Departments of 1Physiology and 2Anesthesiology, Medical College of Wisconsin, Milwaukee, Wisconsin 53226; and the Department of 3Physiology, Louisiana State University Health Science Center, New Orleans, Louisiana 70112
Submitted 10 September 2003 ; accepted in final form 30 December 2003
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ABSTRACT |
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renal ischemia/hypoxia; tubulointerstitial space; renal vascular system
Injury and recovery responses of ARF have been widely studied using rodent models of ischemia-reperfusion (I/R) injury to the kidney (18, 29, 41, 48). Morphologically, outer-medullary proximal tubules are the primary sites of injury in this setting. Proximal tubules have a remarkable ability to be repaired following injurious stimuli, and this process is thought to contribute to the functional recovery from ARF (18, 29, 41, 48).
Similarly, injury to the renal vascular system has also received considerable attention (44). Damage to the renal vascular endothelium results in altered regulation of vascular tone (12, 13, 43). Decreased blood flow in the peritubular capillaries has been demonstrated within minutes of reperfusion injury (50). The importance of endothelial dysfunction in the genesis of I/R injury is further suggested by the observation that administration of endothelial nitric oxide synthase-expressing endothelial cells will hone to the kidney and preserve blood flow and loss of function following I/R (7). In addition, damage to the renal vasculature results in a breakdown of barrier function and increased adhesiveness to inflammatory cells, both of which appear to contribute to the loss of function following ischemic injury (37, 4345).
Recent studies from our laboratory demonstrated that there is a reduction in renal vascular density following I/R injury (3). Interestingly, unlike the renal tubular system, our results suggest that the renal vascular system has little regenerative capacity because vessel number was reduced for up to 40 wk following recovery (3). Furthermore, we showed that postischemic kidneys are more hypoxic than kidneys from shamoperated animals and suggested that the loss of vessel density may predispose secondary complications of chronic renal disease following recovery from ARF (2).
The mechanisms mediating vessel dropout following renal ischemia are largely unexplored. It is clear that there are direct effects of ischemia on cytoskeletal integrity and tight junction stability (45). However, it is also possible that antiangiogenic humoral factors are synthesized in the kidney that may compromise vessel integrity following ischemic injury.
One such potential factor is angiostatin, which inhibits tumor angiogenesis and metastasis and contributes to apoptosis of endothelial cells in vitro (33). Angiostatin is produced via processing of plasminogen by matrix metalloproteinases (MMP)-2, -7, and -9 and potentially other enzymes (25, 34, 35). Angiostatin has recently been identified in myocardial interstitial fluid of dogs undergoing repetitive left anterior descending artery occlusion and also treated with NG-nitro-L-arginine methyl ester; under these conditions, typical collateral development is impaired despite the increased production of vascular endothelial cell growth factor (VEGF) (26).
To date, there have been no studies characterizing the presence of angiostatin in the kidney. In addition, it is clear that there is a breakdown of the extracellular matrix (ECM) of the tubular basement membrane following renal ischemia (47). Despite this fact, there are few studies that addressed the potential alterations in renal MMP activity following ischemic injury. The association between MMP activity, angiostatin, and the potential to mediate blood vessel loss led us to investigate the expression of angiostatin and MMPs in the setting of renal I/R injury. Our results demonstrate that MMP expression is induced and that angiostatin is generated in response to I/R injury. These factors may have implications in the overall balance of renal vessel density and/or the balance of renal ECM content following ischemic injury.
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METHODS |
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Male Sprague-Dawley rats (Harlan, Madison, WI; 250 g) were housed in pairs in standard shoe-box cages with a 12-h light cycle (lights on 06001800) and access to water and standard laboratory rat chow (0.8% NaCl, Purina) available ad libitum. Animals were anesthetized with ketamine (100 mg/kg ip) for 10 min followed by administration of pentobarbital sodium (2550 mg/kg ip). ARF was induced by performing bilateral renal artery clamping for 45 min followed by reperfusion according to procedures previously described (14). After ischemia, reflow was verified visually and animals were allowed to recover for 1, 2, 3, 8, or 35 days postinjury; sham-operated control animals were treated by exposing the kidneys under identical anesthesia but not clamping. Sham-operated controls were performed in the 3-, 8-, and 35-day groups.
Measurement of renal function and harvesting of tissue. Renal functional parameters were measured at 24 h, 7 days, and 35 days. Tail blood samples (0.5 ml) were collected under light halothane anesthesia into heparinized tubes, and plasma was obtained following centrifugation. Urine collection was for 24 h in metabolic cages (Nalgene). Serum and urine creatinine were determined using standard assays (Sigma creatinine kit 555A). Urine volume was determined gravimetrically.
At the time of death, animals were deeply anesthetized with ketamine/xylazine/acepromazine (2.0/0.6/0.3 mg/kg). The kidneys were removed and cut bilaterally; one-half of the kidney was snapfrozen in liquid nitrogen and stored at -70° C for subsequent biochemical analysis. In some studies, tissue was fixed by immersion in Histochoice (Sigma) for subsequent immunofluorescence.
Biochemical analysis of angiostatin and MMPs. Protein extracts from frozen kidneys were prepared according to the method described by Rankin et al. (38). Briefly, kidneys were homogenized in 50 mM Tris·HCl, pH 8.0, containing 10% DMSO, 10 mM E64, and 2 mM PMSF followed by centrifugation at 3,000 g for 5 min. The supernatant protein content was determined with the Bio-Rad protein assay kit using the microassay format based on the manufacturer's recommendation for enhanced sensitivity.
For kidney samples, 20100 µg of total protein (see figure legends) were combined with standard electrophoresis buffer (10 mM Tris, pH 8.3, containing 10% SDS and 700 mM -mercaptoethanol). After being heated for 5 min, samples were electrophoresed on a 12.5% (angiostatin) or 10% (MMP) SDS-PAGE gel and transferred to nitrocellulose. To verify efficiency of transfer and equal loading, the membranes were prestained with Ponceau S (Sigma). After being blocked, the membranes were incubated for 48 h at 4° C with either of the following antibodies: goat anti-human angiostatin (0.2 µg/ml; R & D Systems) or mouse anti-MMP-2 and -MMP-9 (0.5 µg/ml each; Santa Cruz Biotechnology). Peroxidase-conjugated anti-mouse IgG (Bio-Rad) or anti-goat IgG (Sigma) was used with chemiluminescence (SuperSignal, Pierce) for development. An additional primary antibody used in these studies was a peroxidase-conjugated goat anti-human plasminogen (10 µg/µl; Novus Biologicals), which does not require subsequent incubation in secondary antibody before development.
Purified human angiostatin (Alpha Diagnostic International, San Antonio, TX), human plasminogen (Sigma), and human plasmin (Sigma) were included in several gels as positive controls and were used to identify different molecular mass immunoreactive products.
Lysine-Sepharose enrichment. Washed lysine-Sepharose beads (Pharmacia) equivalent to 5 mg of dry gel were combined with 2 mg of renal protein extract or 15 µl of rat plasma in a volume of 1 ml complete RIPA buffer (10 mM HEPES, pH 7.5, 142.5 mM KCl, 0.2% NP-40 supplemented with protease inhibitor cocktail tablets; Roche). In some studies, urine equivalent to 100 µg creatinine or 500 µg urine protein was treated in the same manner. All samples were incubated overnight at 4° C while being shaken. The beads were subsequently washed by consecutive centrifugation and resuspension twice in complete RIPA and twice in RIPA lacking NP-40. After the final spin, adherent proteins were eluted for Western blot analysis by adding 15 µl of electrophoresis sample buffer and boiling the beads for 5 min.
In other studies, adherent proteins were eluted from lysine-Sepharose beads in 200 µl of 0.2 M -aminocaproic acid (Sigma); the buffer was changed to 0.1 M phosphate buffer, pH 7.2, using Sephadex G-25 spin columns (Pharmacia), and samples were immediately frozen at -80° C until use in in vitro biological assays.
In vitro endothelial proliferation assay. Proliferation was detected by direct assessment of cell numbers similar to the method described previously (26). Cultures of confluent human aortic endothelial cells (EC) were trypsinized, and the yield of cells was measured using a hemocytometer. Ten thousand cells were seeded in 10-mm wells (24-well plate) and maintained in DMEM/5% FBS overnight. The cells were then serum starved for 72 h (0.5% FBS) to produce growth arrest and synchronize the cell cycle. After 72 h, EC were maintained in 0.5% FBS/DMEM to establish a baseline. Cells were then stimulated with VEGF (50 ng/ml) and coincubated with or without lysine-Sepharose extracts obtained from kidney or urine samples. The extracts added to these cultures corresponded to either 1.5 µg/µl of kidney lysate protein or 0.5 µg/µl of urine protein, as the original starting material before processing with lysine-Sepharose beads. In some wells, a neutralizing angiostatin antibody was also included in the incubation (4 µg/ml; Pharmingen).
RT-PCR. Total RNA from the kidney or liver was obtained from frozen tissue using Ultraspex RNA isolation solution (Biotecx, Houston, TX). The presence of plasminogen mRNA was determined by running reverse-transcription PCR. Two micrograms of RNA were reverse transcribed with Superscript (Life Technologies) and oligo dT primers for 30 min at 39° C according to the manufacturer's instructions. The resultant RT product was then amplified by PCR using Taq polymerase (Amersham) and the following primers: forward primer 5'-GGATCCATTTAGGTGACACTATAGAACCAGACTGGTC-3'; reverse primer 5'-GAATTCAATCGACTCACTATAGGAGCAAAGAGCAGCAG-3'. The underlined sequences refer to the promoter sequence for SP6 and T7 polymerase, respectively, a feature that was not used in the current study. The resultant amplified products were run on a 1% agarose gel and were consistent with the predicted 404-base region of the rat plasminogen mRNA. As a positive control, primers for -actin were included in the reaction mixture as described previously (49).
SDS-PAGE zymography. Gelatinase activity was evaluated by SDS-PAGE zymography as described previously (26). Protein samples (200 µg) from renal extracts were loaded into the wells of a 10% gelatin gel (Bio-Rad) and electrophoresed. The gel was removed and incubated for 1 h at room temperature in 100 ml of renaturing buffer (2.5% Triton X-100) on a rotary shaker. The buffer was replaced with 100 ml of development buffer (50 mM Tris, pH 7.5, 200 mM NaCl, 5 mM CaCl2, and 0.02% Briji-35). The gel was incubated at 37° C for 1824 h. Each gel was stained with 0.5% Coomassie blue G-250 in 30% methanol and 10% acetic acid for 3 h and destained with several changes in 30% methonal/10% acetic acid. The gels were digitized using a CCD camera frame digitizer system and analyzed using NIH Image (density and band area). Positive controls for MMP-2 and MMP-9 were obtained from Calbiochem.
Immunofluorescence for MMP-2 and MMP-9. Histochoice-fixed kidneys were processed by incubation in 10 mM phosphate-buffered saline, pH 7.4, and subsequent daily changes of increasing sucrose (5, 10, and 20%) in the same buffer. Cryostat sections (10 µm) were thaw-mounted and stored desiccated at 4° C until use. The tissues were rehydrated in PBS and blocked in blocking buffer (PBS containing 0.2% Triton X-100, 5% normal sheep serum, and 0.6% bovin serum albumin). Goat anti-MMP-2 or -MMP-9 (10 µg/ml; Santa Cruz Biotechnology) was applied for 18 h at 4° C. Nonimmune goat IgG (Sigma) at the same concentration was used as a control for specificity. After being rinsed, the tissue was incubated with anti-goat-FITC (Pierce) and visualized using a Nikon E600 microscope equipped with Xenon-Arc epifluorescence and a Princeton Instruments Micromax Cooled CCD camera (RS Princeton Instruments, Trenton, NJ). Image acquisition was aided by a Pentium III grade personal computer running MetaFluor imaging software (Universal Imaging version 4.6, Downingtown, PA). Signals were obtained by excitation at 490 nm and an emission filter of 535/540. Tissue autofluorescence was determined using excitation at 405 nm with the same emission filter and exposure time. A FITC-specific signal was then determined by subtracting the signal obtained at 490-nm excitation from the one obtained at 405-nm excitation using MetaFluor imaging software. Both darkfield and brightfield images were then obtained, and these images were overlaid onto the FITC-specific image for contrast.
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RESULTS |
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A standard Western blot approach using an antibody that recognizes kringle 13 of angiostatin was used to identify plasminogen/angiostatin in renal protein extracts. The antibody effectively recognized proteins consistent with the size of full-length plasminogen (93 kDa) and plasmin (
62 kDa; Fig. 1A). The commonly identified 50- and 38-kDa forms of angiostatin, previously identified by us in canine myocardial interstitial fluid (26), were not consistently observed using this standard approach (Fig. 1A).
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Plasminogen-derived products have high affinity for lysine, and this characteristic has been used to aid in its purification (8, 9, 22). Therefore, renal protein extracts were incubated in the presence of lysine-Sepharose before electrophoresis and Western blot analysis. Ponceau S staining revealed that only a small amount of protein remains following lysine-Sepharose enrichment of up to 2 mg renal protein extract (Fig. 1, D and E). Immunoreactive bands corresponding to plasminogen and plasmin, as well as lower molecular mass species that are consistent with the reported size of angiostatin, were reliably and consistently detectable following lysine-Sepharose enrichment (Fig. 1, B and B'). The relative amount of all plasminogen-derived products, including those corresponding to the 50- and 38-kDa forms of angiostatin, was considerably higher in extracts in the early (days 1-3) postischemic period compared with sham-operated controls. The apparent levels of these proteins diminished over time; however, longer exposure of the gels demonstrated that the 50- and 38-kDa immunoreactive products were persistently elevated for as long 8 and 35 days postischemia when compared with controls (Fig. 1B').
As further verification of these results, lysine-enriched renal protein extracts were probed with a different antibody termed antiplasminogen, which is conjugated directly to peroxidase. The results using this antibody were qualitatively similar to those obtained with the antiangiostatin antibody (Fig. 1, C and C').
Lysine-Sepharose-enriched rat plasma was also probed with either antiangiostatin or antiplasminogen (Fig. 2). At 8 and 35 days postischemia, there was a steady-state increase in all plasminogen-reactive species. The 38-kDa form of plasminogen was largely undetectable in plasma of sham-operated rats but was persistently detected in postischemic rats (Fig. 2, top). Both plasmin and the 50-kDa form of plasminogen were upregulated in postischemic samples. The antiplasminogen antibody revealed qualitatively similar results but did not consistently detect the 38-kDa form of angiostatin (Fig. 2, bottom).
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Lysine-Sepharose enrichment was also used to analyze plasminogen/angiostatin content in the urine of postischemic rats. Samples for Western blots were normalized to either creatinine content (Fig. 3, top) or protein content (Fig. 3, bottom). Regardless of the method of normalization, there was little evidence of immunreactive products in urine from shamoperated control rats. Postischemic animals demonstrated largely elevated urinary plasminogen/angiostatin immunoreactivity, although the pattern of detectable immunoreactive products was inconsistent. However, postischemic urine samples at 2, 8, and 35 days postsurgery tended to contain detectable levels of the 50- and 38-kDa angiostatin isoforms.
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As an additional verification of presence of angiostatin in kidney and urine, lysine-Sepharose-enriched fractions were analyzed for their ability to inhibit VEGF-induced angiogenesis in vitro. VEGF stimulation (50 ng/ml) resulted in a 90% increase in EC number; lysine-Sepharose extracts completely inhibited the VEGF-induced stimulation of EC number (Fig. 4). Under the conditions of the assay used, the inhibition of VEGF-induced stimulation was maximal and identical in extracts derived from either sham-operated or postischemic animals. Data in Fig. 4 represent pooled data from both shamoperated and postischemic kidney and urine extracts. Coincubation of lysine-Sepharose extracts with a neutralizing angiostatin antibody restored VEGF-induced cell proliferation (Fig. 4). These data suggest that the lysine-Sepharose-enriched fraction from urine and kidney homogenates contain biologically active angiostatin.
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Plasminogen is synthesized primarily in the liver, although extrahepatic synthesis of plasminogen has been reported (46). RT-PCR was used to determine if plasminogen synthesis might occur in the kidney following injury. Figure 5 demonstrates that the predicted amplicon corresponding to plasminogen mRNA was generated following RT-PCR of total liver RNA. This product was not detectable by RT-PCR of total RNA from kidney of either sham-operated or postischemic animals. These data suggest that the source of plasminogen is not renal and is likely hepatic.
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It is likely that increased angiostatin expression is the result of increased local conversion of plasminogen within the kidney following I/R injury. Therefore, we sought to determine whether MMP-2 and MMP-9, two enzymes reported to convert plasminogen to angiostatin, were altered in the setting of renal ischemic injury. Western blot analysis (Fig. 6A) and zymography (Fig. 6B) demonstrate the increase in MMP-2 and MMP-9 protein and activity, respectively, in extracts obtained from rat kidney 1, 2, and 3 days postischemia. Densitometric analyses of Western blots revealed significant increases ranging from two- to threefold in both MMP-2 and MMP-9 compared with sham-operated controls (P < 0.05, n = 4/group/time point). Similarly, there were significant increases postischemia in the activities of both MMP-2 and MMP-9. Densitometric values from zymography ranged from two- to fourfold between days 1 and 3 for MMP-2 (P < 0.05 for each time point). Values for MMP-9 activity were not different on day 1, but two- to threefold higher on days 2 and 3 (P < 0.05).
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Immunofluorescence was used to identify the site of increased MMP-2 and -9 expression in tissues of rats 3 days following ischemic injury. In sham-operated controls, fluorescent signal corresponding to MMP-2-like immunoreactivity could be distinguished near the basolateral aspect of tubular structures in the cortex (arrow, Fig. 7A) and this pattern was more prominent in the outer medulla (arrow, Fig. 7B). In contrast, no signal was obtained in sham sections using an antibody for MMP-9 (Fig. 8, A and B) or a nonimmune antibody (not shown). Immunofluorescent signal for MMP-2 was increased moderately in the cortex (Fig. 7C) and more dramatically in the outer medulla (Fig. 7D) 3 days following ischemic injury. Higher-magnification images illustrate that MMP-2 immunoreactivity was primarily localized to the basolateral aspect of the tubule and in the tubulointerstial area (Fig. 7E). When visualized relative to an image obtained from brightfield microscopy, MMP-2 signal can be seen over the basement membrane (arrow, Fig. 7F) and outside the tubule in the interstitial space.
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MMP-9 immunoreactivity was also apparent in the cortex and medulla of postischemic animals, although the pattern of expression was different. Although some interstitial staining could be observed, MMP-9 immunoreactivity was present to a significant degree within cells of the tubular epithelium (arrowhead, Fig. 8, C-E) and in cells residing in the tubulointerstitial area (arrow, 8, C-E). Figure 8F illustrates the lack of signal obtained by a nonimmune goat IgG in the outer medulla of a postischemic rat.
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DISCUSSION |
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We suggest that in the setting of I/R injury, there is a shift in the balance between trophic, angiogenic factors and antiangiogenic factors. Such alterations in the humoral milieu of the postischemic kidney would result in the loss of renal vessels and/or the inhibition of an endogenous repair response, resulting in a net decrease in total vessel number. There are many potential angiogenic and antiangiogenic factors that may be influenced in the setting of ischemic ARF. Although the current study focused on angiostatin, the characterization of several factors with the potential to modulate vessel density in the setting of ischemic injury may likely be warranted in future studies.
Plasminogen produced in the liver has long been recognized as the parent protein of plasmin, by either tissue-type or urokinase-type plasminogen activator. Plasmin degrades plasma fibrin and ECM components either directly or indirectly through the activation of MMPs (39). Plasminogen is also processed by a number of different enzymes, including, but limited to MMP-2, and MMP-9, to generate angiostatin, a molecule with antiangiogenic activity that comprises the kringle 14 domains (K14) of the parent molecule (25, 3335). The K14 angiostatin was originally identified as a 38-kDa peptide (33); however, angiostatin has also been identified as a 45- to 50-kDa form. In a previous study, we identified bands of 50 and 38 kDa in canine myocardial interstitial fluid (26). In this study, bands of
38 and 50 kDa were also identified in postischemic rat kidneys. The possibility that these bands correspond to angiostatin is supported by their physicochemical lysine affinity and the use of two different antibodies against angiostatin/plasminogen. Moreover, lysine-Sepharose extracts suppressed VEGF-induced proliferation of EC cells; an effect that was blocked using a third unique antibody against angiostatin. Although not the focus of this study, it is worth noting that there was a substantial increase in immunoreactivity of two bands
65 kDa, which likely corresponds to plasmin.
Renal gene expression is altered dramatically postischemia. RT-PCR results argue against the possibility that plasminogen is produced locally by injured kidney and suggested that local conversion of angiostatin occurs. The precise localization of angiostatin could not be determined because the antibodies that were used to identify angiostatin also recognize other plasminogen-derived products. However, the analysis of MMP-2 and -9, which are two of many possible enzymes that may mediate cleavage of plasminogen to angiostatin, allows speculation on the possible site of production of angiostatin. Thus it is of interest that the majority of MMP-2 signal appears to be either near the tubular basement membrane or the interstitial space, while MMP-9 is present in tubular cells as well as interstitial cells. The identity of these interstitial cells is not known, but they likely represent an inflammatory cell type. In one previous study, Ziswiler et al. (51) reported that there was no change in the MMP-2 or -9 activities in renal cortex at either 12- or 24-h postischemia in rats. The apparent disparity may relate to the fact that we used whole kidney homogenates containing the outer medullary region. Our immunofluorescence studies suggest the strongest intensities of the MMP-2 and -9 signals were in the outer medulla (Figs. 7 and 8).
The localization of MMPs within the tubulointerstitial area is of significant interest with regard to their potential role in angiostatin generation because this region is the location of peritubular capillaries. Thus the potential local activation of angiostatin by MMPs in close proximity to blood vessels may result in a negative influence on capillary stability. It should also be emphasized that a number of other MMPs can lead to angiostatin generation such as MMP-3 (stomelysin-1), MMP-12 (matrix metalloelastase), and MMP-7 (14, 25, 3335). These have not been addressed in these studies. Moreover, plasminogen activator inhibitor-1 and transforming growth factor- are also capable of affecting angiostatin generation (32), and these are both enhanced in the setting of ischemic ARF in rats (4).
The localization of MMPs in the tubulointerstitial area suggests that it may also influence the regulation of ECM homeostasis in the tubular basement membrane following I/R injury. To our knowledge, no studies evaluated the ramifications of MMP activity on the long-term structure of the tubular basement membrane following ischemic injury. Clearly, further studies will be required to determine whether MMPs influence ECM control and/or angiostatin generation in this setting.
The results from these studies raise the possibility that angiostatin, and potentially other antiangiogenic factors, predisposes vessel dropout and/or inhibits compensatory angiogenic activity post-I/R. With regard to the latter possibility, angiostatin-like proteins were identified in extracts, albeit at reduced levels, for up to 35 days postinjury. It is of interest that elevated angiostatin levels were also identified in plasma. Recent studies showing decreased barrier function of peritubular capillaries following I/R (45) suggest the possibility that plasminogen can leak from the vasculature into the interstitial space where it may be acted on by MMPs. Such a breakdown in barrier function also opens the possibility of backleak into the vascular system, which may account for presence of angiostatin in the plasma. In addition, administration of radiolabeled angiostatin is found in kidney and urine, suggesting that it is cleared by the kidney (28). Therefore, reduced clearance of angiostatin by postischemic kidneys may also account for elevated plasma angiostatin levels. However, determination of these putative mechanisms would require further careful investigation. Whether plasma angiostatin affects other tissues following I/R is unknown.
It is also of interest that some angiostatin was detectable in the urine of postischemic animals, although the pattern of immunoreactive products in urine was not consistent. We suggest that heterogeneity in the reestablishment of the tubular epithelial barrier as well as the level of microalbuminuria may create a very complicated set of parameters that affect the detection of angiostatin in the urine. Nevertheless, it is possible that urinary angiostatin may represent a potentially interesting marker of renal injury in the setting of ARF or delayed graft function.
In this model of reversible ARF, there is a net reduction in the total number of renal vessels, suggesting a shift in the balance between antiangiogenic factors and proangiogenic factors. In the current study, we identified the generation and sustained expression of angiostatin, a well-known antiangiogenic factor, following renal I/R injury. Further studies will be required to determine whether this antiangiogenic factor participates in blood vessel homeostasis and whether angiostatin may also be present in kidney of other progressive disorders characterized by a decrease in blood vessel loss.
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ACKNOWLEDGMENTS |
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GRANTS
D. P. Basile was supported by National Institutes of Health Grants DK-54350 and DK-63114, and W. M. Chilian was supported by Grants HL-32788 and HL-65203.
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FOOTNOTES |
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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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REFERENCES |
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