Departments of 1 Medicine and Physiology and Biophysics, State University of New York, Stony Brook, New York 11794-8152; 2 Tokai University School of Medicine, Kanagawa, Japan; 3 Department of Medicine, Tel Aviv University, Israel; and 4 Department of Medicine, University College London, London, United Kingdom
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ABSTRACT |
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Development of micro- and macrovascular disease in diabetes mellitus (DM) warrants a thorough investigation into the repertoire of endothelial cell (EC) responses to diabetic environmental cues. Using human umbilical vein EC (HUVEC) cultured in three-dimensional (3-D) native collagen I (NC) or glycated collagen I (GC), we observed capillary cord formation that showed a significant reduction in branching when cells were cultured in GC. To gain insight into the molecular determinants of this phenomenon, HUVEC subjected to GC vs. NC were studied using a PCR-selected subtraction approach. Nine different genes were identified as up- or downregulated in response to GC; among those, plasminogen activator inhibitor-1 (PAI-1) mRNA was found to be upregulated by GC. Western blot analysis of HUVEC cultured on GC showed an increase in PAI-1 expression. The addition of a neutralizing anti-PAI-1 antibody to HUVEC cultured in GC restored the branching pattern of formed capillary cords. In contrast, supplementation of culture medium with the constitutively active PAI-1 reproduced defective branching patterns in HUVEC cultured in NC. Ex vivo capillary sprouting in GC was unaffected in PAI-1 knockout mice but was inhibited in wild-type mice. This difference persisted in diabetic mice. In conclusion, the PCR-selected subtraction technique identified PAI-1 as one of the genes characterizing an early response of HUVEC to the diabetic-like interstitial environment modeled by GC and responsible for the defective branching of endothelial cells. We propose that an upregulation of PAI-1 is causatively linked to the defective formation of capillary networks during wound healing and eventual vascular dropout characteristic of diabetic nephropathy.
glycation end products; extracellular matrix; diabetic nephropathy; angiogenesis; cDNA differential display; plasminogen activator inhibitor-1
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INTRODUCTION |
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ADVANCED GLYCATION END PRODUCTS (AGEs) are considered among the leading causes of diabetic complications, especially in the development of atherosclerotic vascular disease (7, 8, 54, 55). These are linked to changes in physicochemical properties of matrix proteins, like collagen-to-collagen crosslinking and tissue rigidity, leading to decreased solubility and susceptibility of proteins to enzymatic digestion (50-57). In fact, infusion of AGEs into nondiabetic animals can reproduce many vascular complications of diabetes (51).
Recently, AGEs have been suggested as modulators of endothelial cell proliferation, migration, and capillary tube formation. In this vein, serious controversies exist as to the spectrum of cell functions perturbed and the direction of functional changes attributed to various glycated proteins (17, 40, 45, 54). This list of contradictory data emphasizes the need to revisit the role of glycated matrix proteins in modulating endothelial cell functions.
Development of diabetic nephropathy is associated with a notable dropout of blood vessels in the kidney. Several investigators have observed a decrease in the cross-sectional surface area of capillaries in renal biopsies from patients with fibrointerstitial disorders including diabetic nephropathy (6, 22, 41). In patients with diabetic nephropathy, among other causes of progressive interstitial fibrosis, the degree of vascularization was found to be significantly decreased, and this single parameter had the highest predictive value for the development of renal dysfunction (41). These findings warrant investigation into the mechanism(s) of defective angiogenesis in diabetic nephropathy. One of these mechanisms could be related to nonenzymatic glycation of deposited collagen I at sites of profibrotic lesions, which may modulate their vascularization. This question was addressed in the present study.
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MATERIALS AND METHODS |
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Cell culture, RNA purification, and cDNA synthesis. HUVEC, used between passages 3 and 8, were cultured in 100-mm dishes in endothelial cell growth medium (EGM)-2 (Clonetics) containing 2% fetal bovine serum (FBS), 100 U/ml penicillin, and 50 µg/ml streptomycin. Immortalized renal microvascular endothelial cells (46) were propagated in the same medium. Cells were trypsinized from the dishes when they reached 80-90% confluence and transferred to 100-mm dishes precoated overnight at 4°C with 2.5 ml nonglycated or glycated collagen I (GC; 50 µg/ml). Twenty-four hours later, cells were harvested, and the total cell RNA was isolated with TRIzol RNA isolation reagent (GIBCO-BRL) according to the protocol provided by the manufacturer. Total RNA was dissolved in 50 µl deionized water, and 250-500 ng RNA were subjected to full-length cDNA synthesis using a SMART PCR cDNA synthesis kit (Clontech) in 100-µl vol and with 18 PCR cycles, according to the manufacturer's instructions. Full-length cDNA was checked on a 1.0% agarose gel, and the cDNA was synthesized in two to three tubes mixed together and purified with Qiaquick PCR purification kit (Qiagen).
Preparation of glycated collagen I and three-dimensional (3-D)
cultures of endothelial cells.
Vitrogen (Cohesion, Palo Alto, CA) was dissolved in 500 mM
D-glucose (dextrose, dissolved in PBS, pH 7.0-7.4) to
reach a concentration of 100 µg/ml, with pH adjusted to 7.4 with 0.1 N NaOH. After sterilization by passing through a 2.2-µm syringe
filter, the solution was incubated at 29°C for 2-3 wk or at
36°C for 4 wk. The endotoxin was removed with a polymixin B
immobilized endotoxin-removing agarose gel column (Detoxi-Gel, Pierce,
Rockford, IL), and the solution was aliquoted and stored at 20°C.
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Subtractive hybridization and selective PCR amplification.
The full-length cDNA prepared from the RNA that was isolated from
cells grown on nonglycated collagen-coated dishes served as driver
cDNA. In contrast, cDNA synthesized from the RNA that was isolated from
cells grown on glycated collagen-coated dishes served as tester cDNA.
Full-length cDNAs (tester and driver), 2-2.5 µg each, were used
as starting material, and the subtraction hybridization and
PCR-selected amplification were performed using the PCR-select cDNA
subtraction kit. Briefly, the tester and driver cDNA were first
digested with Rsa I in a 50-µl reaction mixture containing
15 U of enzyme for 3 h. The cDNAs were then phenol-extracted, ethanol-precipitated, and resuspended in 5.5 µl of dideionized water.
Next, 1 µl of digested tester and driver cDNA was diluted in 5 µl
of H2O, and 2 µl of each diluted cDNA were ligated to 2 µl of adapter 1 and adapter 2R, respectively, in a total volume of 10 µl at 16°C overnight, using 200 units of T4 DNA ligase (New England
Biolabs). After ligation, 1 µl of 0.2 M EDTA-glycogen (1 mg/ml) mix
was added, and samples were heated at 72°C for 5 min to inactivate
the ligase, and stored at 20°C. For the first hybridization, 1.5 µl of Rsa I-digested driver cDNA were added to each of two tubes
containing 1.5 µl of adapter 1- and adapter 2-ligated tester cDNA,
respectively, and 1 µl of 4× hybridization buffer. After an overlay
with mineral oil, the solution was heated to 98°C for 1.5 min to
denature the cDNAs and then allowed to anneal at 68°C for 10 h.
After this first hybridization, the two samples were combined and a
fresh portion of heat-denatured driver cDNA (1 µl driver cDNA that
was 1:4 diluted with 1× hybridization buffer ) was added at the same
time. The sample was then allowed to hybridize for an additional
10 h at 68°C. The final hybridization was then diluted in 200 µl of dilution buffer (20 mM HEPES, pH 8.3, 50 mM NaCl, and 0.2 mM
EDTA), heated at 68°C for 7 min, and stored at
20°C. We performed
two PCR amplifications for each subtraction. The primary PCR was
conducted in 25 µl. It contained 1 µl of the above diluted,
subtracted cDNA, 1 µl PCR primer 1, and 23 µl of PCR master mixture
prepared using the Advantage cDNA PCR kit (Clontech). PCR was performed
using the following parameters: 75°C for 7 min, 27 cycles at 94°C
for 30 s, 66°C for 30 s, and 72°C for 1.5 min. The
amplified products were diluted 10-fold in deionized water. One
microliter of this diluted product was then subjected to secondary PCR
for 12 cycles under the same conditions used for the primary PCR,
except that PCR primer 1 was replaced with nested PCR primer 1 and 2R,
and the annealing temperature was 68°C instead of 66°C.
Cloning and analysis of subtracted cDNA. Products from the secondary PCR were inserted into a PCRII vector with a T/A cloning kit (Invitrogen). Plasmid DNAs were prepared using a QIAprep spin miniprep kit (Qiagen). The plasmid DNAs obtained from the subtracted library were hybridized with forward- and reverse-subtracted cDNA probes according to the manufacturer's instructions for the PCR-select differential screening kit. Briefly, primary PCR product from the forward and reverse subtraction experiments were used in secondary PCR as described above. After Rsa I, Smal I, and Eag II digestion and purification with the QIAquick PCR purification kit (Qiagen), digoxigenin (DIG)-labeled forward and reverse probes were prepared using random labeling procedures (DIG random labeling kit, Boehringer) from the products generated in the secondary PCR. Plasmid DNA were dot-blotted onto nylon membrane and screened by hybridization with the forward and reverse probes according to standard hybridization procedure. Clones representing mRNAs that are differentially expressed, which hybridized strongly (5 times more) with the forward-subtracted probe were sequenced by the chain termination reaction using an ABI Prism BigDye Terminator Cycle Sequencing Ready Reaction Kit (PE Applied Biosystems). Nucleic acid homology searches were performed using the BLAST program.
Northern blot analysis.
Total cell RNA was extracted with the TRIzol RNA isolation
reagent (GIBCO-BRL). The RNA concentration in each sample was
determined by spectrophotometry, and only samples with a 260/280 ratio
close to 1.9 were analyzed. RNAs (15 µg/lane) were separated by
electrophoresis in 1.4% formaldehyde-agarose gels and transferred to
nylon membrane (Millipore) in 10× saline-sodium citrate (SSC).
Membranes were prehybridized for 3 h at 52°C in 50% formamide
(vol/vol) in prehybridization buffer. They were then hybridized for
18 h at 50°C in 50% formamide (vol/vol), 1× hybridization
solution with plasminogen activator inhibitor-1 (PAI-1) cDNA probes
labeled with DIG-dUTP by the random priming method. Afterward, the
membranes were washed twice for 15 min at room temperature in 2× SSC,
0.1% SDS, and twice for 15 min at 68°C in 0.5× SSC, 0.1% SDS. The
CDP-Star Kit (Boehringer Mannheim) was used to detect the DIG-labeled
probes by exposing the filters to X-ray film (Denville Scientific) for
5 min at room temperature followed by densitometric scanning of the
bands. Northern blot analyses of collagen 1 and tissue inhibitor of
metalloproteinases (TIMP)-1 mRNA expression were performed using
[32P]dCTP-labeled cDNA probes, as described previously
(34).
Western blot analysis. Ten micrograms of total protein from each sample were run on 4-20% Tris-glycine gel (Novex). The proteins were transferred to Immobilon-P (Millipore) and blocked for 45 min with 1% casein in PBS. The membranes were incubated at room temperature for 1 h in mouse monoclonal anti-PAI-1 antibody (American Diagnostica) and diluted 1:50 in PBS. After being washed three times in PBS/0.1% Tween, the membranes were incubated in rabbit anti-mouse Ig-horseradish peroxidase (HRP; Amersham) 1:2,000 in PBS for 30 min. Membranes were washed three times in PBS/0.1% Tween and developed using Phototope-HRP (New England Biolabs). The membranes were exposed to Wolf RXB X-ray films.
Reverse zymography. Twenty micrograms of total protein lysate were run on a 4-20% Tris-glycine gel and washed for 10 min in 2.5% Triton X-100 (Fischer Scientific) and 10 min in PBS. The gels were overlaid onto 1.25% agar (Difco) containing 40 mg/ml plasminogen (American Diagnostica), 250 mU/ml human urokinase (Calbiochem), and 2% nonfat dry milk powder (wt/vol) (Carnation) in 0.1 mol/l Tris, pH 8.0 (16). The reverse zymograms were developed at 37°C for 3-6 h and visualized using darkfield illumination. To study the activity of matrix metalloproteases, gelatin-substrate gel zymography was performed as detailed previously (21, 35).
Cell attachment and proliferation assays. After trypsinization, cells were resuspended in endothelial cell medium (ECM; Clonetics) containing 2% FCS (GIBCO-BRL), 10 U/ml penicillin, 100 µg/ml streptomycin, 0.25 µg/ml amphotericin B (antibiotic-antimycotic; Sigma) and plated in 6-cm dishes (105cells/dish) coated with either nonglycated or glycated collagen type I. Cells were assessed qualitatively under phase contrast at 2, 4, 6, and 24 h post-plating. Alternatively, cell attachment was monitored using an electrical cell impedance system, as previously detailed (32).
Cell proliferation was measured using the nonradioactive colorimetric assay described by Oliver et al. (33). Briefly, cells were plated in growth medium in flat-bottomed 96-well plates coated with either native or glycated collagen type I at a concentration of 2 × 103 cells/well and harvested 1, 3, or 5 days after plating. Cells were washed three times with PBS pH 7.2, blotted dry, and fixed in formol saline (10% formaldehyde in 0.9% NaCl) for at least 30 min at ambient temperature. Cells were stained with 0.1% methylene blue (Sigma) in 0.01 M borate buffer, pH 8.5, for 30 min at ambient temperature. Unbound dye was removed by six washes with borate buffer using an automated plate washer (Denley), and the plates were blotted dry on absorbent paper. Dye was eluted from the cells with ethanol-0.1 M HCl (1:1), and absorbance was 650 nm in a plate reader (Titertek).Explant aortic cultures. Thoracic aortas were obtained from 6- to 8-wk-old mice. The wild-type (C57BL/6J) and PAI-1 knockout (C57BL/6J Planh1) mice were utilized for these studies. In a separate series of studies, animals were injected with 100 mg/kg body wt streptozotocin intraperitoneally, and development of hyperglycemia was verified by measurements of glycosuria and blood glucose levels. Animals were studied 8 wk after the induction of diabetes. Mice were anesthetized by intraperitoneal injection with ketamin-xylazine, and a thoracotomy was performed. The full length of the thoracic aorta was aseptically removed and immediately placed into a 35-mm culture dish with ice-cold EBM-2 medium. After removal of the periaortic fibroadipose tissue with fine microsurgical forceps under a dissecting microscope, the vessels were placed in fresh EBM-2 medium as described previously (30, 31). The aortas were then cross-sectioned with a 1-mm interval, and the resulting aortic rings were embedded in 3-D native or glycated collagen I gels in culture chambers (Nalge Nunc), as previously described (30, 31). Collagen I gels were prepared according to Montesano et al. (28). Briefly, 7 vol of vitrogen (Cohesion) were quickly mixed at 4°C with 1 vol 10× EBM-2 and 2 vol of 11.7 mg/ml sodium bicarbonate. Gelation was allowed to take place by incubating the cultures at 37°C for 30 min. After gelation, 400 µl of EBM-2 medium were added to each gel, and the chambers were placed in a 95% air-5% CO2 incubator at 37°C.
Quantitative analysis of angiogenesis in explant cultures was performed using an inverted microscope. Vascular sprouts were counted along the perimeter of each explant by two independent observers blinded to the origin of explant cultures, under ×100 magnification, with images captured using a SONY XC-77 camera and displayed on a video monitor. Newly formed capillary cords were counted daily for 7 days. ![]() |
RESULTS |
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Capillary cord formation, attachment, and proliferation in glycated collagen I. When HUVEC or renal microvascular endothelial cells were cultured in collagen I gels, either native or glycated, the pattern of capillary tube formation showed striking differences. Twenty-four hours after the plating, HUVEC formed elongated, cordlike structures that were densely branched in the native collagen, but much sparser branching was detected in the glycated collagen matrix (Fig. 1, A, C, and D). Initially, these capillary cords formed no lumen, but within 3-5 days lumen was detectable, as demonstrated on an electron micrograph (Fig. 1B). To quantitate this phenomenon, the number of cell bodies between two consecutive branches was counted in each case. Analysis of branching density revealed that 24 h after plating in glycated collagen I, HUVEC displayed about half the density seen in native collagen gels (Fig. 1D; P < 0.05, Wilcoxon test). These differences became less pronounced 48-72 h later.
Qualitative assessment of cell attachment on the surface of gels showed that, 2 h after plating, cells had begun to attach and spread; the number of cells attached and the degree of cell spreading increased with time. There was a delay in attachment and spreading of HUVEC on glycated collagen, compared with the native substrate, but differences disappeared by 2 h postplating (not shown). HUVEC proliferation on glycated collagen showed a less marked increase between 1 and 5 days compared with that seen on nonglycated matrix (not shown). Collectively, the above findings raised the question on the identity of genotypic responses of HUVEC to glycated collagen early after its presentation, when the potential impact of uneven cell attachment has already disappeared, and while the consequences of unequal cell proliferation have not yet resulted in significant changes in cell population. To address this question, we next performed a series of experiments designed to detect potential candidates for delayed branching using a differential DNA screening.Differential screening and identification of differentially
displayed cDNA clones.
Cells presented with control or glycated collagen I were studied
24 h later, at the time when the differences in branching were
maximal. The subtracted library was screened with the forward- and
reverse-subtracted probes. Sixteen clones were found that strongly
hybridized (at least 5 times more) with the forward probe compared with
results obtained with the reverse probe. After partial sequencing and a
search of the gene bank, those 16 clones were identified as 9 different
genes (Fig. 2). Among them, three clones were identified as the PAI-1 gene. One of these clones (A18) containing a 1-kb insert of the PAI-1 gene was chosen as a template for probe labeling that was employed for Northern hybridization.
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Northern blot hybridization, expression, and activity of PAI-1,
matrix metalloproteinase (MMP)s, TIMP-1, and collagen I.
Total RNA extracted from HUVEC grown on collagen I or glycated collagen
I for 12, 18, 24, and 36 h was electrophoretically separated and
hybridized with a DIG-labeled PAI-1 cDNA probe. PAI-1 mRNA showed no
changes with time in collagen I culture but was induced threefold at
12 h in glycated collagen I cultures, with the subsequent return
to baseline at 36 h (Fig.
3A).
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Capillary cord formation in the presence of neutralizing PAI-1
antibodies or activated PAI-1.
To determine the causative relationship between the upregulation of
PAI-1 and the observed delay in capillary cord formation, in the next
series of experiments a neutralizing antibody against PAI-1 was added
to the culture medium of HUVEC grown in glycated and native collagen I. The antibody was introduced at concentrations of 0.05-5.0 µg/ml,
5 h after initiation of cell culture. Results demonstrated that
0.5 and 5.0 µg/ml, but not 0.05 µg/ml, restored the branching
pattern of endothelial cells grown in glycated collagen I to
the level seen on native collagen (Fig.
5). Next, we performed the reverse
procedure, namely, examining capillary cord formation in native
collagen I or Matrigel with the addition of various concentrations of a
recombinant constitutively active human PAI-1 mutant (1-100 ng/ml;
American Diagnostica). HUVEC in collagen I showed a significant
decrease in branching pattern at PAI-1 concentrations of 1 and 10 ng/ml, with 100 ng/ml PAI-1 resulting in a scatter of HUVEC (in all
experiments, PAI-1 was added 12 h after plating). Cells cultured
in Matrigel showed no detectable changes in branching pattern in the
presence of 1 ng/ml PAI-1, but higher concentrations of PAI-1, 10 and
100 ng/ml, resulted in the increased length of capillary cords, which
exhibited reduced branching (Fig. 6).
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Ex vivo aortic explants from PAI-1 /
mice: capillary sprouts in
native and glycated collagen I.
In the next series of experiments we made use of PAI-1 knockout mice
and analyzed the formation of capillary sprouts in explant aortic
cultures. Using this well-established technique (30, 31),
we were able to confirm the endothelial origin of sprouts [staining of
cells with antibodies to the von Willebrand factor, platelet
endothelial cell adhesion molecule (PECAM), and acetylated low-density
lipoproteins; data not shown]. Figure 7
summarizes the results of quantitation of capillary sprouts in ex vivo
cultures. Aortic explants obtained from wild-type mice exhibited a
significantly slower sprouting in GC than in the native 3-D collagen I
lattices. In contrast, aortic explants obtained from PAI-1
/
mice
showed accelerated sprouting in native collagen gels compared with in the wild type, but this process was not inhibited in glycated collagen
(Fig. 7B). These ex vivo observations add further credence to the above in vitro findings obtained in cultured endothelial cells.
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Ex vivo aortic explants from PAI-1 /
mice: angiogenesis in
streptozotocin diabetic mice.
To further challenge the hypothesis on the involvement of PAI-1
in defective angiogenesis in the course of diabetes, explant cultures
were initiated from wild-type and PAI-1
/
mice 8 wk after
streptozotocin injection, as detailed in METHODS. Data
presented in Fig. 8 demonstrate, as
before, the higher propensity for angiogenesis in PAI-1
/
vessels
compared with the wild-type controls. Streptozotocin diabetes resulted
in the suppression of endothelial sprouting in both groups of animals
(Fig. 8B); however, aortic explants obtained from wild-type
diabetic animals showed consistently lower angiogenic capacity than
those obtained from PAI-1
/
diabetic mice (in addition to the
differences on days 4 and 5, as determined by
t-test, ANOVA analysis showed significant difference between the 2 curves). These findings provide ultimate evidence in support of
the proposed role of PAI-1 in defective angiogenesis.
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DISCUSSION |
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The data presented herein show that glycated collagen I delays
endothelial cell branching during in vitro capillary tube formation. This process is chronologically associated with the transcriptional activation of several genes including the PAI-1 gene and induction of
PAI-1 expression and activity. Neutralizing antibody to PAI-1 added to
HUVEC cultured in glycated collagen-1 restores the normal branching
pattern observed in cultures with native collagen-1. The relevance of
these observations is further emphasized by the finding that capillary
sprouting is resistant to glycated collagen lattices when it occurs in
aortic explants from PAI-1 knockout, but not from wild-type mice. The
data are further supported by the finding that streptozotocin diabetis
in PAI-1 /
mice results in a lesser disturbance of angiogenic
capacity than in wild-type animals.
It has been established that patients with diabetic nephropathy exhibit a decrease in the cross-sectional surface area of capillaries in renal biopsies (6, 22, 41). In these patients, the degree of vascularization was found to be significantly decreased, and this single parameter had the highest predictive value of renal dysfunction (41). The question is, Does this vascular dropout represent a cause or a consequence of nephrosclerosis? Given the fact that patients with diabetic nephropathy in the presence of interstitial fibrosis tend to display vascular dropout (6, 22, 41) and that in vitro cultures of endothelial cells on glycated collagen I exhibit a delay in branching of capillary cords, our findings may provide a pathophysiological link among the glycation of matrix proteins, accumulation of extracellular matrix, and failed angiogenic responses. Indeed, studies in streptozotocin-induced diabetes in rats showed that the accumulation of PAI-1 was most pronounced in animals with advanced diabetic nephropathy (Brodsky S, unpublished observations). It remains unclear why the defective angiogenesis in diabetes is confined to the kidney, myocardium, neural trunks, or wound healing, in contrast to the enhanced angiogenesis in the retina (proliferative retinopathy).
In the past, diabetic nephropathy was considered primarily a glomerular disease, but recent evidence has shown that the decline in renal function correlates with the development of tubulointerstitial injury (20). An early pathological change in the renal interstitium is an increase in the tubular basement membrane mass. More advanced disease is characterized by the extracellular matrix expansion of the glomerular mesangium, renal arteriolosclerosis, and tubulointerstitial fibrosis (57). Basement membrane thickening results from the binding of nonenzymatic glycated plasma proteins (AGEs) that alter the meshwork structure of the matrix (1, 2). AGEs are known to elicit changes such as vascular dysfunction, matrix expansion, and glomerulo- and atherosclerosis (7, 8, 49-51). The structural changes induced by AGEs impair the ability of basement membrane components to associate with each other to form the ordered polymeric complex. Some glycated proteins have been associated with impaired cell functions. For instance, nonenzymatic glycation of intact basement membrane or its individual components was reported to result in increased proliferation of retinal endothelial cells and suppressed proliferation of retinal pericytes (17). In addition, glycated fibronectin was found to stimulate cell surface expression of intercellular adhesion molecule-1, vascular cell adhesion molecule, and PECAM-1, thus synergizing with inflammatory mediators in facilitating the diapedesis of inflammatory cells (40). AGE-modified albumin was shown to induce tube formation by human skin microvascular endothelial cells on Matrigel and prevent their apoptosis (51) by acting through the autocrine vascular endothelial growth factor. In HUVEC, AGE-albumin did not affect proliferation, but it stimulated cell migration in a Boyden chamber and capillary tube formation in Vitrogen (45).
The discovery of transcriptional activation of the PAI-1 gene and induction of its product 12-24 h after presenting endothelial cells with glycated collagen I was unexpected but not surprising. Glycation of fibrinogen and several other components of the t-PA-catalyzed plasmin formation resulted in increased plasminogen activation, and glycated fibrin increased PAI-1 binding and activity (5). In addition, it has been reported that AGE-modified albumin induces PAI-1 mRNA concomitant with the increase in abundance of this serine protease inhibitor and its activity in human microvascular endothelial cells (54). Furthermore, serial analysis of gene expression (SAGE) in endothelial cells exposed to atherogenic stimuli revealed 56 differentially expressed genes, PAI-1 being one of them (12). Murphy et al. (29) have observed the induction of several genes, including PAI-1, in mesangial cells incubated in media containing a high glucose concentration. High glucose and hyperosmolality have also been implicated in PAI-1 induction in HUVEC (23). Our own data in endothelial cells confirm PAI-1 induction by high glucose levels, although the dynamics of this process is different from the induction by glycated collagen. In fact, our data on PAI-1 induction by a glycated matrix protein provide, to the best of our knowledge, the first demonstration of the early response of this gene in endothelial cells to environmental cues, reminiscent of the diabetic microenvironment in the interstitium undergoing fibrogenesis, and link it to the defective branching angiogenesis in vitro.
There is growing awareness that PAI-1 may have pathophysiological implications in the course of diabetic nephropathy and/or atherosclerosis. Temelkova-Kurktschiev et al. (44) have found a direct correlation between Hgb A1c, PAI-1, insulin level, and intima-media thickness in 40- to 70-year-old nondiabetic patients with a familial history of type II diabetes mellitus. Accelerated atherosclerosis and thrombosis are the major causes of morbidity and mortality in kidney transplant recipients. In cyclosporin A-treated patients, t-PA and especially PAI-1 activity were increased, suggesting the contribution of PAI-1 to thrombogenicity (48). Recent studies by Kimura et al. (18) have demonstrated a 4/5-guanine tract polymorphism in the promoter region of the PAI-1 gene in diabetic patients experiencing macroangiopathy and showed that the PAI-1 4G4G genotype (which leads to elevated levels of circulating PAI-1 activity compared with the 5G5G genotype) is an independent risk factor for development of macroangiopathy. Expression of PAI-1 promotes antifibrinolysis and accumulation of collagen (4, 47). With consideration of the progressive nature of matrix accumulation and its glycation in patients with diabetic nephropathy, the early-phase induction of the PAI-1 gene and a later sustained elevation in PAI-1 expression may be pathophysiologically linked in orchestrating the permissive mechanisms for development of tubulointerstitial fibrosis and deficient vascularization. Our data demonstrate that, when endothelial cells are repeatedly presented with glycated collagen, the increased expression of PAI-1 persists, in contrast to the single exposure, which is accompanied by a reversible induction of PAI-1. These data suggest that the process may become autocatalytic in the presence of factors promoting glycation of matrix proteins.
Angiotensin II is considered one of the key stimuli for PAI-1 induction
in endothelial and vascular smooth muscle cells (15), which in turn inhibits plasmin-mediated proteolysis resulting in
antifibrinolysis and, more relevant to this study, in the accumulation of extracellular matrix. Sakurai and Nigam (36) provided
evidence that transforming growth factor- is yet another potent
inducer of PAI-1 gene expression in Madin-Darby canine kidney and inner medullary collecting duct (mIMCD3) epithelial cells, where it suppresses tubular branching. The present study has broadened the
repertoire of mechanisms for PAI-1 induction by providing the first
evidence of its early stimulation by glycated collagen I. Whereas the
signaling pathway for PAI-1 induction remains unknown in our cells,
involvement of nuclear factor-
B in the cascade initiated by the
ligation of the receptor for advanced glycation end products has been
reported (38).
Recent studies employing several strains of genetically engineered mice
have shed light on the role of components of fibrinolytic cascade such
as urokinase (uPA) and its cognate receptor uPAR, tissue plasminogen
activator, PAI-1, and plasminogen in regulating wound healing,
fibrogenesis, and cell migration (9, 10, 36). Although
mice overexpressing PAI-1 exhibit pulmonary fibrosis after exposure to
bleomycin, PAI-1 knockout mice appear to be protected against this
complication as well as against hyperoxic lung injury (3,
14). It has been recognized that uPAR and 1-integrins are
colocalized in caveolae, where they regulate integrin function, cell
adhesion, and motility (42). Activated PAI-1 inhibits uPA
interactions, through uPAR adhesion to vitronectin, and interferes with
V
3 receptor-mediated cell migration (11, 19, 43,
56). Such an action of PAI-1 on endothelial cell migration is
probably involved in the observed defective branching in glycated
collagen I. Our experiments in PAI-1 knockout mice support the
conclusion that PAI-1 is intricately involved in angiogenesis. This
model approach will be helpful in further investigation of the
molecular mechanisms of vascular remodeling in diabetes mellitus.
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ACKNOWLEDGEMENTS |
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These studies were supported in part by National Institute of Diabetes and Digestive and Kidney Diseases Grants DK-45462, DK-54602, and DK-45695 (M. S. Goligorsky). The data were presented in a preliminary form at the 32nd Annual Meeting of the American Society of Nephrology, Miami Beach, FL, November 1999.
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FOOTNOTES |
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* J. Chen and S. Brodsky contributed equally to the studies.
Address for reprint requests and other correspondence: M. S. Goligorsky, Dept. of Medicine, State Univ. of New York, Stony Brook, NY 11794-8152 (E-mail: mgoligorsky{at}mail.som.sunysb.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.
Received 18 August 2000; accepted in final form 15 February 2001.
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