Lysophosphatidic acid signaling through its receptor initiates profibrotic epithelial cell fibroblast communication mediated by epithelial cell derived connective tissue growth factor

Abstract

The expansion of the fibroblast pool is a critical step in organ fibrosis, but the mechanisms driving expansion remain to be fully clarified. We previously showed that lysophosphatidic acid (LPA) signaling through its receptor LPA₁ expressed on fibroblasts directly induces the recruitment of these cells. Here we tested whether LPA-LPA₁ signaling drives fibroblast proliferation and activation during the development of renal fibrosis. LPA₁-deficient (LPA₁^-/-) or -sufficient (LPA₁^+/+) mice were crossed to mice with green fluorescent protein expression (GFP) driven by the type I procollagen promoter (Col-GFP) to identify fibroblasts. Unilateral ureteral obstruction-induced increases in renal collagen were significantly, though not completely, attenuated in LPA₁^-/-Col-GFP mice, as were the accumulations of both fibroblasts and myofibroblasts. Connective tissue growth factor was detected mainly in tubular epithelial cells, and its levels were suppressed in LPA₁^-/-Col-GFP mice. LPA-LPA₁ signaling directly induced connective tissue growth factor expression in primary proximal tubular epithelial cells, through a myocardin-related transcription factor-serum response factor pathway. Proximal tubular epithelial cell-derived connective tissue growth factor mediated renal fibroblast proliferation and myofibroblast differentiation. Administration of an inhibitor of myocardin-related transcription factor/serum response factor suppressed obstruction-induced renal fibrosis. Thus, targeting LPA-LPA₁ signaling and/or myocardin-related transcription factor/serum response factor-induced transcription could be promising therapeutic strategies for renal fibrosis.

Keywords: CTGF; LPA(1); fibroblast; fibrosis.

Figure 1. Protection of LPA₁-deficient mice from UUO-induced renal fibrosis

(a) Masson’s trichrome-stained renal sections of LPA₁^+/+Col-GFP mice (upper lane) and LPA₁^−/−Col-GFP mice (lower lane). Representative tissue sections are shown (magnification × 200). Bars, 100 µm. (b) Biochemical analysis of UUO-induced renal fibrosis. Hydroxyproline content was measured in the kidneys of LPA₁^+/+Col-GFP mice and LPA₁^−/−Col-GFP mice ten days after UUO (n = 6 mice/group). Data are expressed as mean ± SEM. (c) Renal expression of COLIα1 mRNA in LPA₁^+/+Col-GFP mice and LPA₁^−/−Col-GFP mice ten days after UUO (n = 6 mice/group). Data are expressed as mean copies of COLIα1 mRNA relative to copies of GAPDH mRNA ± SEM.

Figure 2. UUO-induced renal fibroblast accumulation requires LPA₁

(a) Accumulation of proliferating fibroblasts (GFP⁺PCNA⁺) ten days after UUO. Representative tissue sections stained with anti-GFP antibody/anti-PCNA antibody are shown. Bars, 100 µm. (b) Numbers of GFP⁺ cells in the kidney are expressed as mean ± SEM per HPF (n = 5 mice/group). (c) Numbers of renal GFP⁺PCNA⁺ cells (proliferating fibroblasts) are expressed as mean ± SEM per HPF. (d) Percentages of renal fibroblasts that are proliferating (GFP⁺PCNA⁺ cells/total GFP⁺ cells) are expressed as mean ± SEM per HPF (n = 5 mice/group).

Figure 3. UUO-induced renal αSMA⁺ myofibroblast accumulation requires LPA₁

(a) Accumulation of αSMA⁺ myofibroblasts ten days after UUO. αSMA-stained renal sections of LPA₁^+/+Col-GFP (upper panel) and LPA₁^−/−Col-GFP (lower panel). Representative tissue sections are shown (magnification × 200). Bars, 100 µm. (b) αSMA⁺ areas in the kidney are expressed as mean SEM per HPF (n = 6 mice/group). (c) Renal expression of αSMA mRNA in LPA₁^+/+Col-GFP mice and LPA₁^−/−Col-GFP mice (n = 6 mice/group). Data are expressed as mean copies of αSMA mRNA relative to copies of GAPDH mRNA ± SEM. (d) The expression of αSMA protein in kidney (n = 4 mice/group). Quantification was performed with Image J software and data are expressed as mean dots of αSMA bands relative to dots of GAPDH bands ± SEM. (e) Accumulation of αSMA-expressing fibroblasts (GFP⁺αSMA⁺) ten days after UUO. Representative tissue sections stained with anti-GFP antibody/anti-αSMA antibody are shown. Bars, 100 µm. (f) Percentages of renal fibroblasts that are expression αSMA (GFP⁺αSMA⁺ cells/total GFP⁺ cells) are expressed as mean ± SEM per HPF (n = 5 mice/group).

Figure 4. UUO-induced CTGF expression requires LPA₁, and is predominantly attributable to tubular epithelial cells

(a) Renal expression of CTGF mRNA following UUO for ten days (n = 6 mice/group). Data are expressed as mean copies of CTGF mRNA relative to copies of GAPDH mRNA ± SEM. (b) The production of CTGF protein in kidney ten days after UUO (n = 6 mice/group). Quantification was performed with Image J software and data are expressed mean dots of CTGF bands relative to dots of GAPDH bands ± SEM. (c) The localization of CTGF protein in the kidney. Representative tissue sections are shown (magnification × 200). Bars, 100 µm.

Figure 5. LPA-LPA₁-induced tubular epithelial CTGF drives fibroblast proliferation and αSMA expression

(a, b) LPA induces CTGF mRNA expression in PTECs in a time- and dose-dependent manner (n = 3 cell preparations/group). (c) LPA receptor expression of PTECs. (d) Validation of the inhibitory effects of LPA₁ siRNA and LPA₂ siRNA on the expression of LPA₁ and LPA₂ in PTECS (n = 3 cell preparations/group). (e) Expression levels of LPA-induced CTGF were decreased by knockdown of LPA₁ and LPA₂ by siRNA in PTECs (n = 3 cell preparations/group). (f) Identification of CTGF protein in conditioned media (CM) from PTECs by Western blot. (g, h) Mouse primary renal fibroblasts were transfected with CTGF siRNA, to prevent them from making additional CTGF in response to LPA still present in the CM, and then incubated with CM obtained from PTECs for 48 hours. Fibroblast proliferation and αSMA expression levels were examined (n = 3 cell preparations/group). Data from BrdU proliferation assays are expressed as mean ± SEM of OD value (OD₃₇₀-OD₄₉₂). All data of mRNA expression are expressed as mean ± SEM.

Figure 6. LPA-induced PTEC CTGF expression is dependent on Gα_12/13, Rho, ROCK and actin polymerization

(a) Validation of the inhibitory effects of Gα₁₂ siRNA and Gα₁₃ siRNA on the expression of Gα₁₂ and Gα₁₃ in PTECS (n = 3 cell preparations/group). (b) Expression levels of LPA-induced CTGF were decreased by knockdown of Gα₁₂ and Gα₁₃ by siRNA in PTECs (n = 3 cell preparations/group). (c) Pertussis toxin (PTX)-independent induction of CTGF in response to LPA. PTECs were preincubated with vehicle or 100 ng/ml PTX for 18h followed by the stimulation with 10 µM LPA for 2h (n = 3 cell preparations/group). (d, e) PTECs were preincubated with vehicle (Co), 2.0 µg/ml C3 toxin (C3) for 10h, 5µM Y27632 (Y) for 30 min or 1µg/ml latrunculin B (LB) for 30 min. Cells were then stimulated with 10 µM LPA for an additional 2h (n = 3 cell preparations/group). All data are expressed as mean ± SEM. (f) Immunocytochemical staining for phalloidin in PTECs. PTECs were incubated in serum-free media for 24 hours followed by the stimulation with LPA (10 µM). Some cells were pre-incubated with Y27632 (Y, 5 µM) for 30 min prior to LPA stimulation. All images were captured using identical exposure settings. Scale bars, 25 µm.

Figure 7. LPA promotes the nuclear translocation of MRTF-A and MRTF-B in a ROCK-dependent manner

(a) The subcellular distribution of MRTF-A (upper panel) and MRTF-B (lower panel) in PTECs. PTECs were incubated in serum-free media for 24 hours and then stimulated with LPA (10 µM for 30 min) or control media. PTECs were additionally pre-treated with Y27632 (Y, 5 µM) or control media for 30 min before LPA stimulation. All images were captured using identical exposure settings. (b) Quantification of the subcellular distribution of MRTF-A (upper panel) and MRTF-B (lower panel). Five random fields of view were counted per slide. Subcellular distributions were classified as nuclear (N, nuclear staining > cytoplasmic staining); equal (E, nuclear staining = cytoplasmic staining); or cytoplasmic (C, nuclear staining < cytoplasmic staining). Two independent series of PTECs were analyzed. All data are expressed as mean distribution ± SEM. Scale bars, 25 µm.

Figure 8. LPA enhances MRTF-SRF transcriptional activity through ROCK

(a) LPA induced MRTF-SRF transcriptional activity in PTECs transfected with a transcriptional reporter containing luciferase under the control of a CArG box sequence in a dose-dependent manner (n = 3 cell preparations/group). (b) Pre-treatment with Y27632 (Y, 5 µM for 30 min) reduced the MRTF-SRF transcriptional activity in response to 10 µM LPA (n = 3 cell preparations/group). All data are expressed as mean ± SEM.

Figure 9. LPA-induced PTEC CTGF expression is dependent on MRTF-A, MRTF-B and SRF

(a) Validation of the inhibitory effects of MRTF-A siRNA and MRTF-B siRNA on the expression of MRTF-A and MRTF-B (n = 3 cell preparations/group). (b) Expression levels of LPA-induced CTGF were decreased after knockdown of MRTF-A and MRTF-B by siRNA in PTECs (n = 3 cell preparations/group). (c) Validation of the inhibitory effects of SRF siRNA on the expression of SRF (n = 3 cell preparations/group). (d) Expression levels of LPA-induced CTGF were decreased after knockdown of SRF by siRNA in PTECs (n = 3 cell preparations/group). (e) PTECs were preincubated with 10µM CCG1423 (CCG) for 16h. Cells were then stimulated with 10 µM LPA for an additional 2h (n = 3 cell preparations/group). All data are expressed as mean ± SEM.

Figure 10. Pharmacological inhibition of the MRTF-SRF pathway attenuates UUO-induced renal fibrosis

(a) Representative Masson’s trichrome -stained sections of ligated kidneys. CCG; CCG-203971 (magnification × 200). Bars, 100 µm. (b) Hydroxyproline content in the kidney following UUO for 7 days (n = 5 mice/group). Data are expressed as mean ± SEM. (c–h) Renal expression of COLIα1, CTGF, αSMA, zyxin, vinculin and PDLIM7 following UUO for seven days (n = 5 mice/group). Data are expressed as mean copies of target gene mRNAs relative to copies of GAPDH mRNA ± SEM.

Figure 11. Proposed pro-fibrotic interaction between tubular epithelial cells and fibroblasts in the pathogenesis of renal fibrosis

LPA-LPA₁ signaling contributes to the development of renal fibrosis through interactions between tubular epithelial cells and fibroblasts, in which LPA-LPA₁ signaling stimulates tubular epithelial cells to produce CTGF through a Gα_12/13-Rho-ROCK-MRTF-SRF pathway, and this tubular epithelial cell-derived CTGF in turn induces fibroblast proliferation and αSMA expression.