Mesangial cells secrete various growth factors and cytokines that may regulate endothelial and epithelial cells. Hepatocyte growth factor (HGF) was initially identified as the most potent growth factor for hepatocytes1,2, and is well known as a mesenchyme-derived pleiotropic factor that regulates cell growth, cell motility, and morphogenesis of various types of cells. HGF is also considered to be a humoral mediator of epithelial-mesenchymal interactions responsible for morphogenic tissue interactions during embryonic development and organogenesis3,4,5. Of importance, recent studies suggest that HGF has many actions on the cells of other target organs including the kidney6,7,8,9. Acute renal failure is often reversible, and recovery depends on mitogensis, motogensis and morphogensis (tubular formation) of renal epithelial cells. Thus, rapid regeneration of renal epithelial cells might be important for the treatment of acute renal failure. HGF enhances renal regeneration and suppresses the onset of acute renal failure caused by renal toxins, renal ischemia, or unilateral nephrectomy6,7,8,9. For example, administration of recombinant HGF (rHGF) promoted the regeneration of epithelial cells injured by anti-tumor drugs7. HGF mRNA and blood HGF levels increase markedly after unilateral nephrectomy and in acute renal failure8,9. However, little is known about the local HGF system in the kidney and the mechanisms of the protective actions of HGF in renal cells. As it is apparent that cell-cell interactions among these cells are important in the control of renal function, we wanted to determine the exact mechanisms of maintaining the cell-cell interactions in the kidney, given the presence of a local renal HGF system. On the other hand, it has been hypothesized that endothelial cells may also modulate mesangial cell growth, because many anti-proliferative factors such as nitric oxide (NO) and vascular natriuretic peptides are secreted by endothelial cells10,11,12. It is apparent that dysfunction of endothelial and/or epithelial cells may promote abnormal mesangial cell growth, for example, in glomerulonephritis13,14,15,16. Previously, we have reported that local HGF production by mesangial cells maintains the growth of glomerular endothelial cells17, suggesting the potential contribution of HGF to the maintenance of renal function. In the current study, we examined whether: (1) HGF has a protective role in epithelial and endothelial injury, and (2) HGF has anti-apoptotic actions in the prevention of endothelial and epithelial cell death.
METHODS
In vitro experiments
Cell culture
Rat mesangial cells were donated by Kirin Brewery Co., Ltd. (Tokyo, Japan)17. These cells were used within passages 5 to 7. Rat epithelial cells (NRK-52E) and porcine tubular epithelial cells (LLC-PK1) were purchased from ATCC (American Tissue Culture Collection). Rat mesangial and NRK-52E cells were maintained in Dulbecco's modified Eagle's medium (DMEM) supplemented with 10% fetal calf serum, 100 U/ml penicillin and 100 mg/ml streptomycin. LLC-PK1 cells were maintained in Medium 199 supplemented with 5% fetal calf serum, 100 U/ml penicillin and 100 mg/ml streptomycin. Human endothelial cells (passage 5) were obtained from Clonetics Corp. (San Diego, CA, USA), and cultured in modified MCDB131 medium supplemented with 5% fetal calf serum, 100 U/ml penicillin, 100 
g/ml streptomycin, 10 ng/ml epidermal growth factor, 2 ng/ml basic fibroblast growth factor (bFGF) and 1 M dexamethasone in the standard fashion18. Cells were incubated at 37°C in a humidified atmosphere of 95% air-5% CO2 with media changes every two days. These cells showed the specific characteristics of mesangial, epithelial and endothelial cells, respectively, on immunohistochemical examination and morphological observations.
Cell counting assay
In the preparation of experiments for determination of cell count, the cells were grown to 70% confluence in 96-well culture plates (Becton Dickinson Co., Belgium). After cells reached 70% confluence, the medium was changed to fresh, defined serum-free medium (DSF). DSF containing insulin (5 
10-7 M), transferrin (5 mg/ml), and ascorbate (0.2 mM) maintains cells in a non-catabolic state, as previously reported19. The cells were then incubated overnight. On day 1, the medium was again changed to fresh DSF containing HGF (10 ng/ml), vascular endothelial cell growth factor (VEGF) (10 ng/ml) or vehicle. On day 3, the medium was again changed to fresh DSF containing growth factor or vehicle. On day 4, an index of cell proliferation was determined using WST-cell counting kit (Wako, Osaka, Japan). Tetrazolium salt has been used to develop a quantitative colorimetric assay for cell growth. The assay detects living, but not dead cells. For this purpose, 3-(4,5-dimethylthiazol-2-yl)-2,5,-diphenyl tetrazolium bromide (MTT) is widely used20. In this study, we used an alternative of MTT, that is, sulfonated tetrazolium salt, 4-[3-(4-iodophenyl]-2-(4-nitrophenyl)-2H-5-tetrazolio]-1,3-benzene disulfonate (WST-1), since this compound produces a highly water-soluble formazan dye that makes the assay procedure easier to perform20. The plates were read on a Bio-Rad Model 3550 Microplate reader, using a test wavelength of 450 nm, and a reference wavelength of 650 nm. We confirmed that serum-stimulated increase in cell number is associated with increased absorbance at 450 nm.
Treatment by serum deprivation
In the preparation of experiments for determination of cell death, epithelial cells were grown to confluence. After cells reached confluence, the medium was changed to fresh, defined serum-free (DSF) medium containing HGF, basic fibroblast growth factor (bFGF), VEGF, or vehicle. The cells were then incubated. Every two days the medium was changed to fresh DSF medium containing HGF, bFGF, VEGF, or vehicle. On days 2 and 4, an index of cell proliferation was determined as described above.
Apoptotic cells induced by serum deprivation were also counted at 48 and 96 hours. Cells to be analyzed for apoptosis were stained with propium iodide (PI) and Hoechst 33342, and viewed under fluorescence microscopy as previously described21,22,23,24. Using both membrane-permeable (H33342) and -impermeable (PI) dyes in the assay allowed the determination of cell viability and plasma membrane integrity, and an accounting for any non-apoptotic toxic or necrotic death induced in the study groups. Cells were seeded onto six-well dishes (Lab-Tek) and were cultured in DSF medium for two days, after reaching confluence. To stain the cells for DNA, cells were incubated with Hoechst 33342 (5 g/ml in PBS) for 20 minutes at 37°C. The medium and a PBS rinse of the culture wells were collected before brief addition and decantating of trypsin/EDTA. Culture wells were incubated in residual trypsin/EDTA for three minutes in humidified air at 37°C to achieve maximal cell detachment before rinsing with PBS and collection. The collected medium, including the rinse and trypsinized cells, were pooled and collected by centrifugation at 1000 rpm for five minutes at 4°C. Cell pellets were resuspended in a small volume of serum-containing medium with 5 g/ml Hoechst 33342 and 1 g/ml PI. An aliquot was placed on a glass slide, covered with a glass coverslip, and viewed under fluorescence microscopy. Individual nuclei were visualized at 400 to distinguish the normal uniform nuclear pattern from the characteristic condensed coalesced chromatin pattern of apoptotic cells. For each sample, 300 cells were examined. The number of apoptotic cells was counted under microscopy (magnification, 100) in a blinded manner. The total number of apoptotic cells in each section was summed and expressed as a percentage of the total cell number. At least ten individual sections were evaluated per slide. Samples were coded so that the analysis was performed without knowledge of which treatment each cells had received. The reproducibility of the results was assessed. Intraobserver variability was determined from triplicate measurements performed by one observer for all sections. The mean 
SD difference among measurements made by the same observer was 2.6 0.6%. Interobserver variability was determined from measurements of 10 randomly selected sections performed by a second observer in addition to the first observer. The difference between measurements made by the two observers was 3.0 0.8%. These observers were blinded to other data concerning the cells, as well as to the results of the other observer.
Also, we employed the measurement of cellular DNA fragmentation using cellular DNA Fragmentation ELISA kit (Boehringer Mannheim, Germany), to quantity apoptosis25. Cultured epithelial cells were incubated with 10 M BrdU overnight at 37°C in 5% CO2. At four days after transfection, lysing solution containing BSA, EDTA, and Tween 20 was added to each well. DNA fragments in 100 l cell lysate supernatant were tested by ELISA. The supernatant was transferred to an anti-DNA-precoated microtiter plate and incubated for 60 minutes at 37°C. After washing, the samples were denatured and fixed by microwave irradiation for five minutes. After cooling the microtiter plate for 10 minutes at -20°C, anti-BrdU peroxidase-conjugated solution was added and the plate was incubated for 60 minutes at 37°C. Wells were again washed, TMB substrate solution was added, and the plate was incubated for 30 minutes at room temperature. Stopping solution (25 l 1 M H2SO4) was then added to each well. Absorbance was measured at 450 nm (reference wavelength, 690 nm). Briefly, 10,000 apoptotic cells/well reflects absorbance 1.5 in the manufacturer's recommended condition. The sensitivity of DNA fragmentation ELISA assay is well correlated with the results from the conventional3H-thymidine based DNA fragmentation assay. In our experimental conditions, an increase in absorbance by 0.2 reflected an increase in the cell number from 2,000 apoptotic cells/well.
Preparation of HVJ-liposomes
To produce an HGF expression vector, human HGF cDNA (2.2 kb) was inserted into the Eco RI and Not I sites of pUC-SRa expression vector plasmid. In this plasmid, transcription of the HGF cDNA was under the control of the SR
promoter1,26,27. We employed the hemagglutinating virus of Japan (HVJ)-liposome method27,28,29. Briefly, phosphatidylserine, phosphatidylcholine, and cholesterol were mixed in a weight ratio of 1:4.8:2 in tetrahydrofuran. The lipid mixture (10 mg) was deposited on the sides of a flask by removal of the solvent in a rotary evaporator. High mobility group (HMG) 1 nuclear protein (96 g), purified from calf thymus, was mixed with plasmid DNA (300 g) in 200 l balanced salt solution (BSS; 137 mM NaCl, 5.4 mM KCl, 10 mM Tris-HCl, pH 7.6) at 20°C for one hour, and then added to the dried lipid. Liposome-DNA-HMG 1 complex suspension was mixed by vortex, sonication for three seconds, and shaking for 30 minutes. Purified HVJ (Z strain) was inactivated by UV irradiation (110 erg/mm2/sec) for three minutes immediately prior to use. The liposome suspension (0.5 ml, containing 10 mg lipid) was mixed with HVJ (20,000 hemagglutinating units) in a total volume of 4 ml BSS. The mixture was incubated at 4°C for 10 minutes and then for 30 minutes with gentle shaking at 37°C. Free HVJ was removed from the HVJ-liposomes by sucrose density gradient centrifugation. The top layer of the sucrose gradient containing the HVJ-liposome-DNA complex was collected and used immediately.
Epithelial cells (1 106) were seeded onto six-well plates (Corning, NY, USA) and grown to 80% confluence. Cells were washed three times with BSS containing 2 mM CaCl2 and then incubated with 1 ml HVJ-liposomes-DNA complex (2.5 mg lipid and 10 g encapsulated DNA) at 4°C for five minutes followed by 37°C for 30 minutes (total, 35 min). The cells were then washed and fed fresh medium containing 10% calf serum and placed in a CO2 incubator. To document the successful transfection of cells, we examined the production of HGF. Twenty-four hours after transfection, the medium was changed to fresh DSF and the cells were incubated for an additional 48 hours. To study the release of HGF, transfected cells (48 hr post-transfection) were washed and fed with 1 ml DSF. Twenty-four hours later, conditioned medium was collected, centrifuged at 600 g for 10 minutes and stored at -20°C28. The concentration of HGF in the medium was determined by enzyme-immunoassay using anti- human HGF antibodies27,30. On day 4, an index of cell proliferation was determined using WST-cell counting kit (Wako). The number of apoptotic cells was also counted in a blinded manner.
Reverse transcription-polymerase chain reaction
RNA was extracted from each cell type by RNAzol (Tel-Test Inc., TX, USA), after cells reached confluence under serum stimulation. The level of HGF receptor (c-met) mRNA was measured by reverse transcription-polymerase chain reaction (RT-PCR)31. The 5' primer 2788-2811 complementary to the rat c-met gene was 5'-CAG-TGA-TGA-TCT-CAA-TGG-GCA-AT-3'; the 3' primer 3492-3504 was 5'-AAT-GCC-CTC-TTC-CTA-TGA-CTT-C-3'CAA-TGG-GGG. Extreme care was taken to avoid contamination of tissue samples with trace amounts of experimental RNA. Aliquots of RNA (0.5 g) derived from cultured cells were amplified simultaneously by PCR (35 cycles), with the same reagents, by individuals who were blinded to the identity of the samples, and compared with a negative control (primers without RNA). Amplification products were electrophoresed through 2% agarose gel and stained with ethidium bromide. To ensure that the RT-PCR amplified product reflects transcribed c-met RNA without significant DNA contamination, RNA samples treated with RNase A or amplified without reverse transcriptase were amplified simultaneously as negative controls. These samples did not result in a visual band. Moreover, PCR products were cut by restriction enzymes, and the fragments were identical to the theoretical bands. At least three aliquots of each DNA and RNA sample were subjected to separate PCR amplification in all experiments.
Measurement of hepatocyte growth factor in conditioned medium
Rat mesangial and epithelial cells were seeded on six-well plates (Corning) at a density of 5 104 cells/cm2 and cultured for 24 hours. After replacing the medium with fresh DSF and following culture for 24 hours, the concentration of HGF in the medium was determined by enzyme-immunoassay using anti-rat HGF antibody. This EIA specifically detects rat HGF, because of lack of cross-reactivity against human HGF18,27,30,31.
Co-culture of mesangial cells with epithelial cells
Cells to be co-cultured were seeded onto cell culture inserts (3.0 mM pore size; Becton Dickinson & Company, Belgium) and grown in 10% DMEM. Cells were seeded onto six-well plates (Becton Dickinson), maintained in 10% fetal calf serum and placed in DSF medium for 48 hours following 80% confluence. At confluence, the inserts containing co-cultured cells were put into the wells containing the quiescent cells17,28. Epithelial cells in the lower well were co-cultured for 48 hours with mesangial cells in DSF with 0.5% serum, and cell numbers were assessed by cell counting assay.
Effect of neutralizing anti-hepatocyte growth factor antibody
For each of the antibodies, the IgG fraction (purified with protein A-agarose) was able to neutralize a biological activity of 10 ng/ml HGF, at a concentration of 10 g/ml17. Normal rabbit serum IgG fraction (10 g/ml) was employed as a control. The number of epithelial cells was examined in two groups: group 1, with anti-rat HGF antibody (final 10 g/ml), and group 2, with normal IgG (final 10 g/ml). Neutralizing anti-rat HGF antibody can attenuate only the effects of rat and murine HGF, but not human, bovine or porcine HGF.
Determination of DNA and RNA synthesis
Endothelial cells were seeded onto 24-well tissue culture plates (Corning). At confluence, endothelial cells were rendered quiescent by incubation for 48 hours in DMEM with 0.5% fetal bovine serum. Relative rates of DNA and RNA synthesis were assessed by determination of3H-thymidine or uridine incorporation into trichloroacetic acid (TCA)-precipitable material over the next 24 hours, respectively. HGF or vehicle (fresh DSF containing 0.1% bovine serum albumin) was added 12 hours prior to the addition of3H-thymidine or uridine. Twenty-four hours after the addition of3H-thymidine or uridine, the cells were washed twice with cold PBS, twice with 10% (wt/vol) cold TCA and incubated with 10% TCA at 4°C for 30 minutes. Cells were rinsed in ethanol (95%) and dissolved in 0.25 N NaOH at 4°C for three hours, neutralized, and the radioactivity was determined by liquid scintillation spectrometry28.
Materials
Human and rat recombinant HGF were purified from the culture medium of Chinese hamster ovary cells or C-127 cells, respectively, transfected with expression plasmid containing human or rat HGF cDNA1,26. VEGF and bFGF were obtained from Biosource (Camarillo, CA, USA).
Statistical analysis
All values are expressed as mean SEM. Analysis of variance with subsequent Bonferroni's test was employed to determine the significance of differences in multiple comparisons. Values of P < 0.05 were considered statistically significant.
RESULTS
Effect of recombinant hepatocyte growth factor on epithelial cell death induced by serum deprivation
First, the effect of rat recombinant HGF on the growth of rat epithelial cells (NRK-52E) was examined Figure 1a. The addition of rHGF as well as VEGF stimulated growth of epithelial cells. The specificity of the stimulatory effects of HGF was confirmed by the blockade of cell growth by anti-HGF specific antibody (data not shown). Similarly, rat rHGF stimulated the growth of porcine tubular epithelial cells LLC-PK1; Figure 1b. Previously, we have reported that rat and human rHGF had no mitogenic actions on rat and human mesangial cells, whereas rHGF could stimulate growth of endothelial cells17,32,33. These results indicate that HGF can exert stimulatory effects on growth of epithelial and endothelial cells without replication of mesangial cells. Next, we examined the effect of HGF treatment on cell death induced by serum deprivation and discovered that serum deprivation caused epithelial cell death Figure 2. Twelve hours after serum deprivation, some cells started to become round and eventually detached from the plate, floating in the medium and leaving many holes in the sheet of confluent cells (data not shown). The floating cells could be recovered with the medium, and neither attached onto a new plate nor proliferated. Consistent with this morphological observation, the cell death rate after serum deprivation was significantly increased at 96 hours, as shown in Figure 2. The addition of rHGF (10 ng/ml) resulted in partial attenuation of cell death mediated by serum deprivation in a dose-dependent manner at 96 hours Figure 2. VEGF as well as bFGF also partially prevented epithelial cell death induced by serum deprivation treatment. Cell death injury due to serum deprivation was predominant by apoptosis, consistent with the previous reports34,35. As shown in Figure 3a, morphological studies demonstrated the typical features of apoptotic cells. Indeed, serum deprivation resulted in a significant increase in apoptotic epithelial cells as assessed by morphology Figure 3. Serum deprivation significantly increased the apoptotic cells, which was abolished by addition of serum as assessed by Hoechst staining (P < 0.01). Importantly, the addition of rHGF also prevented apoptosis of epithelial cells induced by serum deprivation in a dose-dependent manner both at 48 and 96 hours Figure 3. Similarly, bFGF as well as VEGF prevented the apoptosis (P < 0.05) both at 48 and 96 hours. Similar results were obtained using PI staining (data not shown). These results were confirmed by the measurement of DNA fragmentation Figure 4. Consistent with nuclear staining, the addition of rHGF significantly decreased the DNA fragmentation index both at 10 and 100 ng/ml of concentration under serum free conditions (P < 0.01). Prevention of apoptosis by HGF was also confirmed using LLC-PK1 cells assessed by Hoechst staining Figure 5.
Figure 1.
Effect of exogenously added rat recombinant hepatocyte growth factor (HGF) on numbers of rat epithelial cells (NRK-52E) (A) and porcine tubular epithelial cells (LLC-PK1) (B). N = 8 per group. Abbreviations are: DSF, vehicle added to cells; HGF, rat rHGF (10 ng/ml) added to cells; VEGF, recombinant vascular endothelial growth factor (10 ng/ml) added to cells. **P < 0.01 versus DSF.
Full figure and legend (53K)Figure 2.
Effects of exogenously added hepatocyte growth factor (HGF), vascular endothelial growth factor (VEGF) and basic fibroblast growth factor (bFGF) on epithelial cell number under serum deprivation. Values are expressed as percentage of cell survival as compared to serum (5% fetal calf serum) treatment. N = 8 per group. Abbreviations are: DSF, vehicle; HGF, human recombinant HGF (1, 10, 100 ng/ml) added to epithelial cells maintained in DSF; VEGF, human recombinant VEGF (100 ng/ml) added to epithelial cells maintained in DSF; FGF, human recombinant bFGF (100 ng/ml) added to epithelial cells maintained in DSF. **P < 0.01 versus serum (5% fetal calf serum) treatment, #P < 0.05, ##P < 0.01 versus DSF.
Full figure and legend (37K)Figure 3.
(A) Morphological changes in nuclei of rat epithelial cells (NRK-52E) treated with serum deprivation. Epithelial cells maintained in DSF exhibited the characteristic features of cell shrinkage, membrane blebbing and rounding, typical of apoptotic death. rHGF as well as serum treatment attenuated these morphological changes. Abbreviations are: FCS, epithelial cells treated with 5% fetal calf serum; serum-free, epithelial cells maintained in DSF; rHGF, epithelial cells treated with DSF medium and human rHGF (100 ng/ml) added to epithelial cells maintained in DSF. **P < 0.01 versus serum (5% fetal calf serum) treatment, #P < 0.05, ##P < 0.01 versus DSF. (B and C, opposite page) Protective effects of exogenously added HGF and VEGF on apoptosis of rat epithelial cells (NRK-52E) under serum deprivation at 48 hours (B) and 96 hours (C) assessed by Hoechst staining. Values are expressed as percentage of apoptotic cells. N = 8 per group. Abbreviations are: DSF, vehicle added to epithelial cells maintained in DSF; rHGF, human rHGF (10 and 100 ng/ml) added to epithelial cells maintained in DSF; 5% FCS, 5% fetal calf serum added to epithelial cells. **P < 0.01 versus DSF.
Full figure and legend (60K)Figure 4.
DNA fragmentation in rat epithelial cells (NRK-52E) treated with rHGF and rVEGF under serum deprivation at 96 hours assessed by DNA fragmentation assay (absorbance at OD 690 nm). N = 6 per group. Abbreviations are: DSF, vehicle; HGF, human rHGF (10 and 100 ng/ml) added to epithelial cells maintained in DSF; VEGF, human recombinant VEGF (100 ng/ml) added to epithelial cells maintained in DSF. **P < 0.01 versus DSF.
Full figure and legend (34K)Figure 5.
Protective effects of exogenously added HGF and bFGF on apoptosis of porcine epithelial cells (LLC-PK1) under serum deprivation at 96 hours assessed by Hoechst staining. Values are expressed as percentage of apoptotic cells. N = 8 per group. Abbreviations are: DSF, vehicle added to epithelial cells maintained in DSF; rHGF, human rHGF (1 and 10 ng/ml) added to epithelial cells maintained in DSF; bFGF, human bFGF (1 and 10 ng/ml) added to epithelial cells maintained in DSF.**P < 0.01 versus DSF, ##P < 0.01 versus rHGF 1 ng/ml.
Full figure and legend (29K)To examine the therapeutic value of HGF as gene therapy in epithelial injury, transfection of human HGF gene was also examined. Rat epithelial cells transfected with the human HGF expression vector synthesized and secreted immunoreactive HGF. The accumulation of immunoreactive human HGF was readily detected in conditioned medium from epithelial cells transfected with HGF expression vector, but not in conditioned medium from untransfected cells or cells transfected with control vector (untransfected; NS; control, NS; HGF, 0.865 0.056 ng/106 cells/24 hr, P < 0.01). The autocrine influence of transfection of HGF vector on epithelial injury is shown in Figure 6. Transfection of epithelial cells with the control expression vector did not alter cell number, compared with cell number in untransfected epithelial cells. However, transfection of human HGF vector attenuated cell death caused by serum deprivation (untransfected, 47,600 4000; control vector, 50,600 1900; HGF vector, 57,400 3200 cells/well, P < 0.01 vs. control vector and untransfected). Moreover, the number of apoptotic cells in cells transfected with HGF expression vector was significantly lower than that in those transfected with control expression vector (P < 0.01; Figure 6). The prevention of apoptosis by transfection of HGF vector was similar to that obtained by addition of 10% fetal calf serum.
Figure 6.
Protective effects of transfected HGF cDNA on number of apoptotic cells in NRK-52E cells. Abbreviations are: DSF, untransfected cells; CV, cells transfected with control vector; HGF, cells transfected with human HGF vector; 10% FCS, cells treated with 10% FCS. N = 6 per group. **P < 0.01 versus DSF, ##P < 0.01 versus CV.
Full figure and legend (30K)Protective action on human endothelial cells
As previously reported, HGF also has a protective action against endothelial injury32,34. Therefore, we further examined the protective actions of HGF against apoptosis of endothelial cells. Under basal conditions, human rHGF as well as VEGF increased the number of endothelial cells in a dose-dependent manner Figure 7a. Our previous study showed that the signal transduction of HGF is different from that of VEGF33. Therefore, we tested the effects of HGF and VEGF on the growth of endothelial cells under TNF- treatment, which is known to mediate apoptosis35,36. Incubation with TNF- resulted in a decrease in DNA synthesis of endothelial cells Figure 7b. Importantly, the addition of human rHGF also significantly attenuated the decrease in DNA synthesis induced by TNF-, and was similar to 10% FCS. In contrast, VEGF failed to show an attenuation of the decrease in DNA synthesis (P < 0.01), whereas VEGF stimulated basal endothelial cell growth. The protective action of HGF was also examined under dexamethasone treatment, as dexamethasone is also known to be a mediator of endothelial injury through apoptosis37. Treatment with dexamethasone also resulted in a decrease in DNA and RNA synthesis of endothelial cells Figure 8. In contrast, the addition of human rHGF resulted in attenuation of the decrease in DNA and RNA synthesis induced by dexamethasone treatment (P < 0.05).
Figure 7.
(A) Effects of exogenously added HGF and VEGF on human endothelial cell number. N = 8 per group. Abbreviations are: DSF, vehicle; HGF, human recombinant HGF (10 and 100 ng/ml) added to endothelial cells; VEGF, human recombinant VEGF (10 and 100 ng/ml) added to endothelial cells; FCS, 10% FCS added to endothelial cells. **P < 0.01 versus vehicle. (B) Effect of exogenously added HGF and VEGF on human endothelial DNA synthesis under TNF- treatment. N = 8 per group. Abbreviations are: DSF, vehicle; HGF, human recombinant HGF (10 ng/ml) added to endothelial cells; VEGF, human recombinant VEGF (10 ng/ml) added to endothelial cells; TNF, human recombinant tumor necrosis factor- (20 nM) added to endothelial cells; FCS, 10% fetal calf serum added to endothelial cells. **P < 0.01 versus DSF, #P < 0.05 versus TNF.
Figure 8.
Effect of exogenously added recombinant HGF on human endothelial cell (A) DNA and (B) RNA synthesis under dexamethasone treatment. N = 8 per group. Abbreviations are: DSF, vehicle; HGF, human recombinant HGF (10 ng/ml) added to endothelial cells; DX, dexamethasone (10-7 M) added to endothelial cells. **P < 0.01 versus DSF, #P < 0.05 versus TNF.
Full figure and legend (28K)Role of hepatocyte growth factor in the regulation of epithelial cells
Hepatocyte growth factor was detected in the conditioned medium of rat mesangial cells by EIA (0.23 0.01 ng/106 cells/24 hr), and was consistent with the previous findings17. On the other hand, we failed to detect immunoreactive HGF in the culture medium of rat epithelial cells (NRK-52E) by EIA. In contrast, the presence of its specific functional receptor (c-met) was readily detected by RT-PCR in both rat (NRK-52E) and porcine epithelial cells (LLC-PK1; Figure 9). Given the presence of a local HGF system in mesangial cells, we hypothesized that local HGF production by mesangial cells may regulate epithelial cells. Using a co-culture system, we further examined this hypothesis that local HGF secretion from mesangial cells regulates the growth of epithelial cells. Co-culture of rat mesangial cells with rat epithelial cells (NRK-52E) resulted in a significant increase in cell number (P < 0.01; Figure 10). Co-incubation of anti-rat HGF antibody in a co-culture system of rat mesangial cells with NRK-52E significantly abolished the mitogenic activity of conditioned medium from mesangial cells (P < 0.01). Moreover, co-culture of rat mesangial cells with porcine tubular epithelial cells (LLC-PK1) also resulted in a significant increase in number of LLC-PK1 [co-culture (-), 0.177 0.004 vs. co-culture (+), 0.192 0.005; P < 0.01].
Figure 9.
Presence of c-met (HGF receptor) and G3PDH mRNAs in renal epithelial cells. 1 = negative control (without reverse transcriptase), 2 = rat epithelial cells (NRK-52E), 3 = porcine tubular epithelial cells (LLC-PK1), M = size marker (X174/HaeIII digest; bonds showed 1353, 1078, 872 and 603 bp from top, respectively). The rat c-met primer yields 725 bp band, and G3PDH primer purchased from Clontech (CA) yields 983 bp. This experiment was performed three times.
Full figure and legend (36K)Figure 10.
Increased epithelial cell (NRK-52E) growth by co-culture with rat mesangial cells. N = 8 per group. Abbreviations are: MC (-), endothelial cells without co-culture with mesangial cells; MC (+), endothelial cells co-cultured with mesangial cells; + control IgG, cells treated with control IgG; + HGF-Ab, ells treated with neutralizing anti-rat HGF antibody.**P < 0.01 versus MC (-).
Full figure and legend (32K)DISCUSSION
The concept of the local control of renal function by locally synthesized compounds has been recently described. It has been hypothesized that locally synthesized growth factors (such as, TGF-
, bFGF) and multiple endothelium-derived substances (PGI2, NO, CNP) also have profound influences on renal function10,11,12,13,14,15. Moreover, following mesangial injury, locally synthesized and/or down-regulated cytokines and growth factors contribute to the injury of endothelial and epithelial cells38,39,40,41. These local systems appear to be independently regulated by regional factors and may play important pathophysiologic roles. Thus, repair of epithelial and endothelial cells may have therapeutic actions in the pathogenesis of renal disease. From this viewpoint, HGF would be of interest, as HGF has many functions in the cells of target organs including the kidney6,7,8,9. Administration of rHGF prevented acute renal failure and accelerated renal regeneration in mice treated with HgCl2 and cisplatin7,9. Recent findings also show that HGF may play an important role in tissue regeneration3,4,5,6,7,8,9. Consistent with the previous findings17,32,33, our present results demonstrated that HGF can stimulate growth of epithelial and endothelial cells Figure 1, while it has no mitogenic action on mesangial cell growth. Therefore, we examined the protective action of HGF against epithelial cell injury induced by serum deprivation, which is a known mediator of apoptosis in epithelial cells34,35. Of interest, HGF could abrogate the death of epithelial cells mediated by serum deprivation Figures 1 and 2. In addition, HGF was shown to prevent apoptosis of epithelial cells induced by serum deprivation assessed by two different kinds of morphological examination and DNA fragmentation assay. HGF should be classed as a new member of the growth factors with anti-cell death actions in epithelial cells through the inhibition of apoptosis. The mechanisms by which HGF prevented epithelial cell death mediated by the conditions in this study are unclear. HGF is known to stimulate phosphatidylinositol-37-kinase (PI3K), protein tyrosine phosphatase 2, phospholipase C-r, pp60c-src, grb2/hSos1, rho and ras2,3,4,42,43,44,45. The activation of these signal transduction pathways suggests that HGF will act to prevent cell death.
On the other hand, secretion of local HGF from mesangial cells also maintains growth of endothelial cells17. Numerous papers have reported the loss of vasodilating properties of resistance vessels in chronic renal failure patients13,14,15. As previously reported33, HGF stimulated exclusively the growth of endothelial cells, but not VSMC. Therefore, we also examined whether HGF has a protective action on endothelial function in various conditions such as TNF- and dexamethasone treatment, which are known to induce apoptosis35,36,37. Interestingly, rHGF attenuated the endothelial injury induced by TNF- and dexamethasone treatment Figure 1b, suggesting that HGF in part may play a pivotal role in endothelial regulation.
What is the role of local renal HGF system in the maintenance of renal function? Mesangial cells, but not epithelial cells, secrete HGF, while epithelial cells have its specific receptor, c-met Figure 9. To examine the new concept that local production of HGF, which stimulates growth of epithelial cells but not mesangial cells, maintains growth of epithelial cells, the effects of local HGF production from mesangial cells on epithelial cell growth were studied using a co-culture system. Co-culture of mesangial cells with epithelial cells resulted in a significant increase in the number of epithelial cells, which was abolished by co-incubation with neutralizing anti-HGF antibody. Since addition of neutralizing anti-HGF antibody could not completely abolish the mitogenic action of co-culture with mesangial cells, other substances may also work to maintain the cell-cell interactions, in addition to HGF. Our previous study demonstrated a marked reduction of local HGF expression in mesangial cells by TGF- and Ang II treatment17. Taken together, down-regulation of local HGF secretion from mesangial cells by TGF- and Ang II may result in renal (epithelial and endothelial) dysfunction. Indeed, our previous report showed a significant decrease in local renal HGF level in spontaneously hypertensive rats as compared to normotensive Wistar-Kyoto rats45. The break of the paracrine-loop of HGF, which maintains epithelial and endothelial cell growth, by TGF- and/or Ang II may result in a dysfunction of the cell-cell regulation in the kidney, which in turn may accelerate the development of renal disease such as glomerulonephritis.
Overall, these results demonstrate that local production of HGF from mesangial cells may maintain epithelial and endothelial cell growth through its anti-apoptotic action. As local HGF secretion from mesangial cells was negatively regulated by Ang II and TGF-17, a decrease in local renal HGF production might be related to the dysfunction of renal cells in renal disease.
References
| 1. | Nakamura T, Hagiya M, Nishizawa T, Seki T, Shimonishi M, Sugiura A, Tashiro K & Shimizu S. Molecular cloning and expression of human hepatocyte growth factor. Nature 1989; 342: 440−443. | Article | PubMed | ISI | ChemPort | |
| 2. | Montesano R, Matsumoto K, Nakamura T & Orci L. Identification of a fibroblasts-derived epithelial morphogen as hepatocyte growth factor. Cell 1991; 67: 901−908. | Article | PubMed | ISI | ChemPort | |
| 3. | Zarnegar R & Michalpoulos GK. Many faces of hepatocyte growth factor: From hepatopoiesis to hematopoiesis. J Cell Biol 1995; 129: 1177−1180. | Article | PubMed | ISI | ChemPort | |
| 4. | Matsumoto K & Nakamura T. Hepatocyte growth factor (HGF) as tissue organizer for organogenesis and regeneration. Biochem Biophys Res Commun 1997; 239: 639−644. | Article | PubMed | ISI | ChemPort | |
| 5. | Matsumoto K & Nakamura T. Emerging multipotent aspect of hepatocyte growth factor. J Biochem 1996; 119: 591−600. | PubMed | ISI | ChemPort | |
| 6. | Igawa T, Kanda S, Kanetake H, Saitoh Y, Ichihara A, Tomita Y & Nakamura T. Hepatocyte growth factor is a potent mitogen for cultured rabbit renal tubular epithelial cells. Biochem Biophys Res Commun 1991; 174: 831−838. | Article | PubMed | ChemPort | |
| 7. | Kawaida K, Matsumoto K, Shimazu H & Nakamura T. Hepatocyte growth factor prevents acute renal failure and accelerates renal regeneration in mice. Proc Natl Acad Sci USA 1994; 91: 4357−4361. | PubMed | ChemPort | |
| 8. | Nagaike M, Hirao S, Tajima H, Noji S, Taniguchi S, Matsumoto K & Nakamura T. Renotropic functions of hepatocyte growth factor in renal regeneration after unilateral nephrectomy. J Biol Chem 1991; 266: 22781−22784. | PubMed | ISI | ChemPort | |
| 9. | Igawa T, Matsumoto K, Kanda S, Saito Y & Nakamura T. Hepatocyte growth factor may function as a renotropic factor for regeneration in rats with acute renal injury. Am J Physiol 1993; 265: F61−F69. | PubMed | ChemPort | |
| 10. | Luscher TF. The endothelium in hypertension: Bystander, target or mediator? J Hypertens 1994; 12: S105−116. | ChemPort | |
| 11. | Suga S, Nakao K, Itoh H, Komatsu Y, Ogawa Y, Hama N & Imura H. Endothelial production of C-type natriuretic peptide and its marked augmentation by transforming growth factor- . Possible existence of "vascular natriuretic peptide system.". J Clin Invest 1992; 90: 1145−1149. | PubMed | ChemPort | |
| 12. | Blantz RC, Lortie M, Vallon V, Gabbai FB, Parmer RJ & Thomson S. Activities of nitric oxide in normal physiology and uremia. Semin Nephrol 1996; 16: 144−150. | PubMed | ChemPort | |
| 13. | Lee LK, Meyer TW, Pollock AS & Lovett DH. Endothelial cell injury initiates glomerular sclerosis in the rat remnant kideny. J Clin Invest 1995; 96: 953−964. | PubMed | ChemPort | |
| 14. | Rigotti P, Ferraresso M, Morpurgo E, Cardrobbi R, Menon F, Bacelle L, Calabrese F, Valente ML, Gatta A & Ancona E. Prevention of renal ischemic injury by endothelial protecting agents. Transplant Proc 1995; 27: 749−751. | PubMed | ChemPort | |
| 15. | Yaqoob M, Patrick AW, Mcclelland P, Stevenson A, Mason H, White MC & Bell GM. Relationship between markers of endothelial dysfunction, oxidant injury and tubular damage in patients with insulin-dependent diabetes mellitus. Clin Sci (Colch) 1993; 85: 557−562. | ChemPort | |
| 16. | Kelley VR, Diaz-Gallo C, Jevnikar AM & Singer GG. Renal tubular epithelial and T cell interactions in autoimmune renal disease. Kidney Int 1993; 43 Suppl 39: S108−S115. |
| 17. | Yo Y, Morishita R, Yamamoto K, Tomita N, Kida I, Hayashi S, Moriguchi A, Kato S, Matsumoto K, Nakamura T, Higaki J & Ogihara T. Actions of hepatocyte growth factor as a local modulator in the kidney: Potential role in pathogenesis of renal disease. Kidney Int 1998; 53: 50−58. | Article | PubMed | ChemPort | |
| 18. | Morishita R, Nakamura S, Nakamura Y, Aoki M, Moriguchi A, Kida I, Yo Y, Matsumoto K, Nakamura T, Higaki J & Ogihara T. Potential role of endothelium-specific growth factor, hepatocyte growth factor, on endothelial damage in diabetes mellitus. Diabetes 1997; 46: 138−142. | PubMed | ChemPort | |
| 19. | Libby P & O'Brien KV. Culture of quiescent arterial smooth muscle cells in a defined serum-free medium. J Cell Physiol 1983; 115: 217−223. | Article | PubMed | ChemPort | |
| 20. | Ishiyama M, Shiga M, Sasamoto K, Mizoguchi M & He P. A new sulfonated tetrazolium salt that produces a highly water-soluble formazan dye. Chem Pharm Bull 1993; 41: 1118−1122. | ChemPort | |
| 21. | Pollman MJ, Yamada T, Horiuchi M & Gibbons GH. Vasoactive substances regulate vascular smooth muscle cell apoptosis; countervailing influences of nitric oxide and angiotensin II. Circ Res 1996; 79: 748−756. | PubMed | ISI | ChemPort | |
| 22. | Bennett MR & Schwartz SM. Apoptosis of human vascular smooth muscle cells derived from normal vessels and coronary atherosclerotic plaques. J Clin Invest 1995; 95: 2266−2274. | PubMed | ISI | ChemPort | |
| 23. | Araki S, Shimada Y, Kaji K & Hayashi H. Apoptosis of vascular endothelial cells by fibroblast growth factor deprivation. Biochem Biophys Res Commun 1990; 168: 1194−1200. | Article | PubMed | ChemPort | |
| 24. | Aoki M, Morishita R, Matsushita H, Nakano N, Hayashi S, Tomita N, Yamamoto K, Moriguchi A, Higaki J & Ogihara T. Serum deprivation induced apoptosis accompanied by up-regulation of p53 and bax in human aortic vascular smooth muscle cells. Heart Vessels 1997; S12: 71−75. |
| 25. | Ito M, Watanabe M, Ihara T, Kamiya H & Sakurai M. Fas antigen and bcl-2 expression on lymphocytes cultured with cytomegalovirus and varicella-zoster virus antigen. Cell Immunol 1995; 160: 173−177. | Article | PubMed | ChemPort | |
| 26. | Tashiro K, Hagiya M, Nishizawa T, Seki T, Shimonishi M, Shimizu S & Nakamura T. Deduced primary structure of rat hepatocyte growth factor and expression of the mRNA in rat tissues. Proc Natl Acad Sci USA 1990; 87: 3200−3204. | PubMed | ChemPort | |
| 27. | Hayashi S, Morishita R, Higaki J, Aoki M, Moriguchi A, Kida I, Sawa Y, Matsumoto K, Nakamura T, Kaneda Y & Ogihara T. Autocrine-paracrine effects of over-expression of hepatocyte growth factor gene on growth of endothelial cells. Biochem Biophys Res Commun 1996; 220: 539−545. | Article | PubMed | ISI | ChemPort | |
| 28. | Morishita R, Gibbons GH, Pratt RE, Tomita N, Kaneda Y, Ogihara T & Dzau VJ. Autocrine and paracrine effects of atrial natriuretic peptide gene transfer on vascular smooth muscle and endothelial cellular growth. J Clin Invest 1994; 94: 824−829. | PubMed | ISI | ChemPort | |
| 29. | Kaneda Y, Iwai K & Uchida T. Increased expression of DNA cointroduced with nuclear protein in adult rat liver. Science 1989; 243: 375−378. | PubMed | ISI | ChemPort | |
| 30. | Yamada A, Matsumoto K, Iwanari H, Sekiguchi K, Kawata S, Matsuzawa Y & Nakamura T. Rapid and sensitive enzyme-linked immunosorbent assay for measurement of HGF in rat and human tissues. Biomed Res 1995; 16: 105−114. | ChemPort | |
| 31. | Nakamura Y, Morishita R, Higaki J, Kida I, Aoki M, Moriguchi A, Yamada K, Hayashi S, Yo Y, Matsumoto K & Nakamura T. Expression of local hepatocyte growth factor system in vascular tissues. Biochem Biophys Res Commun 1995; 215: 483−488. | Article | PubMed | ChemPort | |
| 32. | Nakamura Y, Morishita R, Nakamura S, Aoki M, Moriguchi A, Matsumoto K, Nakamura T, Higaki J & Ogihara T. A vascular modulator, hepatocyte growth factor, is associated with systolic pressure. Hypertension 1996; 28: 409−413. | PubMed | ChemPort | |
| 33. | Nakamura Y, Morishita R, Higaki J, Kida I, Aoki M, Moriguchi A, Yamada K, Hayashi S, Yo Y, Nakano H, Matsumoto K, Nakamura T & Ogihara T. Hepatocyte growth factor (HGF) is a novel member of endothelium-specific growth factors: Additive stimulatory effect of HGF with basic fibroblast growth factor, but not vascular endothelial growth factor. J Hypertens 1996; 14: 1067−1072. | PubMed | ISI | ChemPort | |
| 34. | Morishita R, Higaki J, Hayashi S, Yo Y, Aoki M, Nakamura S, Moriguchi A, Matsushita H, Matsumoto K, Nakamura T & Ogihara T. Role of hepatocyte growth factor in endothelial regulation: Prevention of high D-glucose-induced endothelial cell death by prostaglandins and phosphodiesterase type 3 inhibitor. Diabetologia 1997; 40: 1053−1061. | Article | PubMed | ChemPort | |
| 35. | Cotran RS & Pober JS. Effects of cytokines on vascular endothelium: Their role in vascular and immune injury. Kidney Int 1989; 35: 969−975. | PubMed | ChemPort | |
| 36. | Libby P. Do vascular wall cytokines promote atherogenesis? Hosp Pract 1992; 27: 51−58. | ChemPort | |
| 37. | Lee SW, Tsou AP, Chan H, Thomas J, Petrie K, Eugui EM & Allison G. Glucocorticoids selectively inhibit the transcription of the interleukin 1 gene and decrease the stability of interleukin 1 mRNA. Proc Natl Acad Sci USA 1988; 85: 1204−1208. | PubMed | ChemPort | |
| 38. | Sakai H, Naka R, Suzuki D, Nomoto Y, Miyazaki M, Nikolic-Paterson DJ & Atkins RC. In situ hybridization analysis of TGF-beta in glomeruli from patients with IgA nephropathy. Contrib Nephrol 1995; 111: 107−114. | PubMed | ChemPort | |
| 39. | Border WA. Transforming growth factor-beta and the pathogenesis of glomerular diseases. Curr Opin Nephrol Hypertens 1994; 3: 54−58. | PubMed | ChemPort | |
| 40. | Floege L, Eng E, Yopung BA & Johenson RJ. Factors involved in the regulation of mesangial cell proliferation in vitro and in vivo. Kidney Int 1993; 39: S47−54. |
| 41. | Harris RC & Martinez-Maldonado M. Angiotensin II-mediated renal injury. Miner Electrolyte Metab 1995; 21: 328−335. | PubMed | ChemPort | |
| 42. | Canley LG & Cantley LC. Signal transduction by the hepatocyte growth factor receptor, c-met. Activation of the phosphatidylinositol 3-kinase. J Am Soc Nephrol 1995; 5: 1872−1881. | PubMed | |
| 43. | Ridley AJ, Comoglio PM & Hall A. Regulation of scatter factor/hepatocyte growth factor responses by ras, rac and rho in MDCK cells. Mol Cell Biol 1995; 15: 1111−1122. |
| 44. | Takaishi K, Sasaki T, Kato M, Yamoshi W, Kuroda S, Nakamura T, Takeuchi M & Takai Y. Involvement of Rho p21 small GTP-binding protein and its regulation in HGF-induced cell motility. Oncogene 1994; 9: 273−279. | PubMed | ISI | ChemPort | |
| 45. | Nakano N, Moriguchi A, Morishita R, Kida I, Tomita N, Matsumoto K, Nakamura T, Higaki J & Ogihara T. Role of angiotensin II in the regulation of a novel vascular modulator, hepatocyte growth factor, in experimental hypertensive rats. Hypertension 1997; 30: 1448−1454. | PubMed | ChemPort | |
Acknowledgments
Dr. Ryuichi Morishita is the recipient of a Harry Goldblatt Award from the Council of High Blood Pressure, the American Heart Association. Dr. Shin-ichiro Hayashi is a Research Fellow of the Japan Society for the Promotion of Science. This work was partially supported by grants from the Japan Society for the Promotion of Science, the Japan Heart Foundation: Pfizer Pharmaceuticals Grant for Research on Coronary Artery Disease and Sagawa Cancer Research Foundation. The authors thank Ms. Chihiro Noguchi for excellent technical assistance.
