol/water (60/30/10, vol/vol) supplemented with internal standard for enzyme immunoassay/liquid chromatography/mass spectrometry analysis. Lipids are extracted twice, dried under a stream of nitrogen, and resuspended into 150 μl of 1 mm NH4COOH in 0.2% HCOOH in methanol and analyzed by enzyme immunoassay/liquid chromatography/mass spectrometry.

Mass spectrometry

Sphingolipid fractions were analyzed using a Thermo Finnigan TSQ7000 triple quadrupole mass spectrometer, operating in a multiple reaction monitoring positive ionization mode as previously described (62). Sphingolipid levels were calculated and normalized to lipid phosphate (Pi).

Confocal fluorescence microscopy

For visualization of GFP-SK1 and GFP-S1P1 receptor, 50–60% confluent cultures of mesangial cells in 10-cm dishes were transiently transfected with either GFP-tagged hSK1 or cotransfected with untagged hSK1 and GFP-S1P1 receptor. Untagged SK1 was included in the GFP-tagged S1P1 receptor transfections to increase assay sensitivity, as previously described (38). Transfections were performed using a ratio of 3 μl of FuGENE 6 per μg of plasmid DNA, according to the manufacturer's protocol. Empty pcDNA3.1 vector DNA was added to each transfection as needed to keep the mass of DNA constant. One day after transfection, cells were passed into 35-mm glass bottom dishes (MatTek; Ashland, MA) for confocal microscopy and starved overnight in growth medium supplemented with 0.5% FBS and 10 mm HEPES, pH 7.4. Cells were treated for the time and concentrations indicated in the figure legends, and then fixed for 30 min with 4% paraformaldehyde diluted in PBS, followed by repeated washes with PBS. Confocal microscopy was performed using a Zeiss LSM 510 laser-scanning microscope (Carl Zeiss, Thornwood, NY) with 488-nm excitation and 516–560 nm emission filter sets.

Statistical analysis

Data are expressed as mean ± se and analyzed by Student's t test for unpaired analysis and ANOVA.

Acknowledgments

We thank Charlyne Chassereau and Kent J. Smith for their technical assistance, Department of Medicine and the Hollings Cancer Center Molecular Imaging Facility at Medical University of South Carolina.

This work was supported by NIH Grants HL077192, HL087986 (to A.A.J.), DK55524 (to L.M.L.), the South Carolina COBRE in Lipidomics and Pathobiology (P2 RR17677) (to H.M.E.), and Research Service of the Ralph H. Johnson Veterans Affairs Medical Center (to L.M.L.).

The content of this article does not represent the reviews of the Department of Veterans Affairs or the United States Government.

Disclosure Summary: The authors have nothing to disclose.

Footnotes

Abbreviations:
CTGF
Connective tissue growth factor
DMS
dimethylsphinogsine
GFP
green fluorescent protein
GPCR
G protein-coupled receptor
HMC
human mesangial cell
JNK
c-Jun N-terminal kinase
LDL
low-density lipoprotein
LPPC
lipoprotein preservative/antioxidant cocktail
PMA
phorbol-12 myristate-13 acetate
PTX
pertussis toxin
RMC
rat mesangial cell
S1P
sphingosine-1-phosphate
SK1
sphingosine kinase 1
siRNA
small interfering RNA.

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