点击显示 收起
【摘要】 Ischemic injury to the kidney results in blood vessel loss and predisposition to chronic renal disease. Angiostatin is a proteolytic cleavage product of plasminogen that inhibits angiogenesis, promotes apoptosis of endothelial cells, and disrupts capillary integrity. A combination of lysine-Sepharose enrichment followed by Western blotting was used to study the expression of angiostatin in response to the induction of ischemic renal injury. No angiostatin products were readily detectable in kidneys of sham-operated control rats. In contrast, both 38- and 50-kDa forms of angiostatin were dramatically enhanced in the first 3 days following 45-min ischemia-reperfusion injury. Renal angiostatin levels declined but remained detectable at late time points postrecovery (8-35 days postischemia). Angiostatin-like immunoreactivity was also elevated in the plasma and in urine for up to 35 days following injury. Lysine-Sepharose extracts of either kidney or urine inhibited vascular endothelial cell growth factor-induced proliferation of human aortic endothelial cells in vitro; an effect that was blocked by coincubation with an angiostatin antibody. RT-PCR verified that mRNA of the parent protein plasminogen was produced in the liver, but it was not present in either sham-operated or postischemic kidney. Matrix metalloproteinase (MMP)-2 and MMP-9, which may mediate angiostatin generation, were enhanced in postischemic kidney tissue and were localized to the renal tubules, interstitial cells, and the tubulo-interstitial space. These data indicate the possible local synthesis of angiostatin following acute renal failure (ARF) and suggest a possible role for MMPs in this activity. Renal angiostatin generation following ARF may modulate renal capillary density postischemia and thereby influence chronic renal function.
【关键词】 renal ischemia/hypoxia tubulointerstitial space renal vascular system
RENAL ISCHEMIA / HYPOXIA is a leading cause of acute renal failure (ARF); a clinical condition associated with rapid loss of renal function and high mortality rates. ARF is largely reversible and, in surviving patients, the restoration of renal function is generally observed. However, incomplete recovery and/or a secondary decline of renal function have been reported ( 1, 5, 6, 15, 23, 24, 36, 42 ). Moreover, ischemic injury in the setting of renal transplantation predisposes the graft to the development of future renal complications ( 16, 17, 27, 31 ).
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, 43 - 45 ).
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 N G -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.
METHODS
Animal and surgical procedures. Care of the rats before and during the experimental procedures was conducted in accordance with the policies of the Animal Resource Center, Medical College of Wisconsin, and the National Institutes of Health (NIH) Guide for the Care and Use of Laboratory Animals. All protocols had received prior approval by the Medical College of Wisconsin Institutional Animal Care and Use Committee.
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 0600-1800) 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 (25-50 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, 20-100 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'-GGATCC ATTTAGGTGACACTATAGAA CCAGACTGGTC-3'; reverse primer 5'-GAATTC AATCGACTCACTATAGG AGCAAAGAGCAGCAG-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 CaCl 2, and 0.02% Briji-35). The gel was incubated at 37° C for 18-24 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.
RESULTS
Rats were subjected to 52 min of renal ischemia and allowed to recover for 1, 2, 3, 8, or 35 days postsurgery. Mean serum creatinine values at 24-h postreperfusion were between 3.2 ± 0.3 and 3.7 ± 0.2 mg/dl, indicating evidence of injury ( Table 1 ). Animals that were allowed to recover for 8 days or longer demonstrated a return of serum creatinine to sham-operated control values. In animals surviving for 35 days, daily urinary output remained significantly elevated vs. sham-operated controls, consistent with our previous studies ( 2, 3 ) ( Table 2 ).
Table 1. Renal functional data following ischemiareperfusion or sham operation
Table 2. Renal function of group following recovery from surgery for 35 days
A standard Western blot approach using an antibody that recognizes kringle 1-3 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. 1 A ). 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. 1 A ).
Fig. 1. Angiostatin/plasminogen expression in kidney following ischemia-reperfusion (I/R) injury. A : standard Western blot analysis using an antiangiostatin antibody (R & D Systems) against 100 µg protein from total kidney homogenates. Kidney homogenates were obtained from postischemic rats at the indicated times and from sham-operated rats at 3 days (d) postischemia. Both plasminogen and plasmin bands are easily distinguishable, whereas lower molecular mass species are not consistently distinguished. B : Western blot analysis using the same antiangiostatin antibody following enrichment of 2 mg protein with lysine-Sepharose. Additional bands of 38 and 50 kDa, consistent with the reported size of angiostatin, are distinguishable in postischemic samples 1, 2, or 3 days following I/R injury. B ': longer exposure of the same blot as in B, in which the 38- and 50-kDa bands are distinguishable in samples 8 and 35 days following I/R injury. C and C ': same as B and B ', except blot was probed with antiplasminogen-horseradish peroxidase (HRP)-conjugated antibody (Novus). Similarly sized fragments were detected using this different antibody. D - F : Ponceau S-stained nitrocellulose membranes of the blots probed in A - C, respectively, demonstrating comparable amount of protein loading. Only 1 band was distinguishable in lysine-Sepharose-treated extracts, which did not correspond in size to any immunoreactive product and was not regulated by I/R injury. Results are representative of a minimum of 4 samples/time point. Quantitative densitometric analyses were not conducted because no signal corresponding to angiostatin-like fragments was consistently observed in samples from sham animals.
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. 1 B ' ).
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 ).
Fig. 2. Angiostatin/plasminogen detection in plasma following I/R injury. Top : 15 µl of rat plasma were subjected to lysine-Sepharose enrichment and probed using either the antiangiostatin antibody or antiplasminogen HRP antibody ( bottom ). Shown are samples obtained from 4 sham-operated rats (8 days) and from 4 postischemic rats at 8 and 35 days post-I/R.
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.
Fig. 3. Angiostatin/plasminogen detection in urine following I/R injury. Urine samples were collected for 24 h in metabolic cages from sham-operated and postischemic animals. Aliquots of either 100 µg creatinine ( top ) or 500 µg of urinary protein ( bottom ) were subjected to lysine-Sepharose enrichment and Western blot analysis. Samples were collected from postischemic animals and sham-operated animals at the indicated times; urinary levels from sham-operated controls are shown from 35-day samples. However, these did not differ as a function of time following sham surgery (data not shown). Results shown are those using the R and D antibody. Positive controls shown in standard (std) lane in this blot correspond to purified human plasminogen, plasmin, and angiostatin, as marked.
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.
Fig. 4. Inhibition of endothelial cells (EC) proliferation by lysine-Sepharose-enriched kidney and urine extracts. Human aortic EC were cultured as described in METHODS and stimulated with vascular endothelial cell growth factor (VEGF), with or without lysine-Sepharose-enriched fractions from kidney and urine as shown. Data are means ± SD and are expressed as percent increase in cell number relative to 0.5% FBS control. Positive control (VEGF) is based on n = 8 and VEGF + antibody, n = 4. Data obtained from tissue and urine extracts correspond to the mean values obtained from 4 different animals. As extracts from sham-operated and postischemic animals were both maximally inhibitory, the data were pooled (i.e., 2 sham and 2 postischemic; each extract tested in quadruplicate). * P < 0.05 vs. 0.5% FBS control. P < 0.05 vs. VEGF stimulated. P < 0.05 vs. corresponding antibody-treated samples.
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.
Fig. 5. Plasminogen mRNA is not detectable in kidney by RT-PCR. Primers were designed to amplify mRNA of plasminogen or -actin; 1 µg of total RNA of whole kidney or liver (as labeled) was subjected to RT-PCR as described in METHODS. A minimum of 4 sham and postischemic kidneys (3 days) were tested and failed to demonstrate an amplified product corresponding to plasminogen mRNA in kidney.
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. 6 A ) and zymography ( Fig. 6 B ) 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).
Fig. 6. Expression of matrix metalloproteinase (MMP)-2 and MMP-9 in kidney in response to I/R injury. A : DMSO protein extracts (20 µg) from kidneys of sham-operated (sh) rats or rats 1, 2, or 3 days postischemia were subjected to PAGE, transferred to nylon membranes, and probed with antibodies against MMP-2 ( 65 kDa; top ) or MMP-9 ( 90 kDa). B : gelatin zymography of DMSO protein extracts (50 µg) from kidneys of sham-operated rats or rats 1, 2, or 3 days postischemia. Human MMP-2 (M-2) and MMP-9 (M-9) standards are shown in lanes 1 and 2.
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. 7 A ) and this pattern was more prominent in the outer medulla (arrow, Fig. 7 B ). 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. 7 C ) and more dramatically in the outer medulla ( Fig. 7 D ) 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. 7 E ). When visualized relative to an image obtained from brightfield microscopy, MMP-2 signal can be seen over the basement membrane (arrow, Fig. 7 F ) and outside the tubule in the interstitial space.
Fig. 7. Immunofluorescent localization of MMP-2 in normal and postischemic rat kidneys. Shown are representative frozen sections (10 µm) through the renal cortex ( A, C ) and outer medulla ( B, D, E, F ) of shamoperated rats ( A, B ) or of rats 3 days postischemia ( D - F ). Sections were stained with antibodies against MMP-2 and appropriate FITC-labeled secondary antibody. FITC-specific fluorescence (see METHODS ) was labeled green and is shown relative to the image obtained from darkfield to visualize structure (shown in blue). There is modest MMP-2-immunofluorescent staining in the basolateral aspect of tubules from sham-operated rats ( A, B, arrows). MMP-2-like immunoreactivity is enhanced slightly in cortex ( C ) and more appreciably in the renal medulla ( D ) 3 days following renal injury. E : higher magnification of D, illustrating the localization of MMP-2 in the interstitial space (arrow). F : FITC signal from E is shown relative to image obtained from bright-field microscopy (blue); the basement membrane can be observed as a thin line (arrow). FITC signal can be appreciated over the basement membrane as well as outside the basement membrane area. Bar in A refers to A - D, and bar in E refers to E and F.
Fig. 8. Immunofluorescent localization of MMP-9 normal and postischemic rat kidneys. Shown are representative frozen sections (10 µm) through the renal cortex ( A, C ) and outer medulla ( B, D, E, F ) of sham-operated rats ( A, B ) or of rats 3 days postischemia ( D - F ). Sections were stained with antibodies against MMP-9 and appropriate FITC-labeled secondary antibody. FITC-specific fluorescence (see METHODS ) was labeled green and is shown relative to the image obtained from darkfield to visualize structure (shown in blue). MMP-9 signal was not appreciable in cortex or outer medulla from sham-operated rats. FITC signal was detectable in cortex ( C ) and outer medulla ( D ) of postischemic animals. MMP-9 signal was observed within tubular cells (arrowhead, C, D ), as well as within in cells in the interstitial area (arrow). E : higher magnification of the area highlighted in D using a brightfield image as contrast. FITC-positive signal is observed within the tubule (arrowhead) and in cells outside the basement membrane (arrowhead). No specific FITC signal was apparent in kidneys of postischemic rats following incubation with goat IgG at a concentration equivalent to that used for MMP-2 and MMP9 ( F ). Bar in E refers to E alone; bar in F refers to A - D and F.
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 8 F illustrates the lack of signal obtained by a nonimmune goat IgG in the outer medulla of a postischemic rat.
DISCUSSION
The potential that the kidney may generate molecules with antiangiogenic activity has, to date, received little attention and was the major focus of this study. It is now well-recognized that renal vessel loss is a common feature of several models characterized by tubulointerstitial disease and a predisposition to hypertension ( 10, 19 - 21, 30, 40 ). Similarly, we showed that there is a permanent reduction in the density of renal blood vessels following the apparent recovery from I/R injury ( 3 ). In addition, evidence of peritubular capillary loss has been suggested from analysis of biopsies of patients with tubulointerstitial disease of many etiologies ( 11 ). Hence, it would appear that a complete understanding of the factors involved in maintaining vessel integrity might be useful to our understanding of the pathogenesis of renal diseases.
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 1-4 domains (K1-4) of the parent molecule ( 25, 33 - 35 ). The K1-4 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, 33 - 35 ). 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.
ACKNOWLEDGMENTS
The authors thank D. Donohoe and M. Moniz for technical assistance during these studies and G. Slocum for assistance with fluorescence microscopy.
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.
【参考文献】
Alon US. Neonatal acute renal failure: the need for long-term follow-up. Clin Pediatr (Phila) 37: 387-390, 1998.
Basile D, Donohoe DL, Roethe K, and Mattson DL. Chronic renal hypoxia following ischemia/reperfusion injury: effects of L -arginine on hypoxia and secondary damage. Am J Physiol Renal Physiol 284: F338-F348, 2003.
Basile DP, Donohoe D, Roethe K, and Osborn J. Renal ischemic injury results in permanent damage to peritubular capillaries and influences long-term function. Am J Physiol Renal Physiol 281: F887-F899, 2001.
Basile DP, Martin DR, and Hammerman MH. Extracellular matrix related genes in kidney post-ischemic injury: potential role for TGF- in repair. Am J Physiol Renal Physiol 275: F894-F903, 1998.
Bonomini V, Stefoni S, and Vagelista A. Long-term patient and renal prognosis in acute renal failure. Nephron 36: 169-172, 1984.
Briggs J, Kennedy A, Young L, Luke R, and Gray M. Renal function after acute tubular necrosis. Br Med J 3: 513-516, 1967.
Brodsky SV, Yamamoto T, Tada T, Kim B, Chen J, Kajiya F, and Goligorsky MS. Endothelial dysfunction in ischemic acute renal failure: rescue by transplanted endothelial cells. Am J Physiol Renal Physiol 282: F1140-F1149, 2002.
Cao Y, Veitonmaki N, Keough K, Cheng H, Lee LS, Cao Y, and Zurakowski D. Elevated levels of urine angiostatin and plasminogen/plasmin in cancer patients. Int J Mol Med 5: 547-551, 2000.
Catstellino FJ and Powell JR. Human plasminogen. Methods Enzymol 80: 365-378, 1981.
Choi YJ, Baranska-Daca E, Barrios R, and Nguyen V. Peritubular capillary loss mediates tubular loss and interstitial fibrosis (Absract). J Am Soc Nephrol 10: 70A, 1999.
Choi YJ, Chakraborty S, Nguyen V, Nguyen C, Kim BK, Shim SI, Suki WN, and Truong LD. Peritubular capillary loss is associated with chronic tubulointerstitial injury in human kidney: altered expression of vascular endothelial growth factor. Hum Pathol 31: 1491-1497, 2000.
Conger JD, Robinette JB, and Hammond WS. Differences in vascular reactivity in models of ischemic acute renal failure. Kidney Int 39: 1087-1097, 1991.
Conger JD and Weil JV. Abnormal vascular function following ischemia-reperfusion injury. J Investig Med 43: 431-442, 1995.
Cornelius LA, Nehring LC, Harding E, Bolanowski M, Welgus HG, Kobayashi DK, Pierce RA, and Shapiro SD. Matrix metalloproteinases generate angiostatin: effects on neovascularization. J Immunol 161: 6845-6852, 1998.
Finn WF. Recovery from acute renal failure. In: Acute Renal Failure, edited by Lazarus JM and Brenner B. New York: Churchill Livingstone, 1993, p. 553-596.
Fontan MP, Rodriguez-Carmona A, Bouza P, and Valdes F. The prognostic significance of acute renal failure after renal transplantation in patients treated with cyclosporin. QJM 91: 27-40, 1998.
Hetzel G, Klein B, Brouse M, Westhoff A, Willers R, Sandmann W, and Grabensee B. Risk factors for delayed graft function after renal tranplantation and their significance for long-term clinical outcome. Transpl Int 15: 10-16, 2002.
Humes HD and Liu S. Cellular and molecular basis of renal repair in acute renal failure. J Lab Clin Med 124: 749-754, 1994.
Jaenke RS, Robbins ME, Bywaters T, Whitehouse E, Rezvani M, and Hopewell JW. Capillary endothelium. Target site of renal radiation injury. Lab Invest 68: 396-405, 1993.
Johnson RJ, Alpers CE, Yoshimura A, Lombardi D, Pritzl P, Floege J, and Schwartz SM. Renal injury from angiotensin II-mediated hypertension. Hypertension 19: 464-474, 1992.
Kang DH, Kim YG, Andoh TF, Gordon KL, Suga S, Mazzali M, Jefferson JA, Hughes J, Bennett W, Schreiner GF, and Johnson RJ. Post-cyclosporine-mediated hypertension and nephropathy: amelioration by vascular endothelial growth factor. Am J Physiol Renal Physiol 280: F727-F736, 2001.
Kassam G, Kwon M, Yoon S, Graham K, Young MK, Gluck S, and Waisman D. Purification and characterization of A61: an angiostatin-like plasminogen fragment produced by plamin autodigestion in the absence of sulfhydryl donors. J Biol Chem 276: 8924-8933, 2001.
Kjellstrand CM, Ebben J, and Davin T. Time of death, recovery of renal function, development of chronic renal failure and need for chronic hemodialysis in patients with acute tubular necrosis. Trans Am Soc Artif Intern Organs 27: 45-50, 1981.
Lewers D, Mathew TH, Maher JF, and Schreiner G. Long-term follow-up of renal function and histology after acute tubular necrosis. Ann Intern Med 73: 523-529, 1970.
Lijnen HR, Ugwu F, Bini A, and Collen D. Generation of an angiostatin-like fragment from plasminogen by stromelysin-1 (MMP-3). Biochemistry 37: 4699-4702, 1998.
Matsunaga T, Weihrauch DW, Moniz M, Tessmer J, Warltier DC, and Chilian WM. Angiostatin inhibits coronary angiogenesis during impared production of nitric oxide. Circulation 105: 2185-2191, 2002.
McKay D, Milford E, and Tolkoff-Rubin N. Clinical aspects of renal transplantation. In: The Kidney (6th ed.), edited by Brenner B. Philadelphia, PA: Saunders, 2000, p. 2542-2605.
Molema G, Van Veen-Hof I, Van Loenen-Weemaes A, Proost J, De Leu L, and Meijer D. Pharmokinetics and whole body distribution of elastase derived angiostatin (K1-3) in rats. Int J Cancer 91: 1-7, 2001. <a href="/cgi/external_ref?access_num=10.1002/1097-0215(20010101)91:1
Molitoris BA, Dahl R, and Geerdes A. Cytoskeleton disruption and apical redistribution of proximal tubule Na + -K + -ATPase during ischemia. Am J Physiol Renal Fluid Electrolyte Physiol 263: F488-F495, 1992.
Ohashi R, Kitamura H, and Yamanaka N. Peritubular capillary injury during the progression of experimental glomerulonephritis in rats. J Am Soc Nephrol 11: 47-56, 2000.
Ojo A, Wolfe R, Held P, Port F, and Schmouder R. Delayed graft function: risk factors and implications for renal allograft survival. Transplantation 63: 968-974, 1997.
O'Mahony CA, Albo D, Tuszynski GP, and Berger DH. Transforming growth factor- 1 inhibits generation of angiostatin by human pancreatic cancer cells. Surgery 124: 388-393, 1998.
O'Reilly MS, Holmgren L, Shing Y, Chen C, Rosenthal RA, Moses MA, Lane WS, Sage EH, and Folkman J. Angiostatin: a novel angiogenesis inhibitor that mediates supression of metastases by a Lewis lung carcinoma. Cell 79: 315-328, 1994.
O'Reilly MS, Wiederschain D, Stetler-Stevenson WG, Folkman J, and Moses MA. Regulation of angiostatin production by matrix metalloproteinase-2 in a model of concomitant resistance. J Biol Chem 274: 29568-29571, 1999.
Patterson BC and Sang QX. Angiostatin-converting enzyme activities of human matrilysin (MMP-7) and gelatinase B/Types IV Collegenase (MMP-9). J Biol Chem 272: 28823-28825, 1997.
Polito C, Papale M, and La Manna A. Long-term prognosis of acute renal failure in the full-term neonate. Clin Pediatr (Phila) 37: 381-386, 1998.
Rabb H, Mendiola CC, Dietz J, Saba SR, Issekutz TB, Abanilla F, Bonventre JV, and Ramirez G. Role of CD11a and CD11b in ischemic acute renal failure in rats. Am J Physiol Renal Fluid Electrolyte Physiol 267: F1052-F1058, 1994.
Rankin C, Itoh Y, Tian C, Ziemer D, Calvet J, and Gattone V. Matrix metallopreteinase-2 in a mujrine model of infantile-type polycystic kidney disease. J Am Soc Nephrol 10: 210-217, 1999.
Ray JM and Sttler-Stevenson WG. The role of matrix metalloproteases and their inhibitors in tumour invasion, metastasis and angiogenesis. Eur Respir J 7: 2062-2072, 1994.
Ray PE, Suga S, Liu XH, Huang X, and Johnson RJ. Chronic potassium depletion induces renal injury, salt sensitivity, and hypertension in young rats. Kidney Int 59: 1850-1858, 2001.
Safirstein R. Gene expression in nephrotoxic and ischemic acute renal failure. J Am Soc Nephrol 4: 1387-1395, 1994.
Shaw NJ, Brocklebank JT, Dickinson DF, Wilson N, and Walker DR. Long-term outcome for children with acute renal failure following cardiac surgery. Int J Cardiol 31: 161-166, 1991.
Sheridan AM and Bonventre JV. Cell biology and molecular mechanisms of injury in ischemic acute renal failure. Curr Opin Nephrol Hypertens 9: 427-434, 2000.
Sutton TA, Fisher CJ, and Molitoris BA. Microvascular endothelial injury and dysfunction during ischemic acute renal failure. Kidney Int 62: 1539-1549, 2002.
Sutton TA, Mang H, Campos SB, Sandoval R, Yoder M, and Molitoris BA. Injury of the renal microvascular endothelium alters barrier function following ischemia. Am J Physiol Renal Physiol 285: F191-F198, 2003.
Twining SS, Wilson PM, and Ngamkitidechakul C. Extrahepatic synthesis of plasminogen in the human cornea is upregulated by interleukins-1 and -1. Biochem J 339: 705-712, 1999.
Walker PD. Alterations in renal tubular extracellular matrix components after ischemia-reperfusion injury to the kidney. Lab Invest 70: 339-346, 1994.
Witzgall R, Brown D, Schwarz C, and Bonventre JV. Localization of proliferating cell nuclear antigen, vimentin, c-fos and clusterin in the post-ischemic kidney. J Clin Invest 93: 2175-2288, 1994.
Wu F, Li PL, and Zou AP. Microassay of 5'-nucleotidase and adenosine deaminase activity in microdissected nephron segments. Anal Biochem 266: 133-139, 1999.
Yamamoto T, Tada T, Brodsky SV, Tanaka H, Noiri E, Kajiya F, and Goligorsky MS. Intravital videomicroscopy of peritubular capillaries in renal ischemia. Am J Physiol Renal Physiol 282: F1150-F1155, 2002.
Ziswiler R, Daniel C, Franz E, and Marti HP. Renal matrix metallo-preoteinase activity is unaffected by experimental ischemia reperfusion injury and matrix metalloproteinase inhibition does not alter outcome of renal function. Exp Nephrol 9: 118-124, 2001.
作者单位：Departments of 1 Physiology and 2 Anesthesiology, Medical College of Wisconsin, Milwaukee, Wisconsin 53226; and the Department of 3 Physiology, Louisiana State University Health Science Center, New Orleans, Louisiana 70112