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*
Medical Clinic IV, University of Erlangen-Nuremberg, Erlangen,
Germany
Imperial College School of Medicine, Molecular Pathology Section, London,
United Kingdom.
Correspondence to Dr. Margarete Goppelt-Struebe, Medizinische Klinik IV, Universität Erlangen-Nürnberg, Loschgestrasse 8, D-91054 Erlangen, Germany. Phone: 49-9131-853-92-01; Fax: 49-9131-853-92-02; E-mail: Goppelt-Struebe{at}rzmail.uni-erlangen.de
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Elevated levels of CTGF are also observed in fibrotic lesions
(8,9,10),
and CTGF is suggested to be functionally involved in the development and
progression of fibrotic diseases. In the kidney, CTGF mRNA levels were
elevated in the majority of biopsies obtained from patients with various types
of renal diseases characterized by glomerulosclerosis and tubulointerstitial
fibrosis (11). Expression of
CTGF in diabetic glomerulosclerosis was recently confirmed in the
db/db mouse model
(12). Furthermore, enhanced
CTGF expression was detected in human and rat mesangial cells when the cells
were stimulated with TGF-ß or exposed to elevated levels of glucose
(12,13).
In addition to mesangial cells, CTGF exhibited positive staining in renal
epithelial cells and interstitial fibroblasts in human biopsies
(11). Most of the interstitial
fibroblasts were characterized as myofibroblasts by the expression of
-smooth muscle actin. These fibroblasts are considered to play a key
role in renal interstitial fibrosis by secreting extracellular matrix
components, which lead to disorganization of the normal organ architecture and
loss of function (reviewed in References
14,15,16).
To date, TGF-ß has been characterized as the main growth factor inducing CTGF, with little effect of other growth factors such as platelet-derived growth factor or fibroblast growth factor (4). Very recent studies indicated that, depending on the cell type, bioactive peptides such as thrombin, factor VIIa, or des-Arg10-kallidin or low-molecular weight mediators such as serotonin or lysophosphatidic acid (LPA) might also induce CTGF expression (16,17,18). Signaling pathways leading to CTGF mRNA have not been analyzed in detail in fibroblasts. In normal rat kidney fibroblasts, elevation of intracellular cAMP levels interfered with TGF-ß-mediated induction of CTGF, whereas tyrosine kinase inhibition by herbimycin or activation of protein kinase C by phorbol ester was without effect (19).
LPA is generated by cleavage from membranes of stimulated cells. It is widely distributed in mammalian tissues and serum, reaching concentrations in the micromolar range (20). Cellular effects of LPA can be categorized as growth-related or cytoskeleton-dependent, resulting in the modulation of adhesion, chemotaxis, contraction, or aggregation (21). As a mitogen for fibroblasts and with additional effects on endothelial cells, macrophages, and vascular smooth muscle cells, LPA has been implicated in wound healing. Because of the role of CTGF in wound healing and fibrosis, it was tempting to speculate that, in addition to other early response genes such as c-fos and egr-1, LPA might induce CTGF. Recent results have demonstrated that, in mesangial cells, LPA is indeed able to induce CTGF expression (18). As a model system for the investigation of CTGF expression, we used an immortalized human renal fibroblast cell line exhibiting the major characteristics of nontransformed primary renal interstitial fibroblasts (22).
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Cell Culture
Immortalized human renal fibroblasts were kindly provided by G. A.
Müller (University of
Göttingen,
Göttingen, Germany)
(22). The cells were grown in
Dulbecco's modified Eagle's medium, supplemented with 2 mM L-glutamine, 4.5
g/L glucose, 100 U/ml penicillin, and 100 µg/ml streptomycin, with 10% FCS.
Renal fibroblasts (0.8 to 1.0 x 106 cells/10 ml) were plated
in 100-mm Petri dishes in medium with 10% FCS. At subconfluence (after 2 to 3
d), cells were serum-starved for 1 d in Dulbecco's modified Eagle's medium
containing 0.5% FCS.
Northern Blot Analysis
Northern blot analysis was performed as described previously
(24). After stimulation for
the indicated times, total RNA was extracted according to the protocol of
Chomczynski and Sacchi (25),
with minor alterations. The RNA yield was usually approximately 60 to 80
µg/10-cm Petri dish. Separation of total RNA (20 µg/lane) was achieved
by using 1.2% agarose gels containing 1.9% formaldehyde, with 20 mM
3-(N-morpholino)propanesulfonic acid, 5 mM sodium acetate, 1 mM
ethylenediaminetetraacetate, pH 7.0 as the gel running buffer. Separated RNA
was transferred to nylon membranes by capillary blotting and was fixed by
baking at 80°C for 1 to 2 h. The 18S and 28S rRNA forms were stained with
methylene blue (0.04% in 500 mM sodium acetate, pH 5.2) and directly
quantitated by densitometry.
Hybridization was performed with cDNA probes labeled with [32P]dCTP, using a NonaPrimer kit (Appligene, Heidelberg, Germany). A cDNA specific for human CTGF was obtained by reverse transcription-PCR amplification of the entire coding region of the gene and was cloned into the pTracer-CMV2 vector (Invitrogen BV, Groningen, The Netherlands). The primer sequences used for the amplification were 5'-GCCAACCATGACCGCCGCCAG-3' (sense) and 5'-TGCCATGTCTCCGTACATCTTCCTG-3' (antisense).
Blots were prehybridized for at least 1 h at 40°C. The probes were allowed to bind overnight at the same temperature. Washing at 40°C was performed for 2 x 15 min under high-salt conditions [2 x SSC/0.2% sodium dodecyl sulfate (SDS)] and for 2 x 15 min under low-salt conditions (0.2x SSC/0.2% SDS). DNA/RNA hybrids were detected by autoradiography using Kodak X-Omat AR film (Eastman Kodak, Rochester, NY). Quantitative analysis was performed by densitometric scanning of the autoradiographs (Bioprofil; Froebel, Wasserburg, Germany). All values were corrected for differences in RNA loading by calculation of the CTGF/18S RNA expression ratio.
Western Blot Analysis
Cellular proteins were isolated using RIPA buffer [50 mM Tris-HCl, pH 7.5
1% (vol/vol) Triton X-100, 0.1% (wt/vol) deoxycholic acid, 0.1% (wt/vol) SDS,
150 mM NaCl, 1 mM phenylmethylsulfonyl fluoride, 1 mM sodium vanadate, 14
µg/ml aprotinin]. For Western blot analysis, 50 to 100 µg of protein
were separated by SDS-polyacrylamide gel electrophoresis (10% polyacrylamide)
and transferred to a polyvinylidene difluoride membrane (Pall Biosupport
Division, Dreieich, Germany). Blots were incubated in 5% skim milk, followed
by overnight incubation with rabbit anti-human CTGF antibody. The membranes
were then washed and incubated for 1 h with horseradish peroxidase-conjugated
swine anti-rabbit IgG (DAKO, Glostrup, Denmark). Bound antibody was detected
with the enhanced chemiluminescence reagent luminol (Autogen Bioclear UK,
Wiltshire, UK).
Staining of Actin Filaments
Cells were cultured and growth-arrested on glass, eight-well, multitest
slides (ICN, Cleveland, OH) placed in a Petri dish. Further treatments with
different toxins and stimuli were performed in wet chambers. After treatment,
cells were fixed with 3% paraformaldehyde in phosphate-buffered saline for 10
min and then permeabilized with 0.2% Triton X-100 in phosphate-buffered saline
for 7 min at room temperature. For examination of DNA fragmentation, cells
were incubated with Hoechst 33258 (8 µg/ml) for 5 min. After washing, the
actin cytoskeleton was stained with rhodamine-phalloidin (Molecular Probes,
Leiden, The Netherlands) for 20 min.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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LPA binds to heptahelical receptors, which couple to different types of G proteins (21). Preincubation of the renal fibroblasts with pertussis toxin did not affect LPA-mediated induction of CTGF (Figure 2A). This finding suggested the involvement of G proteins of the Gq/11 or G12/13 type. The mitogen-activated protein (MAP) kinases p42/44 and p38 are signaling modules for Gi-mediated gene induction by LPA (26,27). Inhibition of p42/44 MAP kinase activation by PD-98059 and of p38 MAP kinase activity by SB-203580 did not significantly reduce the LPA-mediated induction of CTGF (Figure 2, B and C), indicating that neither pathway was essential for CTGF induction.
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RhoA-Mediated Modulation of the Actin Cytoskeleton
LPA was demonstrated to activate the small GTPase RhoA via pertussis
toxin-insensitive signaling
(28). This activation was
related to changes in cell morphologic features attributable to increased
organization of the actin cytoskeleton
(29). Cultured human renal
fibroblasts demonstrated expression of actin stress fibers, which increased
after treatment with LPA (Figure
3A). Inactivation of the GTPases Rho, Rac, and Cdc42 by toxin B
(30) completely disassembled
the actin cytoskeleton within 1 h, comparably to cytochalasin D (which
directly affects actin polymerization)
(Figure 3A). No signs of
apoptosis were detectable with nuclear DNA staining with Hoechst 33258 (data
not shown). Specific inhibition of RhoA by C3 toxin disassembled the actin
stress fibers, whereas the cortical actin fibers remained intact. The pattern
of actin fibers in cells treated with C3 toxin was in accordance with the
induction of lamellipodia with membrane actin ruffles by Rac and the induction
of filopodia with actin microspikes by Cdc42; Rac and Cdc42 are not
inactivated by C3 toxin (31).
Resolution of actin stress fibers was also obtained when the Rho kinases,
which are downstream targets of RhoA, were inhibited by Y-27632
(Figure 3A). Stimulation of the
cells with LPA did not restore the actin cytoskeleton after disassembly by
toxin B, cytochalasin D, or C3 toxin (data not shown). The alterations of the
actin cytoskeleton were also visible as changes in cell morphologic features
detected by light microscopy. As an example, contraction of the cells and
development of a more spindle-like appearance after treatment with C3 toxin
are shown in Figure 3B. The
phenotype observed after treatment with C3 toxin was not reversed by
subsequent treatment with LPA.
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Involvement of RhoA in the LPA-Mediated Induction of CTGF mRNA
Interference with the activation of the Rho proteins RhoA, Rac, and Cdc42
by toxin B completely prevented LPA-mediated CTGF mRNA induction
(Figure 4, A and D). Similarly,
the basal expression of CTGF was abolished after treatment with toxin B.
Baseline levels were barely affected by cytochalasin D, whereas destruction of
the actin cytoskeleton by this compound prevented LPA-mediated induction of
CTGF (Figure 4, A and D).
Interference with CTGF mRNA induction was specifically related to inhibition
of RhoA, as indicated by the strong inhibitory effect of C3 toxin, which
specifically targets RhoA without effects on Rac or Cdc42
(Figure 4, B and D). Various
proteins are involved in RhoA-mediated regulation of the actin cytoskeleton,
including the Rho kinase family
(32). Inhibition of these
kinases by Y-27632 reduced CTGF expression, with maximal inhibition being
observed with 5 to 10 µM Y-27632 (Figure
4, C and D). Apparent changes in basal CTGF mRNA expression did
not prove to be statistically significant.
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Regulatory Roles for RhoA and the Actin Cytoskeleton in
TGF-ß-Mediated Induction of CTGF
Disruption of the cytoskeleton by inhibition of the Rho kinases by Y-27632
or cytochalasin D also affected TGF-ß-induced CTGF mRNA expression. When
the cells were preincubated with either compound for 45 min and then
stimulated with TGF-ß for 6 h, induction of CTGF was almost completely
prevented (Figure 5).
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Reversal of cAMP-Mediated Changes in Cell Morphologic Features by
LPA
Elevated levels of cAMP and subsequent activation of protein kinase A (PKA)
also led to the disassembly of actin stress fibers in fibroblasts
(33). Incubation of human
renal fibroblasts with the cell-permeable cAMP analogue cBIMPs, which
specifically activates PKA, or the adenylyl cyclase activator forskolin
induced destruction of the actin fibers in <1 h
(Figure 6A). Subsequent
treatment with LPA completely restored the actin filaments. The disassembly of
the actin cytoskeleton produced by elevated cAMP levels led to changes in cell
shape, characterized by rounding of the cell bodies and the development of
elongated processes. As an example, cells treated with forskolin are shown in
Figure 6B. After treatment with
LPA, the cells flattened and became indistinguishable from untreated control
cells.
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Interference with LPA-Mediated Induction of CTGF by PKA
Activation
Incubation of the renal fibroblasts with forskolin or prostaglandin
E1, which also elevated intracellular cAMP levels by stimulating
adenylyl cyclase, reduced the basal expression of CTGF. LPA-mediated induction
of CTGF was greatly impaired when the cells were preincubated with either
forskolin or prostaglandin E1
(Figure 7, A and C). Similar
effects were observed when the cells were incubated with cell-permeable
analogues of cAMP, i.e., 8-(4-chlorophenylthio)-cAMP or cBIMPs
(34)
(Figure 7, B and C). Again, LPA
did not overcome the inhibitory effect of elevated cAMP levels.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Rho proteins are involved in gene expression but are also prominent regulators of the actin cytoskeleton. Rho-associated serine/threonine kinase isozymes (Rho kinases) regulate the polymerization of stress fibers via inactivation of myosin light chain phosphatase (38). Inhibition of these kinases with the specific inhibitor Y-27632 (39) also impaired CTGF induction, suggesting a link between cytoskeletal integrity and the induction of CTGF. This was confirmed when the actin stress fibers were depolymerized by the direct action of cytochalasin D. Cytochalasin D impaired LPA-mediated CTGF induction to a similar extent, compared with C3 exotoxin. An important role of RhoA and the cytoskeleton in the regulation of CTGF was recently observed in mesangial cells, which in many other respects exhibit different signaling properties, compared with fibroblasts (18). As an example, induction of the early response gene cyclooxygenase 2 was positively regulated by the viral kinase pp60v-src in fibroblasts (40,41), whereas cyclooxygenase 2 was not inducible in v-src-transformed mesangial cells (42). Similarly, elevation of cAMP levels had no effect on cyclooxygenase 2 expression in mesangial cells (43) but induced cyclooxygenase 2 expression in fibroblasts (44). Regulation of CTGF by RhoA, in contrast, does not seem to be restricted to a specialized cell type.
Cytoskeletal changes were also observed when the intracellular levels of cAMP were elevated by cell-permeable cAMP analogues or by adenylyl cyclase activation by forskolin. A similar actin pattern was observed when the cells were treated with prostaglandin E1, suggesting activation of PKA via EP2 or EP4 receptors (45). The morphologic changes were reversed when the PKA-activated cells were subsequently treated with LPA; in <1 h, the cells rounded and the actin cytoskeleton reorganized into stress fibers. Interactions between cAMP-activated pathways and Rho-mediated structural changes have been observed in different cell types. In epithelial cells, activation of PKA caused rapid changes in cell morphologic features, which were attributable to inactivation of RhoA. Pre-treatment of these cells with thrombin, which activated RhoA, prevented PKA-mediated morphologic changes (46). In these cells, the thrombin pathway dominated the cAMP pathway. The opposite was observed in PC12 cells, where elevated cAMP levels protected cells against LPA-mediated neurite retraction (47). The interaction between LPA and PKA seems to be dependent on the cell type, because LPA was able to reverse the morphologic changes caused by cAMP in astrocytes (48). Recombinant RhoA is a target of PKA, and phosphorylation of Ser-188 increased the interaction with the guanine nucleotide dissociation inhibitor, which translocates RhoA from the membrane to the cytosol (49). In line with this interpretation, elevated levels of cAMP resulted in enhanced cytosolic levels of RhoA and seemed to prevent membrane association (50). Activation of PKA thus results in inactivation of RhoA and may thus oppose the effects of RhoA activators such as LPA. The morphologic observations for renal fibroblasts are in agreement with the changes observed in astrocytes, because LPA was able to reverse the effects of cAMP in both cell types.
In contrast to the morphologic changes, however, LPA was not able to overcome the cAMP-mediated blockade of the signaling pathway leading to gene expression; LPA-mediated CTGF expression was inhibited by all compounds that elevated cAMP levels. If an equilibrium between phosphorylated and nonphosphorylated RhoA is assumed, then LPA might be able to shift the equilibrium to the nonphosphorylated form, thus reversing the morphologic effects. Because nothing is known regarding the phosphatases mediating RhoA dephosphorylation or the kinetics of those reactions, it might be speculated that the transition is too slow to allow the rapid signaling that induces CTGF mRNA expression. Furthermore, we cannot exclude additional, RhoA-independent mechanisms by which PKA might interfere with LPA-mediated CTGF expression.
The regulatory role of RhoA is not unique for LPA signaling. TGF-ß-mediated induction of CTGF was strongly reduced when RhoA signaling was inhibited by a Rho kinase inhibitor or when the actin cytoskeleton was directly disassembled by cytochalasin D. These data are in agreement with data published previously; in H-ras-transformed fibroblasts, inhibition of RhoA by C3 exotoxin abrogated the ability of TGF-ß to induce actin stress fibers (51). Furthermore, elevated levels of cAMP blocked TGF-ß-induced changes in cell morphologic features and CTGF expression in normal rat kidney fibroblasts (19). Recent data obtained in a different cell type, i.e., renal mesangial cells, also indicated that inhibition of Rho proteins interfered with TGF-ß-mediated CTGF induction (18). Therefore, the regulatory roles of RhoA and the cytoskeleton are not restricted to a specific stimulus or cell type but seem to represent more general features of CTGF regulation.
Given the role of CTGF in matrix synthesis and its proposed role in the development of fibrosis, targeting of RhoA might be a way to interfere with these processes. Rho proteins are inactivated by various bacterial toxins, as used in this study. At the pharmacologic level, hydroxymethyl glutaryl-CoA reductase inhibitors might be more appropriate. In addition to their lipid-lowering effects, these compounds interfere with the isoprenylation, and thus the activation, of Rho proteins (52). Preliminary data indicate that lovastatin and simvastatin, two clinically used statins, indeed interfere with CTGF synthesis.
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Acknowledgments |
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References |
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