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Am J Physiol Renal Physiol 287: F1223-F1232, 2004. First published August 17, 2004; doi:10.1152/ajprenal.00245.2004
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Hypoxic induction of Ctgf is directly mediated by Hif-1

Debra F. Higgins,1 Mangatt P. Biju,1 Yasuhiro Akai,1 Anton Wutz,2 Randall S. Johnson,3 and Volker H. Haase1

1Department of Medicine, University of Pennsylvania, Philadelphia, Pennsylvania 19104-6144; 2Research Institute of Molecular Biology, A-1030 Vienna, Austria; and 3Molecular Biology Section, Division of Biology, University of California San Diego, La Jolla, California 92093-0366

Submitted 1 July 2004 ; accepted in final form 12 August 2004


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
CTGF plays a significant role in the development of renal fibrosis by mediating the fibrotic effects of transforming growth factor (TGF)-{beta}1 and has been shown to be hypoxia inducible in human breast cancer cells. It has been suggested that hypoxia is an important underlying cause for the development of renal fibrosis through the modulation of profibrotic genes. One of the key mediators of the cell's response to lowered oxygen environments is hypoxia-inducible-factor-1 (HIF-1), a basic helix-loop-helix transcription factor, which enables cells to adapt to hypoxia by regulating the expression of genes involved in increasing oxygen availability (VEGF, erythropoietin) and enhancing glucose uptake and metabolism (Glut-1, PGK). In this paper, we have used primary tubular epithelial cell cultures from a tetracycline-inducible-Hif-1{alpha} knockout murine model to further elucidate the role of Hif-1 in the hypoxic-induction of Ctgf expression. We show that hypoxia response elements present upstream of Ctgf enable direct interaction of Hif-1 transcription factor with the Ctgf promoter, resulting in increased transcription of Ctgf mRNA. Cells deficient in Hif-1{alpha} were incapable of inducing Ctgf mRNA in response to hypoxia, suggesting an absolute requirement of Hif-1. Furthermore, the observed Hif-1-mediated hypoxic stimulation of Ctgf expression was found to occur independently of TGF-{beta}1 signaling. Our findings have important implications for a number of fibrotic disorders in which hypoxia, CTGF, and TGF-{beta}1 are involved, including renal, dermal, hepatic, and pulmonary fibrosis.

renal fibrosis; transforming growth factor-{beta}; hypoxia; hypoxia response elements

END-STAGE RENAL DISEASE (ESRD) is typified by significant tubulointerstitial fibrosis irrespective of the initiating insult. Progression to ESRD and loss of renal function correlate with the degree of tubulointerstitial fibrosis (1, 6), which is characterized by the loss of renal tubular epithelial cell integrity, loss of peritubular capillaries, and a significant increase in myofibroblasts in the interstitium, which contribute to the increased deposition of extracellular matrix molecules. A significant proportion of myofibroblasts have been shown to derive from the transition of tubular epithelial cells toward a myofibroblast phenotype by a process known as epithelial-mesenchymal transition (EMT), highlighting the importance of tubular epithelial cells in the pathogenesis of renal fibrosis (33). We recently identified upregulation of Ctgf (also known as fisp-12, ccn-2) in vivo during unilateral ureteral obstruction, a model of tubulointerstitial fibrosis, and also in tubular epithelial cells undergoing EMT in response to transforming growth factor (TGF)-{beta}1 in vitro (24). Expression of CTGF has been positively correlated with the degree of fibrosis, and indeed CTGF protein excretion in urine samples from patients with kidney disease has been identified as an effective indicator of renal function, with higher levels of CTGF protein reflecting increased renal damage (19, 49).

CTGF is a matricellular protein belonging to the CCN family (reviewed in Ref. 2) and is involved in development and cellular differentiation. Expression of CTGF is upregulated during wound repair (28), inflammation (7), fibrotic disorders (24, 27, 29, 37), tumor growth (16, 61, 69, 71), and angiogenesis (31, 36, 59). Biological properties of CTGF include involvement in cell proliferation, adhesion, chemotaxis, and extracellular matrix production in a variety of cell types such as fibroblasts, mesangial cells, endothelial cells, and chondrocytes (15, 43, 60). Since CTGF appears to be a critical factor in the development of tubulointerstitial fibrosis, acting as a downstream mediator of TGF-{beta}1 signaling (20), and is differentially regulated during EMT (17, 18, 24, 37), understanding the mechanisms controlling CTGF expression is of paramount importance in defining the role of this molecule in renal disease.

Since microvascular changes are observed during renal fibrosis, evidence is now accumulating which suggests that localized hypoxia may contribute to the development of fibrotic regions within the kidney (10, 12, 44). Cells respond to low oxygen levels by stabilizing the transcription factor hypoxia-inducible factor-1 (HIF-1), which leads to upregulation of genes involved in vasculogenesis (VEGF), erythropoiesis (EPO), glucose metabolism (PGK, LDH), glucose uptake (Glut-1) (42), and fibrogenesis [matrix metalloproteinase (MMP)-1, tissue inhibitor of MMP (TIMP)-3, plasminogen activator inhibitor (PAI)-1] (35, 44, 46). HIF-1 is a basic helix-loop-helix transcription factor that is composed of an oxygen-sensitive {alpha}-subunit, HIF-1{alpha}, and a constitutively expressed {beta}-subunit, HIF-1{beta} [also known as aryl hydrocarbon receptor nuclear translocator (ARNT)] (67). HIF-1{alpha} protein stability is tightly regulated in an oxygen-dependent manner; in the presence of oxygen, HIF-1{alpha} is hydroxylated by prolyl hydroxylase enzymes (9) enabling interaction with pVHL (26, 30, 34). pVHL is the substrate recognition component of an E3-ubiquitin ligase complex (32) that targets HIF-1{alpha} for degradation by the 26S proteasome (32, 39). In the absence of oxygen, prolyl hydroxylase activity is inhibited, which results in the stabilization and nuclear translocation of the HIF-1{alpha}-subunit, enabling it to bind to HIF-1{beta} and form transcriptionally active HIF-1.

HIF-1 regulates gene expression through interaction with sequence-specific hypoxia response elements (HREs) present in either upstream, downstream, or intronic regions of hypoxia-responsive genes. The HRE motif was first identified as a 50-bp sequence in the 3'-flanking region of the human erythropoietin (EPO) gene (57). Similar regulatory elements were found in the 5'-upstream sequences of the VEGF gene (14, 38). Subsequent identification of HREs in other hypoxia-responsive genes revealed that the pentanucleotide 5'-RCGTG-3' was sufficient for binding of HIF-1 (56).

CTGF mRNA and protein levels have been shown to increase in response to hypoxia in the human breast cancer cell line MDA231 (59), although HREs were not observed in the 800 bases located immediately upstream of CTGF or in sequences downstream of the coding region, suggesting that the observed hypoxic regulation was not mediated through HIF (36). However, we observed a significant increase in Ctgf mRNA expression by oligonucleotide microarray analysis of murine kidneys with constitutively active Hif-1 [achieved through inactivation of pVhl (21), data not presented here], which suggested that Ctgf may be regulated by a Hif-1-dependent pathway. Hif-1 regulation of Ctgf was further supported by analysis of murine strains in which both Vhl and Hif-1{alpha} were deleted (Haase VH, unpublished observations), and similar regulation was observed in other cell types, for example, in cerebral cortex and in thymocytes (Haase VH, unpublished observations), suggesting a general role for Hif-1 in the regulation of Ctgf mRNA expression rather than a cell-specific mechanism. To further define the role of Hif-1 in the regulation of Ctgf expression during hypoxia, we analyzed Ctgf expression in primary tubular epithelial cells (PTCs) which were either wild-type or mutated for Hif-1{alpha}.

In this report, we describe our findings on the hypoxic regulation of Ctgf gene expression and the specific involvement of Hif-1 in PTCs. In particular, we show that Hif-1 can directly interact with HREs located in the 5'-region of Ctgf. We show, by use of a transgenic murine model, that Hif-1 is unequivocally required for hypoxic induction of Ctgf in primary renal epithelial cells and describe the absolute requirement of two HRE motifs for increased promoter activity in response to hypoxia. Furthermore, we show that TGF-{beta}1 signaling is not required for hypoxic induction of Ctgf expression. Taken together, our results provide compelling evidence that Hif-1 is directly involved in the regulation of Ctgf at the transcriptional level.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Generation of tetracycline-inducible Cre recombinase Hif-1{alpha} 2-lox mice. Mice were generated by breeding tetracycline-controllable Lc-1 Cre-transgenic mice (55) to mice homozygous for the Hif-1{alpha} 2-lox allele (52) and that carried the reverse transactivator 2 cDNA (rtTA2) under the control of the ubiquitously expressed ROSA26 promoter (70).

Cell culture. Primary tubular epithelial cells were isolated from kidney cortex of 3-wk-old mice (Hif-1{alpha}2-lox/2-lox; R26-rtTA2; Lc-1) as previously described (64, 65) and grown in DMEM/Ham's F-12 (Invitrogen, Carlsbad, CA) supplemented with 50 ng/ml insulin, 200 ng/ml hydrocortisone, 5 µg/ml apotransferrin (Sigma, St. Louis, MO), 1% penicillin, and 1% streptomycin. Primary cells were characterized by immunocytochemistry for anti-E-cadherin (Transduction Laboratories, Lexington, KY), anti-cytokeratin (Sigma), and anti-{alpha}-smooth muscle actin (Neomarkers, Lab Vision, Fremont, CA) according to standard procedures. For Hif-1{alpha} exon 2 deletion, primary cells were treated with 4 µg/ml doxycycline 1 day after isolation, and this treatment was continued for 4 days; doxycycline was withdrawn at least 24 h before cell stimulation. MCT (23) and LLC-PK (ATCC) cells were grown in DMEM/Ham's F-12 medium (Invitrogen) supplemented with 10% fetal calf serum, 1% penicillin, and 1% streptomycin. Cells were subjected to hypoxic treatment of 0.5% O2-5% CO2 in an Invivo2 hypoxia chamber (Ruskinn Technologies, Leeds, UK) or cultured under normoxic conditions of 21% O2-5% CO2. Deferoxamine mesylate (DFX; 100 µM, Sigma) was added to cells for at least 6 h to induce HIF-1{alpha} stabilization.

Genotyping for Hif-1{alpha}. Genomic DNA was isolated from primary cell cultures using the TRIzol LS reagent (Invitrogen) after the initial isolation of RNA according to the manufacturer's protocol. Hif-1{alpha} 1-lox and 2-lox alleles were detected with the following primers: forward 1: 5'-TTG GGG ATG AAA ACA TCT GC-3'; forward 2: 5'-GCA GTT AAG AGC ACT AGT TG-3'; and reverse: 5'-GGA GCT ATC TCT CTA GAC C-3'. The 2-lox allele was identified as a ~260-bp band and 1-lox allele as a ~270-bp band.

RNA isolation and analysis. Total RNA was isolated from cell cultures using the TRIzol LS reagent (Invitrogen) as per the manufacturer's instructions. Fifteen micrograms of RNA were run on a 1.2% formaldehyde gel, blotted to GeneScreen Plus Hybridization Membrane (New England Nuclear Life Science Products, Boston, MA), and probed with radiolabeled probes for Pgk and Ctgf. Probes were prepared by random primer labeling, using a Prime It II Kit (Stratagene, Cedar Creek, TX), of PCR-amplified products from a cDNA template using specific primers: PGK forward: 5'-CAA ACA ACC AAA GGA TCA AGG-3'; PGK reverse: 5'-CCC AAG ATA GCC AGG AAG G-3'; CTGF forward: 5'-CTA AGA CCT GTG GAA TGG GC-3'; and CTGF reverse: 5'-CTC AAA GAT GTC ATT GTC CCC-3'. Membranes were hybridized for 1 h at 65°C in Quick Hyb Buffer (Stratagene), washed, and exposed to Biomax film (Kodak). Densitometric analysis of autoradiographs was performed using Scion Image (Release Beta 4.0.2, Scion). RNase protection analysis (RPA) was performed using a RiboQuant Ribonuclease Protection Assay kit (Pharmingen) with an mAng-1 multiprobe template set.

EMSA. Nuclear and cytoplasmic protein isolation and EMSA were performed using Epo-HRE as described previously (3). Epo-HRE, HRE 5.1, HRE 5.2, {Delta}HRE 5.1, and {Delta}HRE 5.2 were generated as single-strand templates (Sigma Genosys).

Western blot analysis. For detection of Hif-1{alpha} and Hif-2{alpha}, nuclear extracts were prepared as described previously (3). Fifteen micrograms of nuclear protein extract were size separated by 3–8% gradient SDS-PAGE (Invitrogen) and electrotransferred to nitrocellulose membranes (Amersham Pharmacia Biotech, Piscataway, NJ). Equivalent loading was assessed by staining with Ponceau S. Membranes were blocked with 5% nonfat dry milk in 0.1% Tween 20/Tris-buffered saline for 20 min and incubated with the primary rabbit anti-murine Hif-1{alpha} antiserum (Cayman Laboratories) or rabbit polyclonal anti-Hif-2{alpha} antiserum (generated against murine Hif-2{alpha} peptide 580–693) overnight at 4°C. Blots were washed, incubated with horseradish peroxidase-conjugated secondary antibody at room temperature for 1 h, washed, and developed with ECL Plus Reagent (Amersham Pharmacia Biotech) according to the manufacturer's instructions.

Generation of reporter luciferase constructs. A 3.8-kb fragment upstream of Ctgf containing a full-length promoter region and part of exon 1 (from –14 to –3873 relative to the transcription start site) was PCR-amplified from murine genomic DNA using ctgfgen forward: 5'-ACG CGT CGA CGA AGC CAG TTC CCC AAG G-3' and ctgfgen reverse: 5'-ACG CGT CGA CAG GAG GAT GCA CAG CAG G-5' (both of which incorporate a SalI restriction site) and cloned into TOPO XL plasmid (Invitrogen). Identity of inserted DNA was confirmed by sequencing. Luciferase promoter constructs were generated by inserting a 3.8-kb HinDIII/EcoRV fragment, containing both HRE 5.1 and HRE 5.2, into XhoI-blunt/HinDIII-pGL3 basic plasmid (Promega, Madison, WI). The 1.0-kb construct was generated by inserting a HinDIII/XbaI fragment, containing neither HRE, into HinDIII/NheI-pGL3 basic plasmid. The 1.7-kb constructs, containing HRE 5.2, were generated by PCR amplification from the 3.8-kb pGL3 construct using a forward primer, 5'-ATT TGT CTC GAG AAA CAC CAA CTA TGA GAG G-3', which contained a XhoI restriction site, and a reverse primer, 5'-TCA TAG AAT TAT GCA GTT GC-3', which recognized part of the pGL3 basic plasmid. The PCR product was digested with XhoI/HinDIII and ligated into pGL3 basic. Mutation of the HRE sites and SMAD binding element was performed using a Site Directed Mutagenesis Kit (Stratagene). Primers designed for mutagenesis were {Delta}HRE 5.1 forward: 5'-TGG GTT TTG GCT TTG AGT Cga aTT CCA CCA GTA TGT TTC CC-3'; {Delta}HRE 5.1 reverse: 5'-GGG AAA CAT ACT GGT GGA Att cGA CTC AAA GCC AAA ACC C-3'; {Delta}HRE 5.2 forward: 5'-ACA CAC ACA CAC ACA CAC ACA aAa GCA AAG AGA GAC AGA GAG-3'; {Delta}HRE 5.2 reverse: 5'-CTC TCT GTC TCT CTT TGC tTt TGT GTG TGT GTG TGT GTG TG-3'; {Delta}SMAD forward: 5'-GAG CTA AAG TGT GCC AGC TTT Tgg Atc CGG AGG AAT GTG GAG TGT C-3'; and {Delta}SMAD reverse: 5'-GAC ACT CCA CAT TCC TCC Gga TcC AAA AGC TGG CAC ACT TTA GCT C-3'. HRE and SMAD elements are shown underlined and in bold; mutated nucleotides are shown in lowercase.

Luciferase assays. For luciferase assays, 20 x 104 MCT or LLC-PK cells were inoculated onto 24-well plates (Costar, Corning, NY) and transiently transfected with 0.7 µg of each pGL3 basic construct along with 0.1 µg Renilla luciferase plasmid (Promega) per well using a 3:1 ratio of Lipofectamine 2000:DNA in OptiMEM medium (Invitrogen). Cells were transfected for 4–6 h, and the medium was replaced with serum-free DMEM/Ham's F-12 with antibiotics. Cells were grown overnight and stimulated with 0.5% O2 for 12 h or 5 ng/ml TGF-{beta}1 (Sigma) for 6 h. Cell lysates were obtained by scraping cells into passive lysis buffer and were analyzed for firefly and Renilla luciferase activity using the Dual Luciferase Reporter Assay System (Promega). Luminescence was detected using a Microplate Luminometer and Simplicity version 2.1 software (Berthold Detection Systems). Transfections were performed in triplicate, and all experiments were repeated at least three times. Firefly activity was normalized to Renilla activity to compensate for differences in transfection efficiency, and results are displayed as relative luciferase activity ± SD. Statistical analysis was performed using Student's t-test. A P value <0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Ctgf mRNA is regulated by hypoxia in PTCs. Oligonucleotide microarray analysis of kidneys in which Hif-1 was stabilized revealed an increase in Ctgf mRNA expression (Haase VH, unpublished observations), highlighting the possible regulation of this gene by Hif-1. To assess the contribution of hypoxia to Ctgf mRNA levels, PTCs were isolated, grown to 80% confluency, and subjected to 0.5% O2 for 1–24 h or kept at normoxia for 24 h. PTC cultures were characterized by immunocytochemistry and were found to stain positive for the epithelial markers E-cadherin and cytokeratin and negative for the mesenchymal marker {alpha}-smooth muscle actin (data not shown). The hypoxic response was determined by monitoring the gene expression of known hypoxia-regulated genes such as Pgk (Fig. 1A). Ctgf mRNA levels were increased 2.64 ± 0.51-fold (P < 0.005) after 1 h at 0.5% O2 and remained elevated up to 24 h (Fig. 1).



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  		Fig. 1. Time course of Ctgf mRNA expression in primary renal tubular epithelial cells (PTCs) in response to hypoxia. A: PTCs were isolated, cultured until 80% confluent, and exposed to 0.5% O2 for 0–24 h. Ctgf and Pgk transcripts were detected by Northern blot analysis; a representative blot is shown along with ethidium bromide-stained 28S and 18S rRNA for loading. Results were repeated at least 3 times. Densitometric analysis for Ctgf mRNA levels are shown normalized to 18S rRNA level; 0 h was given an arbitrary value of 1. Values are means ± SD of samples from 3 Northern blots. *P < 0.01 and **P < 0.05 compared with 0-h sample.

 
Conditional targeting of Hif-1{alpha} in PTCs. To further characterize the role of the transcription factor Hif-1 in the hypoxic induction of Ctgf mRNA, murine strains in which Hif-1 could be conditionally inactivated by Cre-loxP-mediated recombination were used for the isolation of PTCs. Mice containing a tetracycline-inducible Cre recombinase (Lc-1) under the control of the tet-operon and containing reverse transactivator 2 cDNA (rtTA2) expressed from the ROSA26 promoter were crossed to murine strains homozygous for the Hif-1{alpha} 2-lox allele (Fig. 2A). PTCs were isolated and for some experiments cultured in the presence of 4 µg/ml doxycycline for approximately 4 days, which resulted in the activation of Cre recombinase through binding of the reverse transactivator to the tet-operon, and the subsequent conversion of the Hif-1{alpha} 2-lox allele to the nonfunctional 1-lox allele. In the Hif-1{alpha} 2-lox allele, exon 2, which encodes the basic helix-loop-helix domain required for dimerization and DNA binding, is flanked by two loxP sites (floxed), which enable deletion of this region and generation of an out-of-frame mutation. The efficiency of recombination of the alleles was determined by PCR (Fig. 2B). In cells treated with doxycycline, 2-lox alleles were not detectable by PCR, suggesting complete deletion of the floxed exon 2 region. Western blot analysis of nuclear extracts from wild-type cells revealed a low level of Hif-1{alpha} stabilization under normoxic conditions, which was greatly enhanced on exposure to hypoxic conditions of 0.5% O2 for 6 h (Fig. 2C). Normoxic stabilization of Hif-1{alpha} has been reported previously and has been correlated with cell culture confluency (58). Given the nature of epithelial growth displaying high cell-to-cell contact, we feel that this normoxic level of Hif-1{alpha} is representative of normal epithelial cell cultures. In doxycycline-treated PTCs, Hif-1{alpha} was undetectable in both normoxic and hypoxic nuclear extracts, confirming complete absence of Hif-1{alpha} protein (Fig. 2C). Hif-2{alpha}, a homolog of Hif-1{alpha}, can also bind the Hif-1{beta} subunit to form functional Hif-2 transcription factor. Western blot analysis for Hif-2{alpha} revealed the presence of a nonspecific band (Fig. 2C, nonspecific) with a slightly higher electrophoretic mobility than that in the positive control lane [Fig. 2C, lane D, DFX-treated distal tubular cells (48); DFX is a hypoxia mimetic that results in the stabilization of Hif-1{alpha} through the inactivation of prolyl hydroxylase activity (34, 68)]. We conclude from this that Hif-2{alpha} was not significantly expressed in PTCs and that deletion of Hif-1{alpha} did not result in compensatory expression of Hif-2{alpha}. Furthermore, RPA analysis revealed increased expression of Vegf under hypoxic conditions in wild-type cells only (Fig. 2D). Since Vegf is a known Hif-2 target gene, the absence of its induction in hypoxic Hif-1{alpha}-null cells suggests that PTCs do not express Hif-2{alpha} protein at a functional level and that Hif-1 is the main hypoxia-inducible-factor produced in these cells. Vegf expression was slightly higher in wild-type normoxic cells compared with normoxic Hif-1{alpha}-null cells, supporting our observation of a baseline stabilization of Hif-1 under normoxia.



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  		Fig. 2. Efficient generation of hypoxia-inducible-factor-1 (Hif-1{alpha})-deficient primary PTCs by tetracycline-inducible Cre recombinase-mediated deletion. A: mice expressing reverse-transactivator 2 (rtTA2) under the control of the ROSA26 promoter and containing Lc-1 Cre recombinase (Lc-1 Cre) under the control of the tet-operon (tetO) were crossed with mice containing the Hif-1{alpha} 2-lox allele (HIF-1 ex2, loxP). In the presence of doxycycline (+DOX), the reverse transactivator ({bullet} and {circ}) binds the tet-operon and induces Cre recombinase expression, which mediates the deletion of Hif-1{alpha} exon 2 between the loxP sites. As Hif-1{alpha} exon 2 encodes the basic helix-loop-helix domain required for dimerization to Hif-1{beta}, stable Hif-1 transcription factor is not formed. B: DNA was isolated from PTCs cultured in the absence (–) or presence (+) of 4 µg/ml DOX as described in EXPERIMENTAL PROCEDURES. Genotyping for wild-type (2-lox) or deleted (1-lox) Hif-1{alpha} exon 2 was performed, and aliquots of the PCR reaction were visualized on 3% agarose gels along with known positive genomic tail DNA controls for heterozygous 1-lox/2-lox (1/2) or homozygous 2-lox mice (2/2). C: Western blot analysis for Hif-1{alpha} and Hif-2{alpha} in wild-type and DOX-treated PTCs. N, normoxia; H, hypoxia; D, positive control from DFX-treated DT cells; n.s., nonspecific. D: RPA analysis for Vegf mRNA in wild-type or DOX-treated PTCs. Note that hypoxic induction of Vegf only occurs in cells with wild-type Hif-1{alpha}. L32 levels are shown as loading controls.

 
Functional Hif-1 is required for hypoxic induction of Ctgf mRNA. PTCs, wild-type or deficient for Hif-1{alpha}, were exposed to 0.5% O2 for 0–24 h. PCR genotyping of DNA from each treatment confirmed the conversion of the 2-lox allele in wild-type cells to a 1-lox allele in doxycycline-treated cells (Fig. 2B). mRNA levels of the hypoxia-responsive gene Pgk were assessed to monitor the functional efficiency of Hif-1{alpha} gene deletion. In Hif-1{alpha}-deficient cells, hypoxic-induction of Pgk mRNA was no longer detectable, confirming the absence of functional Hif-1 (Fig. 3A comparing wild-type with HIF–/– lanes). Ctgf mRNA displayed a time-dependent hypoxic induction in wild-type cells; however, this induction was not paralleled in Hif-1{alpha}-deficient cells, as assessed by Northern blot (Fig. 3A) and densitometric analysis (Fig. 3B). These results suggest that Hif-1 is either directly or indirectly involved in the hypoxic induction of Ctgf mRNA.



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  		Fig. 3. Hif-1 is required for hypoxic induction of Ctgf mRNA. A: wild-type or Hif-1{alpha}-deficient PTCs (Hif –/–, generated by exposure to DOX, which was removed at least 24 h before hypoxic stimulation) were subjected to 0.5% O2 for 0–24 h. Ctgf and Pgk mRNA levels were assessed by Northern blot analysis. Ethidium bromide-stained 28S and 18S rRNA are shown for loading. A representative blot of 3 independent experiments is shown. B: densitometric analysis of Ctgf mRNA levels normalized to 18S rRNA levels; 0 h was given an arbitrary value of 1. Values are means ± SD for samples from 3 Northern blots. {blacktriangleup}, Wild-type; gray filled squares, Hif-1{alpha} –/–. *P < 0.05 compared with 0-h sample for Hif-1{alpha} wild-type samples.

 
Functional HREs present in 5'-regulatory region of Ctgf. To examine whether Hif-1 is able to interact directly with the Ctgf promoter, we examined regions of genomic DNA flanking Ctgf for the presence of specific HREs. Previous reports have described a number of transcription factor binding elements contained within the 800-base promoter region of CTGF, including TATA box, Sp1 motif, basal control element-1 (BCE-1; required for the basal constitutive expression of CTGF) (2); however, HREs were not found in this region (36). Since our data suggested that Hif-1 is required for the hypoxic induction of Ctgf, we searched a larger region of ~4,000 bases of an upstream genomic sequence and ~1,100 bases downstream for HRE motifs. We specifically looked for the core HRE sequence 5'-RCGTG-3', which is critical for Hif-1 binding, using MatInspector software (Genomatix, Munich, Germany). We identified two potential HRE sites, which we designated HRE 5.1, located at –3741 to –3745, and HRE 5.2, located at –1554 to –1558, relevant to the transcription start site, which was designated +1 (Fig. 4A). The core HRE sequence was not detected in the downstream region. To assess whether HRE 5.1 and HRE 5.2 could represent functional HRE sites, we performed EMSAs using radiolabeled probes containing the individual putative HREs (Fig. 4B). Mutated EMSA probes were designed in which the 5'-ACGTG-3' motif of HRE 5.1 was replaced with 5'-AttcG-3', introducing an EcoR1 restriction site ({Delta}HRE 5.1). The 5'-GCGTG-3' motif of HRE 5.2 was mutated to 5'-GCtTt-3' ({Delta}HRE 5.2, Fig. 4B, lower case represents the nucleotides targeted for mutation).



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  		Fig. 4. Hypoxia-response elements (HREs) present upstream of Ctgf can bind Hif-1. Hif-1 transcription factor binds the HRE denoted by the core sequence 5'-RCGTG-3'. This motif was identified twice in genomic DNA sequence spanning 4,000 bases upstream of the transcription start site for Ctgf using the MatInspector program (Genomatix Software). HRE 5.1 was located at –3741 to –3745, and HRE 5.2 was located at –1554 to –1558 (nos. refer to base no. upstream of transcription start site designated +1). Shown are depictions of previously characterized transcription factor binding sites for TATA box, basal control element-1 [BCE-1; previously termed transforming growth factor (TGF)-{beta} response element (TGF-{beta}RE)] and SMAD binding element in the 5'-upstream genomic sequence of Ctgf. Also shown are the 2 putative HREs showing the core motif 5'-RCGTG-3' on the complementary strand (arrows show direction of the motif). B: DNA probes designed for EMSA showing the HRE binding motif underlined and in bold. Mutated probes ({Delta}HRE 5.1 and {Delta}HRE 5.2) were also generated to confirm the specificity of the Hif-1 interaction with the HRE motif; mutated nucleotides in the HRE are shown in lower case. {blacktriangleup}, Mutation in HRE 5.1 that introduced a novel EcoRI restriction site. C: EMSA of putative HRE elements in the upstream sequence of Ctgf showing their ability to bind Hif-1 in vitro. Nuclear extracts from MCT cells (cortical tubular epithelial cell line) treated with or without the hypoxic mimetic deferoxamine mesylate (DFX) were incubated with radiolabeled DNA probes containing either the HRE for the erythropoietin gene (Epo-HRE), HRE 5.1, or HRE 5.2. Left: EMSA using Epo-HRE as a positive control for Hif-1 supershift. Right: EMSA with wild-type and mutated probes (4 panels). Ap-1 (A) or cold {Delta}HRE 5.2 probe ({Delta}5.2) was added to show specificity of interaction between the nuclear extracts and the DNA probes. Hif-1{alpha} antibody was used to supershift the nuclear protein-DNA probe complex and confirm Hif-1 as the nuclear factor which binds the HRE sites.

 
Nuclear extracts were obtained from the murine cortical tubular epithelial cell line MCT treated with the hypoxia mimetic DFX or under control normoxic conditions. EMSA was performed with DFX-treated nuclear extracts rather than hypoxia-treated extracts to maximize detection of Hif binding to our EMSA probes, as DFX specifically results in accumulation of the Hif-{alpha} subunit. The HRE from the erythropoietin gene (Epo-HRE) was used in a parallel EMSA as a positive control. Interaction with nuclear protein(s) was detected for the Epo-HRE, HRE 5.1, and HRE 5.2 probes (Fig. 4C). Hif-1 binding was determined by the addition of HIF-1{alpha} antibody, which resulted in the supershift of the specific Hif-1 protein-DNA complex (Fig. 4C; arrows indicate specific HIF-binding and supershift). The specificity of protein/DNA binding was shown by competition with either AP-1 for the Epo-HRE and HRE 5.1 probe or cold-mutated probe for HRE 5.2. Hif-1/DNA binding was increased in response to DFX treatment compared with control, and mutation of the putative HRE sites in {Delta}HRE 5.1 and {Delta}HRE 5.2 resulted in loss of the specific Hif-1/DNA interaction, showing the absolute requirement of the 5'-RCGTG-3' motif. Nuclear binding was observed for the {Delta}HRE 5.2 probe, which displayed a similar electrophoretic mobility to that of the Hif-1/DNA interaction; however, this could not be supershifted with Hif-1{alpha} antibody, suggesting interaction with an as yet unidentified nuclear protein. Specific Hif-1 binding of all three DNA probes in normoxic control extracts may be explained by the low level of Hif-1{alpha} stabilization in confluent epithelial cell cultures as observed by Western blot analysis; however, in all cases, a significant increase in binding was detected with DFX treatment, suggesting further stabilization of Hif-1 in these cells. Our results suggest that the HREs identified in the upstream sequences of Ctgf genomic DNA represent functional hypoxia binding elements and can bind Hif-1 in vitro in response to Hif-1{alpha} stabilization.

Ctgf promoter assays further confirm the hypoxic regulation of Ctgf transcription by Hif-1. Since we have identified novel HREs in upstream sequences of the Ctgf genomic DNA that were capable of binding to Hif-1 in vitro, we needed to assess whether these sequences could enhance mRNA transcription levels in vivo in response to hypoxia. Luciferase constructs were prepared that contained either full-length 3.8-kb upstream fragments, including both HRE sites (3.8-kb construct) along with mutated constructs containing the individual HRE 5.2 site (1.7-kb construct), or neither HRE elements (1.0-kb construct). The genomic sequences were cloned upstream of the luciferase gene into pGL3 basic (Promega), which lacks promoter elements; therefore, luciferase expression occurs only in response to promoter elements present in the cloned 5'-region. Luciferase constructs were cotransfected into MCT or LLC-PK cells (immortalized renal tubular epithelial cell lines) with the Renilla luciferase plasmid under the control of SV40 promoter elements, since the low transfection efficiencies in our primary cultures precluded this analysis in these cells. Firefly luciferase activity was normalized to Renilla luciferase activity to account for variation in transfection efficiency. We consistently obtained higher overall luciferase activity with the shorter constructs (Fig. 5B comparing normoxic baseline values for 3.8-, 1.7-, and 1.0-kb constructs), which we believe was due to differences in transfection efficiency rather than to the presence of negative regulating factors in the full-length constructs, since transfection with equimolar amounts of each construct reduced the differences in basal normoxic activity; however, we cannot fully rule out this possibility. In addition, full-length constructs were prepared in which the putative HRE sites were mutated either individually ({Delta}5.1 and {Delta}5.2) or in combination ({Delta}{Delta}HRE), which eliminated the difference in baseline activity of the various constructs. A significant 2.0 ± 0.46-fold increase in luciferase activity was observed in the full-length 3.8-kb promoter construct exposed to hypoxic conditions of 0.5% O2 for 12 h (Fig. 5B, P < 0.05). This is comparable to the increase observed in endogenous Ctgf expression in response to hypoxic stimulation. No difference was observed for the 1.7-kb (containing 1 putative HRE site, HRE5.2) or the 1.0-kb constructs (containing neither HRE sites) under normoxia or hypoxia treatments. Similarly, the mutated full-length constructs {Delta}5.1, {Delta}5.2, and {Delta}{Delta}HRE did not show any increased activity under hypoxia, suggesting that both HRE sites are necessary for the observed increase in Ctgf mRNA in hypoxia-treated cells. Results are shown for transfections in LLC-PK cells, and similar results were obtained in the MCT cell line.



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  		Fig. 5. Ctgf promoter constructs containing 2 HREs display hypoxia response. Luciferase promoter constructs were prepared by inserting 3.8 kb of an upstream genomic DNA sequence for Ctgf into pGL3 basic luciferase plasmid, which does not contain any promoter elements. The presence of hypoxia-responsive promoter elements in this upstream sequence drives the expression of luciferase (LUC) in response to hypoxic stimulation. Deletion constructs were prepared in which either HRE 5.1 was absent (1.7 kb) or both HRE 5.1 and 5.2 were removed (1.0 kb). Full-length constructs were also prepared in which the HRE motifs were mutated either individually ({Delta}5.1, {Delta}5.2) or in combination ({Delta}{Delta}HRE). Mutations of the HRE sites were as described in Fig. 4B for the mutated probes used in the EMSA. A: schematic of the constructs displaying various mutated sites (X) along with (B) their respective luciferase values under normoxic (21%; open bars) and hypoxic (0.5% O2; gray bars) stimulation for 12 h. Transfections were performed in triplicate, and experiments were repeated at least 3 times. A representative experiment in LLC-PK cells is shown. SMAD/BCE-1, SMAD binding element and the basal control element-1; RLUS, relative light units. Error bars represent the SD of the triplicate samples. *P < 0.05, 2.0-fold increase over normoxia.

 
TGF-{beta}1 signaling is not required for hypoxic induction of Ctgf promoter activity. Ctgf is known to mediate certain effects of TGF-{beta}1, including upregulation of collagen I and fibronectin production in a number of renal cell types, including mesangial cells and fibroblasts (5, 8, 15). Hypoxia has been shown to increase TGF-{beta}1 in a number of cell types, including human renal fibroblasts (44) and human hepatoma cells (47). Since TGF-{beta}1 can induce Ctgf mRNA expression, we wanted to determine whether the observed increase in Ctgf promoter activity under hypoxia was partially mediated through increased TGF-{beta}1 activity. TGF-{beta}1 upregulates Ctgf mRNA expression via a SMAD-dependent signaling pathway (25). Activation of the TGF-{beta}1 receptor results in recruitment of SMAD 3 and its subsequent phosphorylation, enabling it to dimerize with SMAD 4 (41). The SMAD 3/4 complex enters the nucleus, where it binds to specific promoter elements, recruiting various transcription factor subunits and regulating the expression of genes including Ctgf. It has previously been shown that mutation of the SMAD binding site in the CTGF promoter can inhibit TGF-{beta}1-induced CTGF mRNA production (25). We utilized this mutation in our luciferase constructs to eliminate the effect of TGF-{beta}1 signaling on Ctgf expression during hypoxia. Constructs (3.8 and 1.0 kb) harboring the mutated SMAD sequence, from wild-type 5'-CAGACGG-3' (–396 to –390) to mutated 5'-gAtCCGG-3' (introducing a novel BamH1 restriction site), showed loss of TGF-{beta}1-induced luciferase activity compared with the wild-type constructs (Fig. 6A). The wild-type 3.8-kb construct showed a 2.1 ± 0.27-fold increase over control cells (P < 0.05), whereas the 3.8{Delta}SMAD construct displayed only a 1.2 ± 0.06-fold increase (P < 0.001) in response to TGF-{beta}1 stimulation for 6 h. Similar loss of TGF-{beta}1-induced transcriptional activity was observed in the 1.0-kb construct, from 2.4 ± 0.07-fold (P < 0.0001) in the wild-type SMAD construct to 1.2 ± 0.05-fold (P < 0.01) in 1.0{Delta}SMAD (Fig. 6A). When these constructs were subjected to combined TGF-{beta}1 and hypoxia treatments, the 3.8{Delta}SMAD construct displayed a 1.91 ± 0.16-fold increase (P < 0.01) in promoter activity, which must have been in response to increased Hif-1 levels observed during hypoxia given that the 1.0{Delta}SMAD construct did not show a similar change (Fig. 6B). In response to hypoxia alone, both the 3.8-kb and the 3.8{Delta}SMAD constructs showed similar increases in promoter activity (1.6 ± 0.14, P < 0.01 and 1.5 ± 0.04-fold, P < 0.0001, respectively), suggesting that TGF-{beta}1 does not play a significant role in Ctgf regulation during hypoxic conditions (Fig. 6C). The full-length constructs containing mutated HRE sites, with and without mutated SMAD sites, showed no changes in response to hypoxia (Fig. 6C), again supporting our finding that both HREs present in the 3.8-kb construct are required for the observed increase in Ctgf expression in response to hypoxia.



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  		Fig. 6. Hypoxia increases Ctgf promoter activity in the absence of TGF-{beta}1 signaling. A: mutation of the SMAD binding element renders the Ctgf promoter unresponsive to TGF-{beta}1 stimulation. MCT cells were transiently transfected with Ctgf promoter constructs (schematics depicting mutated regions shown in C) as described in EXPERIMENTAL PROCEDURES and treated with vehicle or 5 ng/ml TGF-{beta}1 for 6 h. B: 3.8-kb Ctgf promoter retains its hypoxia response in the absence of functional SMAD binding in the presence of combined TGF-{beta}1 and hypoxia treatment. C: hypoxia response of mutated and deleted constructs show that the full-length 3.8-kb construct contains sequences that are required for hypoxic induction of Ctgf promoter activity. Transfections were performed in triplicate, and experiments were repeated at least 3 times. A representative experiment in LLC-PK cells is shown. Values are means ± SD of the triplicate samples. Open bars, normoxic control; dark gray bars, 5 ng/ml TGF-{beta}1; black bars, hypoxia (0.05% O2) and 5 ng/ml TGF-{beta}1; light gray bars, hypoxia. #P < 0.05, *P < 0.01, and **P < 0.001 compared with normoxic control samples.

 

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CTGF has been identified as a major pathogenic molecular marker of renal fibrosis. Increased occurrence of CTGF has been associated with deteriorating renal function in a number of renal diseases that culminate in the development of tubulointerstitial fibrosis, including diabetic nephropathy, focal segmental glomerulosclerosis, IgA nephropathy (29), and obstructive nephropathy (24). CTGF acts as a mediator of TGF-{beta}1 signaling, specifically toward the dysregulation of extracellular matrix molecules including collagen I and fibronectin (20). Because TGF-{beta}1 is a key factor in the development of tubulointerstitial fibrosis, CTGF therefore represents a remarkable target to ablate the extensive TGF-{beta}1-induced effects on fibrosis. Understanding the pathways involved in regulating CTGF expression is of paramount importance in the development of strategies to target CTGF and modify its expression. Here, we report on the ability of Hif-1 to interact with promoter elements in genomic sequences upstream of Ctgf which increases promoter activity and mRNA transcription in response to hypoxia.

HIF-1 has been identified as a main molecular mediator enabling cell adaptation to hypoxia by facilitating vasculogenesis and increased O2 delivery as well as alteration of cell metabolism. More recently, it has been appreciated that HIF-1 can regulate genes involved in fibrogenesis, and it is now thought that a dysregulation of gene expression in response to lowered O2 levels within the kidney may contribute to the development of interstitial fibrosis (11, 12, 40, 44). Reduced peritubular capillary density and perfusion have been associated with regions of interstitial hypoxia during the development of renal disease (1, 45, 50). Therefore, it is plausible that HIF-1, stabilized in response to low O2 levels, regulates the expression of genes that underlie the development of renal tubulointerstitial fibrosis. Furthermore, nonhypoxic stimuli also result in HIF-1 stabilization, including nitric oxide, TNF-{alpha} (54, 75), IL-1 (66), angiotensin II, PDGF (66), and HGF (63), all of which have been implicated in the development of renal fibrosis. This further highlights the relative importance of HIF-1 as a possible mediator of tubulointerstitial fibrosis, because Hif-1 can increase expression of profibrogenic genes such as MMP-1, TIMP-3, and PAI-1 (35, 44, 46). To further define the role of hypoxia and HIF-1 in mediating the development of renal fibrosis, we investigated the effect of hypoxia on the regulation of the profibrogenic molecule Ctgf.

CTGF has previously been shown to be regulated by hypoxia in a human breast cancer cell line. Although HREs were not identified in the 800 bases immediately upstream of CTGF in this report (36), expression data generated in our laboratory from kidney-specific Vhl-null mice (i.e., mice with stabilized Hif-1{alpha} protein) and Vhl/Hif-1{alpha}-null mice pointed to a possible regulation of Ctgf by Hif-1 (Haase VH, unpublished observations). Furthermore, our findings suggested that Hif-1 was directly involved in the regulation of Ctgf expression because primary epithelial cell cultures deficient for Hif-1 no longer displayed hypoxic induction of Ctgf mRNA. To confirm a direct regulation of Ctgf by Hif-1 through interaction with classic HRE elements, we identified two HRE sites that were subsequently determined to bind Hif-1 directly in electrophoretic mobility shift assays and that displayed increased promoter activity in response to hypoxia as determined by in vitro reporter luciferase assays. Further characterization of these upstream sequences revealed a requirement for the presence of both HRE motifs for induction of a hypoxic response.

Hypoxic treatment led to an approximately twofold elevation in Ctgf mRNA; although we have not measured Ctgf protein levels, previous reports would indicate that a change in Ctgf mRNA expression of this magnitude may have pathophysiological implications. Studies have shown a strong correlation between Ctgf mRNA-expressing cells and sites of tubulointerstitial fibrosis (29). Analysis of unilateral ureteral obstructed kidneys displaying advanced stages of tubulointersitital fibrosis revealed a twofold increase in the in vivo level of Ctgf mRNA (24, 73). Furthermore, TGF-{beta}1 stimulation of tubular epithelial cells resulted in approximately a twofold mRNA increase (24), which was similar to that observed for other cell types, including mesangial cells (22), cardiac fibroblasts (4), and corneal fibroblasts (13). These findings would suggest that a twofold increase in Ctgf mRNA level is sufficient to exert a pathophysiological effect.

TGF-{beta}1 is a major contributing factor to the development of fibrosis and can induce CTGF expression through interaction of the SMAD 3/4 complex with the SMAD binding element situated in the CTGF promoter (25). TGF-{beta}1 activity can be increased in response to hypoxia (46, 47, 74); therefore, to ensure that the observed hypoxia-induced Ctgf expression was not a result of increased bioactivity of TGF-{beta}1, we analyzed a number of luciferase promoter constructs that were rendered unresponsive to TGF-{beta}1 treatment by a previously described mutation of the SMAD binding element (25). As expected, mutation of the SMAD binding element inhibited the ability of TGF-{beta}1 to increase Ctgf promoter activity. However, increased promoter activity was observed in response to hypoxia in these constructs, confirming the presence of functional HRE sites that could positively regulate promoter activity without the requirement for TGF-{beta}1 signaling. Although a synergistic increase in Ctgf promoter activity in response to combined TGF-{beta}1 and hypoxic stimulation was not observed in our reporter assays, we cannot disregard the possibility that these two factors may act synergistically, as in the case of VEGF mRNA regulation (53). Because our reporter assays were terminated after 12 h of stimulation, it is possible that longer stimulation past 24 h may reveal synergism of these two factors.

Although we cannot rule out the possibility of downstream Hif-1 target genes positively influencing the activity of the Ctgf promoter under hypoxia, our data from the luciferase constructs containing mutated HRE sites suggest that Hif-1 can directly interact with these elements and is required for the observed increase in transcriptional activity under hypoxia. For instance, VEGF is a known target for HIF-1 and has been shown to enhance CTGF mRNA expression (62). However, the kinetics of VEGF expression in response to hypoxia would preclude it from playing a role in our observed hypoxic induction of Ctgf. VEGF protein levels are statistically increased after 12 h of hypoxic stimulation at 1% O2 in Hep3B cells (53) to levels that are capable of inducing CTGF mRNA expression; however, stimulation with VEGF typically takes a further 6 h to statistically increase CTGF mRNA levels (62). Because we have observed increased expression of Ctgf mRNA as early as 1 h of hypoxic stimulation, we can conclude that Vegf is not actively inducing Ctgf in our assays.

Analysis of the distribution of Hif-1 and Hif-2 in the kidney revealed that Hif-1 is predominantly expressed in tubular cells in response to systemic or local hypoxia induced by anemia, carbon monoxide, or cobalt chloride, whereas Hif-2 was predominantly expressed in interstitial endothelial and peritubular cells (51). We show here that PTCs can respond to lowered levels of O2 by stabilizing Hif-1. In support of previous evidence regarding a lack of Hif-2 expression in tubular epithelial cells (51), we were unable to detect significant Hif-2{alpha} protein levels in our primary cell cultures. Furthermore, when Hif-1{alpha} was deleted from our PTCs, hypoxic induction of the Hif-2 target gene Vegf was no longer observed by RPA analysis, confirming Hif-1 as the predominant hypoxia-inducible factor produced in these cells. It has previously been suggested that cell cultures may not reflect the true in vivo situation in that both Hif-1 and Hif-2 can be expressed in culture, whereas in vivo often selective expression of only one Hif is observed (11). Here, we report good correlation between the in vivo situation and the observed in vitro expression of hypoxia-inducible factors. Furthermore, we report on a novel gene-targeting system for Hif-1{alpha}, where primary cell cultures can be grown as wild-type cells and then mutated for Hif-1{alpha} by doxycycline treatment. This system enables precise deletion of Hif-1{alpha} while avoiding confounding issues such as cell toxicity, which is usually associated with adenoviral or similar transfection techniques.

HIF-1 is an important cellular factor that enables cells to adapt to hypoxic environments. Loss of HIF-1 expression in our conditionally mutated primary epithelial cell cultures, however, did not affect the ability of the cells to survive exposure to hypoxic conditions of 0.5% O2. Indeed, in our system, survival studies have shown that loss of Hif-1 had no discernable effect on cell viability when cultures were exposed to O2 levels as low as 0.2% for 48 h provided the glucose availability was not limiting (Biju MP and Haase VH, unpublished observations). Because our hypoxic experiments reported here were performed at 0.5% O2 for a maximum of 24 h in the presence of nonlimiting glucose levels, differences in cell viability were considered negligible.

In conclusion, we show here that hypoxia regulates Ctgf expression through a Hif-1-dependent and TGF-{beta}1-independent pathway, suggesting that although CTGF appears to mediate many of the profibrogenic effects of TGF-{beta}1, it may also display TGF-{beta}1-independent profibrotic properties. While CTGF has been suggested as a possible therapeutic target against ESRD and development of fibrosis, pharmacological manipulation of HIF-1, which is an important regulator of profibrotic gene expression, may offer therapeutic potential. In summary, our findings highlight Hif-1 as a regulator of hypoxic Ctgf expression, which is not only relevant for renal fibrosis but has pathogenic implications for fibrotic diseases in general, as well as for cancer as increased CTGF positively correlates with metastatic potential (59, 72).


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This work was supported in part by grants from the National Institute of Diabetes and Digestive and Kidney Diseases (R03-DK-62060 to V. H. Haase), the Charlotte Geyer Foundation, and seed money from the Department of Medicine (V. H. Haase). D. F. Higgins was supported by a Postdoctoral Fellowship from the American Heart Association.


   ACKNOWLEDGMENTS
 
We thank Fuad Ziyadeh for critical review of the manuscript, Eric Neilson and Masayuki Iwano for providing the MCT cell line, Peter Friedman for providing the immortalized distal tubular cell line, and Hermann Bujard for the kind gift of the Lc-1 mouse.

Current address of Y. Akai: Nara Medical Univ., First Dept. of Internal Medicine, 840 Shijo-cho, Kashihara, Nara 634-8522, Japan.


   FOOTNOTES
 

Address for reprint requests and other correspondence: V. H. Haase, Dept. of Medicine, 700 Clinical Research Bldg., 415 Curie Bld., Philadelphia, PA 19104-6144 (E-mail: vhaase{at}mail.med.upenn.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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