Central mechanisms leading to ischemia induced allograft rejection are apoptosis and inflammation, processes highly regulated by the urokinase-type plasminogen activator (uPA) and its specific receptor (uPAR). Recently, up-regulation of uPA and uPAR has been shown to correlate with allograft rejection in human biopsies. However, the causal connection of uPA/uPAR in mediating transplant rejection and underlying molecular mechanisms remain poorly understood. In this study, we evaluated the role of uPA/uPAR in a mice model for kidney ischemia reperfusion (IR) injury and for acute kidney allograft rejection. uPAR but not uPA deficiency protected from IR injury. In the allogenic kidney transplant model, uPAR but not uPA deficiency of the allograft caused superior recipient survival and strongly attenuated loss of renal function. uPAR-deficient allografts showed reduced generation of reactive oxygen species and apoptosis. Moreover, neutrophil and monocyte/macrophage infiltration was strongly attenuated and up-regulation of the adhesion molecule ICAM-1 was completely abrogated in uPAR-deficient allografts. Inadequate ICAM-1 up-regulation in uPAR−/− primary aortic endothelial cells after C5a and TNF-α stimulation was confirmed by in vitro experiments. Our results demonstrate that the local renal uPAR plays an important role in the apoptotic and inflammatory responses mediating IR-injury and transplant rejection.

Urokinase-type plasminogen activator (uPA)3 is a multifunctional molecule that serves either as a proteolytic enzyme or as a signal-inducing ligand. The urokinase receptor (uPAR; CD87) was originally identified as a proteinase receptor for uPA, directing pericellular proteolysis. However, accumulating data clearly demonstrate that uPAR can also activate a variety of intracellular signal pathways via its lateral interaction with different cell surface proteins such as integrins, growth factor receptors, and G-protein-coupled membrane proteins. These interactions enable the uPA/uPAR system not only to control pericellular fibrinolytic and proteolytic activities, but also to modulate cell adhesion, migration, proliferation, and differentiation (1, 2). Moreover, the important role of the uPA/uPAR system has recently been demonstrated in both innate and adaptive immune-mediated responses (3). The uPA/uPAR system modulates Ag processing and presentation (4), lymphocyte activation (5), generation of pro- and anti-inflammatory signals (6), activation of intracellular signaling pathways (7), cytotoxicity (8), cell adhesion (9), and migration (10, 11, 12), all of which are critical steps in cell-mediated immune responses. Furthermore, uPA potentiates neutrophil activation (7) and superoxid production (13). Recently, we were also able to demonstrate an important link between the uPA/uPAR and complement system in the regulation of immunological responses in kidney (14) and lung tissues (15).

Renal ischemia reperfusion (IR) injury after transplantation leads to acute renal failure, which profoundly affects both early and long-term allograft function (16). Accumulating evidence demonstrates that prolonged cold ischemia time is a strong risk factor for unfavourable outcome after allogenic kidney transplantation (17, 18) and suggests that the severity of IR injury to the allograft determines its immunogenicity and subsequent graft fate (19). The mechanisms underlying IR damage of kidney tissue seem to be multifactorial and interdependent. The oxygen supply to the tissue by reperfusion leads to the generation of reactive oxygen species (ROS) exceeding the protective anti-oxidative capacity of kidney cells. Oxidative stress is known to be a major apoptotic stimulus in allograft nephropathy (20). Furthermore, IR injury stimulates the components of innate immune response, such as complement activation and up-regulation of multiple proinflammatory genes including chemokines, cytokines, cytokine receptors, and adhesion molecules. This inflammatory response induced by innate mechanisms early after transplantation is markedly amplified by the subsequent adaptive response (16). Therefore, IR injury initiates and induces the alloimmune response leading to acute and chronic allograft rejection (21, 22).

Recently, up-regulation of the uPA/uPAR system has been demonstrated under hypoxia/reoxygenation conditions in vitro (23, 24). Moreover, uPA/uPAR activation has been shown to correlate with allograft rejection in a human biopsy study (25, 26) implying a probable involvement of this system in acute and chronic allograft rejection. However, the causal connection of uPA/uPAR in mediating of IR injury and transplant rejection as well as the possible molecular mechanisms for uPA/uPAR mediated initiation and perpetuation of inflammatory reaction after transplantation remain unexplored.

The aim of our study was to assess the role of the renal uPA/uPAR system in renal IR injury and in allogenic kidney transplant rejection. We demonstrate that uPAR but not uPA deficiency of the allograft protected from IR injury and acute allogenic transplant rejection. As underlying mechanisms we could elucidate impaired susceptibility of the uPAR-deficient allografts to IR-induced ROS-mediated apoptosis and reduced expression of adhesion molecules, leading to impaired migration of host monocytes/macrophages and neutrophils into uPAR−/− allografts.

The uPA- and uPAR-deficient mice (uPA−/− and uPAR−/−), generated as previously described (27), were a gift from Peter Carmeliet and Mieke Dewerchin (Center for Transgene Technology and Gene Therapy, University of Leuven, Belgium). uPA/uPAR deficiency was verified by PCR genotyping. The uPA−/− mice on a C57BL/6J background and the corresponding WT controls (WT1), as well as uPAR−/− mice on a mixed C57BL/6J (75%) × 129/Sv (25%) background and their WT littermate controls (WT2) served as kidney donors (H2b). BALB/c (H2d) mice served as recipients in kidney transplantation experiments and were supplied by Charles River Laboratories. The animals were bred under pathogen-free conditions in the animal facility of Phenos GmbH (Hannover, Germany) and cared for in accordance with our institution’s guidelines for experimental animals. All experiments were approved by the animal protection committee of the local authorities. Mice weighing 25–30g were used for all experiments. For IR injury 7–12 mice were used in each group (8 uPAR−/− mice, 7 WT2 mice, 10 uPA−/− mice, and 12 WT1 mice). For transplant experiments 6 mice for each group were used for the survival study and additional 6 mice for each group and each time point were sacrificed at 4 h, at day 1, and at day 6 after transplantation for histological and molecular analysis.

Renal IR-injury was induced in homozygous male uPA−/−, uPAR−/−, WT1, and WT2 mice by bilateral clamping of both renal pedicals. The animals were anesthetized with isoflurane. After median laparatomy, renal pedicals were bluntly dissected and a nontraumatic vascular clamp was applied for 35 min. At 24 h, postischemia kidney function was estimated by serum creatinine measurement using an automated method (Beckman Analyzer).

For transplant experiments, homozygous female uPA−/−, uPAR−/−, WT1, and WT2 mice were used as kidney donors and female BALB/c (H2d) mice served as recipients. Vascularized kidney transplantation was performed as described previously (28). In brief, the animals were anesthetized with isoflurane, and the left donor kidney attached to a cuff of the aorta and the renal vein with a small caval cuff and the ureter were removed en bloc. After left nephrectomy of the recipient, the vascular cuffs were anastomosed to the recipient abdominal aorta and vena cava, respectively, below the level of the native renal vessels. The ureter was directly anastomosed to the bladder (29). The times of cold and warm ischemia of allografts were 60 and 30 min, respectively. The right native kidney was removed through flank incision either on the day of transplantation or four days later. After transplantation, kidney function was estimated at designated time points (ranging from 24 h to several weeks) by serum creatinine level measurement. The general physical condition of the animals was monitored for any evidence of rejection.

Kidney grafts were harvested 24 h and six days after transplantation and one half of each allograft was immediately fixed in buffered formalin and embedded in paraffin. Sections of 3 μm were stained with hematoxylin-eosin and periodic acid-Schiff stain, and evaluated according to the updated Banff classification (30) by a nephropathologist, who was masked to the experimental groups. Immunohistochemistry was performed using the following primary Abs: rat anti-mouse monocytes/macrophages (F4/80; Serotec), polyclonal rabbit anti-mouse active caspase 3 (BD Pharmingen), monoclonal rat anti-mouse neutrophils (Gr-1, a gift from Prof. Hoffmann, Hannover Medical School, Hannover, Germany), monoclonal rat anti-mouse T lymphocytes (CD4 and CD8; BD Pharmingen), rabbit polyclonal anti-mouse CD25 (Santa Cruz Biotechnology), rabbit polyclonal anti-mouse uPAR (Santa Cruz Biotechnology). For indirect immunofluorescence, nonspecific binding sites were blocked with 10% normal donkey serum (Jackson ImmunoResearch Laboratories) for 30 min. Thereafter, sections were incubated with the primary Ab for 1 h. All incubations were performed in a humid chamber at room temperature (RT). For fluorescent visualization of bound primary Abs, sections were further incubated with Cy3 conjugated secondary Abs (Jackson Immuno Research Laboratory) for 1 h. Specimens were analyzed using a Zeiss Axioplan-2 imaging microscope with the computer program AxioVision 3.0 (Zeiss). Analysis of inflammation was done by semiquantitative scoring of the infiltrating cells in 10 randomly chosen, nonoverlapping fields of cortex and outer medulla (original magnification, ×200). Score: 0, no; 1, weak; 2, moderate; 3, high; and 4, very high numbers of infiltrating cells. For CD4 and CD25 expression, absolute cell numbers were counted in 20 nonoverlapping view fields each specimen. Active caspase-3 expression in the outer strip of the outer medulla was scored as follows: 0 < 10%, 1 = 10–30%, 30–50%, 4 > 50% of the tubuli affected. ICAM-1 expression in the cortex was scored as follows: 0 < 10%, 1 = 10–30%, 30–50%, 4 > 50% of the glomeruli affected. The analysis was done without knowledge of the animal assignment.

Priming of alloantigen-specific T cells from kidney graft recipients was investigated by performing MLR assay based on the measurement of BrdU incorporation during DNA synthesis. The responder spleen cells obtained from naive BALB/c mice or from WT2 or uPAR−/− allograft recipients at day 6 after transplantation were treated with ammonium chloride solution (Cell Systems) to lyse erythrocytes, washed three times, and resuspended at 3 × 106 cells/ml in complete RPMI 1640 medium (Life Technologies) supplemented with 10% FCS (Sigma-Aldrich), 2 mM l-glutamine, 100 U/ml penicillin, 100 mg/ml streptomycin, and 100-μl aliquots were delivered in triplicate to the wells of a 96-well, flat-bottom tissue culture plate. Stimulator cells were prepared from the spleens of syngeneic (i.e., BALB/c) and allograft donors (i.e.WT2, uPAR−/−). After lysis of erythrocytes the stimulator cells were treated with 50 μg/ml mitomycin C for 30 min at 37°C, washed and resuspended in culture medium at 3 × 106 cells/ml, and 100-μl aliquots per well were added to responder cells and cultured for 48 h. Afterward, BrdU was added and 18h later responder cell proliferation was quantified using colorimetric Cell Proliferation ELISA kit (Roche Diagnostics GmbH) in accordance to the manufacturer’s instructions. Syngeneic stimulator cells were used as background controls and were subtracted from alloresponses.

Priming of alloantigen-specific T cells from kidney allograft recipients was also tested by enumerating of IFN-γ-producing T cells using a mouse IFN-γ ELISpot kit (BD Biosciences) in accordance to the manufacturer’s instructions. Spleen cell suspensions were prepared from naive BALB/c mice or from allograft recipients at day 6 after transplantation and used as responder cells. Spleen cells from WT2 or syngeneic BALB/c mice were prepared and treated with mitomycin C for use as stimulator cells in the assay as described above. Responder and stimulator cells were cultured together for 24 h at 37°C in 5% CO2. The resulting spots were counted using the A.EL.VIS 4-Plate ELISPOT reader with the A.EL.VIS ELISPOT analysis software.

The redox-sensitive fluorophore dihydroethidium (DHE) was used to evaluate O2-production in the kidney in situ (31). Frozen tissue Cryosections of 6 μM were incubated with 0.1 mM DHE dissolved in HEPES-Tyrode buffer solution (132 mM NaCl, 4 mM KCl, 1 mM CaCl2, 0.5 mM MgCl2, 9.5 mM HEPES, and 5 mM glucose) for 12 min RT. After incubation, images were obtained using the Leica imaging system IM 500 (Ex, 520 nm; Em, 605 nm). Semiquantitative scoring was performed as follows: score: 0, no; 1, weak; 2, moderate; 3, high; and 4, very high intensity.

For TUNEL-assay (terminal deoxynucleotidyl transferase-mediated dUTP nick-end labeling) 2 μm sections of 4% paraformaldehyde (PFA)-fixed paraffin-embedded tissues were deparaffinized, treated with the terminal deoxynucleotidyl transferase enzyme and incubated in a humidified chamber at 37°C for 1 h. After washing, the tissue was treated with FITC-labeled anti-digoxygenin, incubated for 60 min, and washed. Negative controls were prepared under the same conditions, with the omission of the terminal deoxynucleotidyl transferase enzyme. TUNEL pos. cell numbers were counted in 20 nonoverlapping view fields each specimen without knowledge of the animal assignment.

Frozen kidneys were grinded in liquid nitrogen and total RNA was extracted using Trizol reagent (Invitrogen). For real-time quantitative PCR (qPCR), 1 μg of DNase-treated total RNA was reverse transcribed using Superscript II Reverse transcriptase (Invitrogen) and qPCR was performed on an SDS 7700 system (Applied Biosystems) using Rox dye (Invitrogen), FastStart taq Polymerase (Roche Diagnostics) and gene specific primers and FAM-Tamra-labeled probes (BioTez). PCR amplification was conducted at 10 min 96°C and 40 cycles at 10 s 95°C and 1 min at 60°C. β-actin served as the reference gene for normalization. Primer sequences are available on request. Quantification was conducted using qgene software (32).

Isolation of mouse aortic endothelial cells from 6- to 8-wk-old WT or uPAR−/− mice was performed as described previously (33). The cells were cultivated in medium consisting of endothelial cell growth medium 2 (Clonetics/Cambrex) and DMEM (1:1) supplemented with 20% FCS, 100 μg/ml endothelial cell growth supplement (Sigma-Aldrich), 100 U/ml penicillin, 100 mg/ml streptomycin, 2 mM l-glutamine, 0.5% nonessential amino acids, and 0.1 mg/ml heparin. MAEC at passage 3 were used for endothelial cell characterization and for all experiments. The endothelial nature of cells was confirmed by the typical cobblestone morphology of confluent monolayers, by Dil-Ac-LDL uptake and by surface expression of CD31 and CD106 analyzed by immunnocytochemistry (data not shown).

WT and uPAR−/− MAEC were seeded and cultured on glass coverslips. Serum-starved cells were fixed with 4% PFA in PBS for 20 min at RT. Nonspecific binding was blocked by 2 h incubation at RT with 3% BSA in PBS; the preparations were washed three times with PBS. Incubations with primary Ab (rat anti-mouse receptor for C5a anaphylatoxin (C5aR) clone 20/70 from Hycult Biotechnology or polyclonal rabbit anti-TNFR1 from Santa Cruz Biotechnology) were performed for 2 h at RT. Incubations with Alexa 488-conjugated secondary Abs (Molecular Probes) were performed for 1 h. After staining, cells were embedded in Poly-Mount mounting medium (Polysciences). The fluorescence cell images were captured using a Leica TCS-SP2 AOBS confocal microscope (Leica Microsystems). All the images were taken with oil-immersed × 63 objective, NA = 1.4.

Cell surface expression of ICAM-1 on MAEC was measured by cell ELISA as described previously (34). In brief, MAEC were grown in 96-well plate until 80% confluent, starved for 4 h and then stimulated for 16 h with 100 ng/ml recombinant murine C5a (mrC5a; Sigma-Aldrich) or with 5 ng/ml recombinant murine TNF-α (mrTNFα; R&D Systems). These concentrations were chosen in preliminary experiments ranging from 0.5 ng/ml to 200 ng/ml for each stimulus. For inhibition experiments MAEC were preincubated for 2 h with 100 ng/ml pertussis toxin (Sigma-Aldrich) or with 20 μg/ml hamster anti-mouse-TNFR1 mAb (R&D Systems). MAEC incubated in medium without stimuli served as a control. After fixation with 3% PFA and blocking with 3% BSA to prevent nonspecific binding the cells were incubated for 2 h with polyclonal rabbit-anti-mouse ICAM-1 Ab (Santa Cruz Biotechnology).The specific binding of Abs was then evaluated by incubation of cells for 1 h with secondary peroxidase conjugated goat anti-rabbit IgG (Santa Cruz Biotechnology) followed by addition of tetramethylbenzidine substrate solution (R&D Systems), stopping the reaction with 0.5 M H2SO4 and measuring the OD at 450 nm. The substrate was then washed away with deionized water, the plate allowed to dry and 0.5% trypan blue was added to stain for the number of cells/well. Excess of trypan blue was washed away and 1% SDS was added to solubilize the trypan blue stained cells. Each well was then read at 595 nm. The OD of ICAM-1 staining was divided by the OD of trypan blue staining to yield ICAM-1 expression for each well.

The adhesion of WT2 PBMC isolated by Ficoll-Paque separation to the endothelial surface of aortas obtained from WT2 or uPAR−/− mice was determined by the counting of adherent cells fluorescently labeled with the acetyloxymethyl ester of calcein (Calbiochem). A total of 1–2 mm pieces of aortas cleaned carefully of periadventitial fat and connective tissue and opened longitudinally were placed adventitia-side down on collagen I coated 96-well plates containing 10 μl of endothelial cell basal medium 2 (Clonetics/Cambrex) supplemented with 5% FCS to allow adherence of the aortic pieces to the substratum. When the pieces were well-attached (after 4 h), 200 μl of EBM-2 medium containing 50 ng/ml mrC5a, or 50 ng/ml mrTNFα was added. These concentrations were chosen in preliminary experiments ranging from 0.5 ng/ml to 200 ng/ml for each stimulus. For inhibitory experiments the pieces were preincubated for 2 h with 100 ng/ml pertussis toxin or with 20 μg/ml neutralizing hamster anti-mouse-TNFR1 mAb. Aorta pieces incubated in medium alone served as a control. After 16 h, the aorta pieces were washed twice and 100∗103 fluorescently labeled WT2 PBMC in 200 μl medium were added. The cells were allowed to adhere for 45 min at 37°C. Unbound cells were removed by washing three times. Photographs of aorta pieces were then made using the Leica imaging microscope with the digital image-processing program. The bound leukocytes were counted without knowledge of the group assignment in four different view-fields per aorta piece.

Data are shown as mean ± SEM. Normal distribution was analyzed by Klomogorov-Smirnov-test and statistical significance was calculated by Student’s t test for independent groups. SPSS 12.01 software was used.

The uPA/uPAR system is involved in a variety of signaling cascades which mediate IR injury. We first tested whether uPA or uPAR are mediators or effectors of renal IR injury. We performed bilateral renal pedical clamping in uPA−/− and uPAR−/− mice and their corresponding wild-type controls (WT1 and WT2, respectively). Both WT1 and WT2 controls and uPA−/− mice demonstrated severe loss of renal function within 24 h postischemia, as reflected in elevation of serum creatinine levels. In contrast, uPAR-deficient animals were protected from IR injury and demonstrated statistically significant attenuated loss of renal function (Fig. 1,A). Because apoptosis is a hallmark of IR injury, we performed a TUNEL assay in normal kidney tissue and in kidneys 24 h after ischemia (Fig. 1 B). No TUNEL-positive cells were detected in normal kidneys in any of the four groups of animals (data not shown). We detected increased numbers of TUNEL-positive nuclei in WT1, WT2 and uPA−/− kidneys. Because WT1 and WT2 mice showed almost identical results, only the data from WT2 mice are shown. However, in uPAR−/− kidneys the amount of TUNEL-positive cells was significantly reduced. These results suggest that uPAR contributes to hypoxia-induced apoptosis independently from uPA.

FIGURE 1.

Attenuation of IR-induced renal dysfunction and apoptosis in uPAR−/− mice. A, Renal function was investigated by measurement of serum creatinine level before (control) and at 24 h after IR. IR-injury caused severe renal dysfunction with elevation of serum creatinine level in WT1, WT2, and uPA-deficient mice. In contrast, uPAR deficiency led to markedly attenuated loss of renal function after IR injury (∗∗∗, p < 0.001 vs WT2, 7–12 mice were investigated for each group). B, Representative specimens after TUNEL assay 24-h postischemia are shown (original magnification, ×200). Because WT1 and WT2 mice demonstrated comparable results, representative data from WT2 kidney is shown. WT kidneys (a) and uPA-deficient kidneys (b) had increased numbers of apoptotic cells mainly in the tubular epithelium. In contrast, postischemic uPAR-deficient kidneys (C) showed almost no TUNEL positive cells. The results of quantification are presented as mean ± SEM in the lower panel (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group).

FIGURE 1.

Attenuation of IR-induced renal dysfunction and apoptosis in uPAR−/− mice. A, Renal function was investigated by measurement of serum creatinine level before (control) and at 24 h after IR. IR-injury caused severe renal dysfunction with elevation of serum creatinine level in WT1, WT2, and uPA-deficient mice. In contrast, uPAR deficiency led to markedly attenuated loss of renal function after IR injury (∗∗∗, p < 0.001 vs WT2, 7–12 mice were investigated for each group). B, Representative specimens after TUNEL assay 24-h postischemia are shown (original magnification, ×200). Because WT1 and WT2 mice demonstrated comparable results, representative data from WT2 kidney is shown. WT kidneys (a) and uPA-deficient kidneys (b) had increased numbers of apoptotic cells mainly in the tubular epithelium. In contrast, postischemic uPAR-deficient kidneys (C) showed almost no TUNEL positive cells. The results of quantification are presented as mean ± SEM in the lower panel (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group).

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Because activation of uPA/uPAR system has been shown to correlate with allograft rejection in a human biopsy study (25), we tested whether or not a similar phenomenon may be observed in kidney allografts in mice. We performed allogenic kidney transplantation using WT2 mice (H2b) as donors, and BALB/c mice (H2d) as recipients. The expression of uPA and uPAR was then investigated in normal WT2 kidneys and in WT2 kidney allografts at 4 and 24 h after transplantation. We detected strong up-regulation of uPAR (Fig. 2,A) mRNA 4 h after transplantation with further increase after 24 h. The up-regulation of uPA mRNA was less pronounced but also could be observed at 24 h after transplantation (Fig. 2,B). At protein level (Fig. 2 C), the weak expression of uPAR seen in normal WT2 kidney was strongly up-regulated within 24 h after transplantation and further increased at day 6. As expected, no positive signal was detected in uPAR−/− allografts (data not shown). These data demonstrate that transplantation results in local activation of the uPA/uPAR system in rejecting allografts from WT mice.

FIGURE 2.

uPA and uPAR expression is up-regulated in WT kidney allografts. uPAR (A) and uPA (B) mRNA expression in WT2 kidney allografts was investigated by qPCR before (control) and at 4 and 24 h after allogenic kidney transplantation. Both uPAR and uPA mRNA expression was time-dependently up-regulated after transplantation (∗, p < 0.05; ∗∗, p < 0.01 vs control, six mice were investigated for each group). C, The uPAR protein expression in WT2 allografts was investigated by immunocytochemistry. The upper row, uPAR expression in the cortex area; the lower row, uPAR expression in the outer strip of the outer medulla. Low baseline uPAR protein expression seen in normal WT2 kidney (control) was increased within 24 h and even more at 6 days after transplantation. (Original magnification ×200; six mice were investigated for each group).

FIGURE 2.

uPA and uPAR expression is up-regulated in WT kidney allografts. uPAR (A) and uPA (B) mRNA expression in WT2 kidney allografts was investigated by qPCR before (control) and at 4 and 24 h after allogenic kidney transplantation. Both uPAR and uPA mRNA expression was time-dependently up-regulated after transplantation (∗, p < 0.05; ∗∗, p < 0.01 vs control, six mice were investigated for each group). C, The uPAR protein expression in WT2 allografts was investigated by immunocytochemistry. The upper row, uPAR expression in the cortex area; the lower row, uPAR expression in the outer strip of the outer medulla. Low baseline uPAR protein expression seen in normal WT2 kidney (control) was increased within 24 h and even more at 6 days after transplantation. (Original magnification ×200; six mice were investigated for each group).

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Further, we analyzed an influence of uPA or uPAR deficiency of the allograft on recipient survival (Fig. 3 A). We performed allogenic kidney transplantation using uPA- and uPAR-deficient mice (H2b) and their corresponding WT controls as donors, and BALB/c mice (H2d) as recipients. All recipients of WT as well as uPA−/− allografts died shortly after allogenic kidney transplantation. In contrast, recipients of uPAR-deficient allografts showed superior survival for more than 20 wk after transplantation.

FIGURE 3.

Effect of allograft uPA and uPAR deficiency on survival, renal function and renal morphology after allogenic kidney transplantation. A, Survival analysis of mice receiving uPA- and uPAR-deficient allografts as compared with the animals receiving corresponding WT allografts is shown. WT1 and WT2 allografts revealed identical results and are presented as WT. Survival of recipients of uPA−/− allografts was comparable to WT allograft controls. The majority of all these transplant recipients died within 4 wk after transplantation. In contrast, recipients of uPAR−/− allografts showed prolonged allograft survival over 20 wk after transplantation. B, Renal function was estimated in recipient mice before (day 0) and at designated time points after transplantation by serum creatinine measurement. Recipients of uPA−/− and both WT1 and WT2 allografts had a similar steep rise of serum creatinine at 24 h and at 6 days after transplantation, correlating with acute rejection. In contrast, recipients of uPAR-deficient allografts had only a moderate initial rise in creatinine within 24 h after transplantation and remained stable at slightly increased serum creatinine levels thereafter. The results are presented as mean ± SEM (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group). C, Morphological changes one day after kidney transplantation are shown. All WT (a) and the uPA-deficient allografts (b) showed diffuse severe ATN. In contrast, the uPAR-deficient allograft recipients (c) had no signs of ATN. (PAS stain, original magnification ×200; six mice were investigated for each group).

FIGURE 3.

Effect of allograft uPA and uPAR deficiency on survival, renal function and renal morphology after allogenic kidney transplantation. A, Survival analysis of mice receiving uPA- and uPAR-deficient allografts as compared with the animals receiving corresponding WT allografts is shown. WT1 and WT2 allografts revealed identical results and are presented as WT. Survival of recipients of uPA−/− allografts was comparable to WT allograft controls. The majority of all these transplant recipients died within 4 wk after transplantation. In contrast, recipients of uPAR−/− allografts showed prolonged allograft survival over 20 wk after transplantation. B, Renal function was estimated in recipient mice before (day 0) and at designated time points after transplantation by serum creatinine measurement. Recipients of uPA−/− and both WT1 and WT2 allografts had a similar steep rise of serum creatinine at 24 h and at 6 days after transplantation, correlating with acute rejection. In contrast, recipients of uPAR-deficient allografts had only a moderate initial rise in creatinine within 24 h after transplantation and remained stable at slightly increased serum creatinine levels thereafter. The results are presented as mean ± SEM (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group). C, Morphological changes one day after kidney transplantation are shown. All WT (a) and the uPA-deficient allografts (b) showed diffuse severe ATN. In contrast, the uPAR-deficient allograft recipients (c) had no signs of ATN. (PAS stain, original magnification ×200; six mice were investigated for each group).

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Next, we studied the effect of uPA/uPAR deficiency on renal function after transplantation by measuring serum creatinine levels (Fig. 3,B). To monitor the effects of rejection on renal function we removed both recipient kidneys and transplanted the donor kidney in the same operation. An increase in serum creatinine was detected in all groups 24 h after transplantation; however this effect was strongly attenuated in mice which received uPAR−/− allografts. Six days after transplantation, recipients of uPA-deficient allografts and WT allografts developed deleterious loss of renal function as reflected in severe creatinine elevation. In contrast, the renal function of recipients of uPAR-deficient allografts remained stable without any significant increase in serum creatinine. Histological analysis revealed severe diffuse acute tubular necrosis (ATN) in WT and uPA−/− but not in uPAR−/− allografts at d1 after transplantation (Fig. 3 C). These results indicate that the uPAR but not uPA of allograft origin contributes significantly to the loss of renal function and allograft rejection in this model of allogenic kidney transplantation. Because uPA deficiency had no protective effects in either the IR model or in kidney transplantation, the following experiments were performed using only uPAR-deficient animals and their respective WT2 controls.

Because hypoxia-induced apoptosis is the pathophysiological correlate for ATN, we performed a TUNEL assay to investigate the cell death rate in uPAR−/− and WT2 kidney allografts (Fig. 4). WT2 allografts ubiquitously developed TUNEL positive nuclei as early as 4 h (a) and more pronouncedly 24 h (c) after transplantation. In contrast, uPAR−/− allografts demonstrated almost no signs of apoptosis at these time points (b and d). To elucidate whether the differences in cell death between WT2 and uPAR−/− allografts were due to necrosis or to apoptosis we performed immunohistochemistry for cleaved caspase-3. Especially in the outer stripe of the outer medulla, the area which is most sensitive to hypoxic damage, and in the medium of the vessels, cleaved caspase-3 level was increased in WT2 allografts at 24 h after transplantation (data not shown) and even more so at day 6 after kidney transplantation (e). In comparison, uPAR−/− allografts (f) showed markedly lower levels of cleaved caspase-3. These results were in line with our previous finding in the IR injury model and clearly demonstrate that local uPAR expression is also pivotal in ischemia triggered apoptosis in the allogenic kidney transplantation model.

FIGURE 4.

Allograft uPAR-deficiency strongly protected kidneys from transplantation-induced apoptosis. TUNEL assay was performed in WT2 and uPAR−/− allografts at 4 (upper row) and 24 h (middle row) after transplantation. WT2 allografts (a and c) had many TUNEL positive nuclei at 4 h (a) with further increase at 24 h (c) after transplantation. In contrast, uPAR-deficient allografts had almost no TUNEL positive cells at (b) and 24 h (d) after transplantation. (TUNEL assay, original magnification ×200). The quantification results are presented as mean ± SEM in the right-hand panel (∗∗, p < 0.01; ∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group). The immunohistochemistry for cleaved caspase-3 showed in WT2 allografts an intense up-regulation of cleaved caspase-3 level in the tubuli of the outer stripe of the outer medulla, the area most sensitive to hypoxic damage (e). uPAR-deficient allografts in contrast showed only few cleaved caspase-3 positive tubuli (f).

FIGURE 4.

Allograft uPAR-deficiency strongly protected kidneys from transplantation-induced apoptosis. TUNEL assay was performed in WT2 and uPAR−/− allografts at 4 (upper row) and 24 h (middle row) after transplantation. WT2 allografts (a and c) had many TUNEL positive nuclei at 4 h (a) with further increase at 24 h (c) after transplantation. In contrast, uPAR-deficient allografts had almost no TUNEL positive cells at (b) and 24 h (d) after transplantation. (TUNEL assay, original magnification ×200). The quantification results are presented as mean ± SEM in the right-hand panel (∗∗, p < 0.01; ∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group). The immunohistochemistry for cleaved caspase-3 showed in WT2 allografts an intense up-regulation of cleaved caspase-3 level in the tubuli of the outer stripe of the outer medulla, the area most sensitive to hypoxic damage (e). uPAR-deficient allografts in contrast showed only few cleaved caspase-3 positive tubuli (f).

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Hypoxia leads to tissue damage via generation of ROS contributing significantly to the mitochondrial apoptotic pathway (35). Scavengers for ROS, H2O2, and superoxide have been shown to inhibit apoptosis induced by ischemia-reperfusion (36). Therefore, we tested the hypothesis that strongly reduced apoptosis in uPAR−/− allografts might be due to decreased production of ROS. DHE staining (Fig. 5) performed in normal WT2 (a) and uPAR−/− (b) kidneys revealed similar basal levels of ROS. However, in contrast to WT allografts demonstrating severe up-regulation of ROS at 4 (c) and 24 h (e) after transplantation, ROS generation was markedly reduced in uPAR-deficient allografts (d and f). This result suggests that uPAR contributes to the initial apoptosis triggering step - the generation of ROS.

FIGURE 5.

uPAR-deficiency of the allograft strongly reduces ROS generation after transplantation. Generation of ROS was investigated by DHE stain. The upper row shows low basal level of DHE staining in WT2 (a) and uPAR−/− (b) kidneys before transplantation. An increase in generation of ROS in the tubulo-interstitial compartment was detected in WT2 allografts as early as 4 h (c) after transplantation; this effect was still detectable 24 h after transplantation (e). In contrast, uPAR-deficient allografts showed impaired generation of ROS at 4 (d) and 24 h (f) after transplantation. The results of the semiquantitative analysis are presented as mean ± SEM in the right-hand panel (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group).

FIGURE 5.

uPAR-deficiency of the allograft strongly reduces ROS generation after transplantation. Generation of ROS was investigated by DHE stain. The upper row shows low basal level of DHE staining in WT2 (a) and uPAR−/− (b) kidneys before transplantation. An increase in generation of ROS in the tubulo-interstitial compartment was detected in WT2 allografts as early as 4 h (c) after transplantation; this effect was still detectable 24 h after transplantation (e). In contrast, uPAR-deficient allografts showed impaired generation of ROS at 4 (d) and 24 h (f) after transplantation. The results of the semiquantitative analysis are presented as mean ± SEM in the right-hand panel (∗∗∗, p < 0.001 vs WT2, six mice were investigated for each group).

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The inflammatory cell infiltration of the allograft is a hallmark of allograft rejection (37, 38). Therefore, we performed immunohistochemistry for different cell subsets to elucidate the composition of the cell infiltrates. No differences were observed between the groups in CD8 and CD4 infiltrates. Moreover, the CD25 expression on CD4 T cells was comparable in WT2 and uPAR−/− allografts (Fig. 6,A). In line with this observation the MLR assay did not reveal any differences between splenocytes obtained from WT2 and uPAR−/− allograft recipients at day 6 after transplantation (Fig. 6,B). The splenocytes from WT2 and uPAR−/− allograft recipients showed a similar increase of proliferative response to WT2 stimulator cells compared with splenocytes from naive BALB/c mice. The frequency of alloantigen-specific IFN-γ-producing T cells tested by ELISPOT assay was also comparable in WT2 and uPAR−/− allograft recipients (Fig. 6 C). This result shows that uPAR deficiency did not decrease the level of donor-reactive T cell priming in this model.

FIGURE 6.

uPAR-deficiency of the allograft does not change host T cell infiltration and alloantigen-specific T cell priming after transplantation. A, At day 6 after transplantation, WT2 allografts (a) and uPAR-deficient allografts (b) showed similar CD8 positive T cell infiltration. Furthermore, CD4 positive T cell infiltrates of similar density were seen in both WT2 and uPAR−/− allografts (c and d). No differences were observed concerning CD25 positive T cells between WT2 and uPAR−/− allografts (e and f). The results of the quantification analysis are presented as mean ± SEM in the right-hand panel six mice were investigated for each group). B, In vitro MLR assay was performed with splenocytes isolated from naive BALB/c mice and from recipients of WT2 and uPAR−/− kidney allografts at day 6 after transplantation. No differences between WT2 and uPAR−/− allograft recipients were observed. Experiments were performed in triplicate, n = 5 mice for each group, data are presented as mean ± SEM. C, IFN-γ-producing cells at day 6 after transplantation in spleen cells of naive BALB/c mice and recipients of WT2 and uPAR−/− kidney allografts were determined by ELISPOT analysis. Numbers of IFN-γ-producing cells were similar in uPAR−/− and WT2 allograft recipients. Experiments were performed in triplicate, n = 5 mice for each group, data are presented as mean ± SEM.

FIGURE 6.

uPAR-deficiency of the allograft does not change host T cell infiltration and alloantigen-specific T cell priming after transplantation. A, At day 6 after transplantation, WT2 allografts (a) and uPAR-deficient allografts (b) showed similar CD8 positive T cell infiltration. Furthermore, CD4 positive T cell infiltrates of similar density were seen in both WT2 and uPAR−/− allografts (c and d). No differences were observed concerning CD25 positive T cells between WT2 and uPAR−/− allografts (e and f). The results of the quantification analysis are presented as mean ± SEM in the right-hand panel six mice were investigated for each group). B, In vitro MLR assay was performed with splenocytes isolated from naive BALB/c mice and from recipients of WT2 and uPAR−/− kidney allografts at day 6 after transplantation. No differences between WT2 and uPAR−/− allograft recipients were observed. Experiments were performed in triplicate, n = 5 mice for each group, data are presented as mean ± SEM. C, IFN-γ-producing cells at day 6 after transplantation in spleen cells of naive BALB/c mice and recipients of WT2 and uPAR−/− kidney allografts were determined by ELISPOT analysis. Numbers of IFN-γ-producing cells were similar in uPAR−/− and WT2 allograft recipients. Experiments were performed in triplicate, n = 5 mice for each group, data are presented as mean ± SEM.

Close modal

Furthermore, we analyzed infiltration of neutrophils (Fig. 7,A, upper panel) and monocytes/macrophages (Fig. 7 A, middle panel). We found that uPAR deficiency of the allograft decreased the number of infiltrating neutrophils and monocytes/macrophages as compared with WT2 control allografts. These results underline the importance of local renal uPAR for monocyte/macrophage and neutrophil infiltration.

FIGURE 7.

uPAR-deficiency of the allograft protects from host leukocyte infiltration after transplantation. A, At day six after transplantation, WT2 allografts (a) showed more tubulo-interstitial infiltrates of Gr-1 positive neutrophils as compared with uPAR-deficient allografts (b). Dense infiltration of F4/80 positive monocytes/macrophages seen in the tubulo-interstitium of WT2 allografts (c) was strongly reduced in uPAR-deficient allografts (d). ICAM-1 expression was studied by immunohistochemistry. Twenty-four hours after transplantation ICAM-1 was heavily up-regulated in the glomeruli of WT2 allografts (e) whereas uPAR−/− allografts (f) showed almost no ICAM-1 up-regulation. The results of the semiquantification analysis are presented as mean ± SEM in the right-hand panel (∗∗, p < 0.01; ∗∗∗, p < 0.001 vs WT2, original magnification ×200, six mice were investigated for each group). B, The expression of MCP-1 (upper panel) and MIP-2 (low panel) mRNA was analyzed by TaqMan RT-PCR in normal kidneys (control) and in WT2 and uPAR−/− allografts obtained at 24 h and day 6 after transplantation. Data are expressed as the mean ± SEM (∗∗, p < 0.01 vs WT2, six mice were investigated for each group).

FIGURE 7.

uPAR-deficiency of the allograft protects from host leukocyte infiltration after transplantation. A, At day six after transplantation, WT2 allografts (a) showed more tubulo-interstitial infiltrates of Gr-1 positive neutrophils as compared with uPAR-deficient allografts (b). Dense infiltration of F4/80 positive monocytes/macrophages seen in the tubulo-interstitium of WT2 allografts (c) was strongly reduced in uPAR-deficient allografts (d). ICAM-1 expression was studied by immunohistochemistry. Twenty-four hours after transplantation ICAM-1 was heavily up-regulated in the glomeruli of WT2 allografts (e) whereas uPAR−/− allografts (f) showed almost no ICAM-1 up-regulation. The results of the semiquantification analysis are presented as mean ± SEM in the right-hand panel (∗∗, p < 0.01; ∗∗∗, p < 0.001 vs WT2, original magnification ×200, six mice were investigated for each group). B, The expression of MCP-1 (upper panel) and MIP-2 (low panel) mRNA was analyzed by TaqMan RT-PCR in normal kidneys (control) and in WT2 and uPAR−/− allografts obtained at 24 h and day 6 after transplantation. Data are expressed as the mean ± SEM (∗∗, p < 0.01 vs WT2, six mice were investigated for each group).

Close modal

Adhesive interaction between up-regulated adhesion molecules on activated endothelium with blood leukocytes is a crucial step for leukocyte infiltration into the site of inflammation. Therefore, we analyzed the expression of ICAM-1 in WT2 and uPAR−/− allografts after kidney transplantation. Baseline ICAM-1 expression of uPAR−/− and WT kidneys was comparable (data not shown). However, a strong ICAM-1 up-regulation seen in WT2 allografts after transplantation was significantly impaired in uPAR−/− allografts (Fig. 7 A, lower panel).

Members of the chemokine family play a central role in inflammatory cell infiltration into extravascular sites by attracting and stimulating specific subsets of leukocytes (39). MCP-1 is an important mediator for monocyte recruitment (40) whereas MIP-2 is necessary for an adequate neutrophil infiltration (39). Therefore, we compared the transplantation-induced changes of MCP-1 and MIP-2 mRNA expression in WT2 and uPAR−/− allografts. As expected, we observed a strong up-regulation of these proinflammatory mediators in WT2 allografts. Surprisingly, uPAR−/−allografts demonstrated a practically identical pattern of the transient overexpression of MIP-2 mRNA, with a maximum at 24 h after transplantation. The kinetics of MCP-1 mRNA up-regulation was also similar in WT and uPAR−/− allografts, and moreover, the grade of MCP-1 mRNA up-regulation was even significantly higher in uPAR−/− kidney (Fig. 7 B). These results suggest that attenuated leukocyte infiltration into uPAR-deficient allografts may be due to impaired up-regulation of ICAM-1 but not to altered chemokine expression.

Recently, the interaction of complement activation product C5a with its receptor (C5aR) has been shown to induce a strong increase in gene expression for cell adhesion molecules in endothelial cells similar to those induced by TNF-α (41). Because both TNF-α and activated complement are important mediators of IR injury and transplant rejection (42, 43), we hypothesized that TNF-α and/or C5a-dependent up-regulation of endothelial adhesion molecules may be impaired in uPAR−/− allografts and may be an explanation for the reduced monocyte/macrophage and granulocyte infiltration. To test this hypothesis, we performed additional in vitro experiments with the primary culture of MAEC from WT2 and uPAR−/− mice. After demonstration of similar cell surface expression of both TNFR1 and C5aR on WT2 and uPAR−/− MAEC by immunocytochemistry (Fig. 8,A) we investigated the expression of ICAM-1 after stimulation of cells with TNF-α and C5a for 16 h by cell ELISA. The treatment with 5 ng/ml mrTNFα resulted in a strong up-regulation of ICAM-1 expression in both WT2 and uPAR−/− MAEC, however, this effect was significantly decreased in uPAR−/− MAEC compared with WT2. The pretreatment with TNFR1 blocking Ab strongly decreased the TNF-α-induced up-regulation of ICAM-1 suggesting the involvement of TNFR1 (Fig. 8,B). The treatment of MAEC with 100 ng/ml mrC5a resulted in a moderate but significant up-regulation of ICAM-1 in WT2 MAEC, however, this effect was completely abrogated in uPAR−/− MAEC. The pretreatment with the C5aR blocking pertussis toxin prevented the C5a-induced up-regulation of ICAM-1 in WT2 MAEC verifying the role of C5aR (Fig. 8 C).

FIGURE 8.

uPAR-deficiency strongly reduces C5a/TNF-α-induced up-regulation of ICAM-1 in endothelial cells and completely abrogates C5a/TNF-α-induced increase of WT leukocyte adhesion in vitro. A, Nonstimulated serum-starved MAEC obtained from WT and uPAR−/− mice were immunostained with monoclonal anti-C5aR (upper panel) or polyclonal anti-TNFR1 (lower panel) Abs and Alexa 488-conjugated secondary Abs. Basal expression of C5aR and TNFR1 was comparable in uPAR−/− and WT MAEC. (The frame size of images is 240 × 240 μm.) B, mrTNFα (left histogram) stimulation resulted in a strong up-regulation of ICAM-1 expression in WT2 and to a lesser extent also in uPAR−/− MAEC. Pretreatment with TNFR1 blocking Ab strongly decreased the TNF-α-induced up-regulation of ICAM-1. Nonstimulated cells served as controls. mrC5a stimulation (right histogram) resulted in a moderate but significant up-regulation of ICAM-1 in WT2 MAEC, however, this effect was completely abrogated in uPAR−/− MAEC. Pertussis toxin prevented the C5a-induced up-regulation of ICAM-1 in WT MAEC. (Data are expressed as the mean ± SEM from three independent experiments, performed in four parallels for each condition. Significant differences were determined by Student’s t test for C5a/TNF-α-stimulated vs nonstimulated cells (∗∗, p < 0.01; ∗∗∗, p < 0.001) and for WT2 vs uPAR−/− cells (++, p < 0.01; +++, p < 0.001). C, Adhesion of WT2 PBMC to the endothelial surface of aortas was studied in vitro. Aortas from WT2 or uPAR−/− mice were preincubated or not with 100 ng/ml pertussis toxin or with 20 μg/ml anti-TNFR1 Abs for 2 h and then stimulated for 16 h with 50 ng/ml C5a or TNF-α. Unstimulated aortas served as a control. C5a- and TNF-α-dependent increase of WT2 PBMC adhesion to uPAR−/− aorta was completely abrogated as compared with WT2 aortas. The data are presented as mean ± SEM for five individual experiments performed in triplicate (∗∗, p < 0.01 between C5a/TNF-α-stimulated and nonstimulated aortas and ++, p < 0.01; +++, p < 0.001 between WT2 and uPAR−/− aortas under the same stimulation conditions).

FIGURE 8.

uPAR-deficiency strongly reduces C5a/TNF-α-induced up-regulation of ICAM-1 in endothelial cells and completely abrogates C5a/TNF-α-induced increase of WT leukocyte adhesion in vitro. A, Nonstimulated serum-starved MAEC obtained from WT and uPAR−/− mice were immunostained with monoclonal anti-C5aR (upper panel) or polyclonal anti-TNFR1 (lower panel) Abs and Alexa 488-conjugated secondary Abs. Basal expression of C5aR and TNFR1 was comparable in uPAR−/− and WT MAEC. (The frame size of images is 240 × 240 μm.) B, mrTNFα (left histogram) stimulation resulted in a strong up-regulation of ICAM-1 expression in WT2 and to a lesser extent also in uPAR−/− MAEC. Pretreatment with TNFR1 blocking Ab strongly decreased the TNF-α-induced up-regulation of ICAM-1. Nonstimulated cells served as controls. mrC5a stimulation (right histogram) resulted in a moderate but significant up-regulation of ICAM-1 in WT2 MAEC, however, this effect was completely abrogated in uPAR−/− MAEC. Pertussis toxin prevented the C5a-induced up-regulation of ICAM-1 in WT MAEC. (Data are expressed as the mean ± SEM from three independent experiments, performed in four parallels for each condition. Significant differences were determined by Student’s t test for C5a/TNF-α-stimulated vs nonstimulated cells (∗∗, p < 0.01; ∗∗∗, p < 0.001) and for WT2 vs uPAR−/− cells (++, p < 0.01; +++, p < 0.001). C, Adhesion of WT2 PBMC to the endothelial surface of aortas was studied in vitro. Aortas from WT2 or uPAR−/− mice were preincubated or not with 100 ng/ml pertussis toxin or with 20 μg/ml anti-TNFR1 Abs for 2 h and then stimulated for 16 h with 50 ng/ml C5a or TNF-α. Unstimulated aortas served as a control. C5a- and TNF-α-dependent increase of WT2 PBMC adhesion to uPAR−/− aorta was completely abrogated as compared with WT2 aortas. The data are presented as mean ± SEM for five individual experiments performed in triplicate (∗∗, p < 0.01 between C5a/TNF-α-stimulated and nonstimulated aortas and ++, p < 0.01; +++, p < 0.001 between WT2 and uPAR−/− aortas under the same stimulation conditions).

Close modal

These results were confirmed by a PBMC adhesion assay ex vivo. In these experiments, adhesion of normal WT2 PBMC to the endothelial surface of aortas obtained from WT2 or uPAR−/− mice and stimulated in vitro with mrC5a or mrTNFα was investigated. As demonstrated in Fig. 8 D, 16-h stimulation of WT2 aorta with TNF-α or C5a induced significant increase in adhesion of WT2 PBMC. These effects could be strongly reduced or completely abolished when aortas were pretreated with TNFR1 blocking Ab and pertussis toxin, respectively. In contrast, the TNF-α and C5a-dependent up-regulation of endothelial adhesion was completely abolished in uPAR−/− aortas. These results clearly demonstrate that uPAR expression on endothelial cells is necessary for adequate TNF-α and C5a signaling leading to up-regulation of adhesion molecules and mediating leukocyte adhesion.

Our data provide the first evidence that local renal uPAR but not uPA expression plays a pivotal role in the pathogenesis of IR injury and allogenic transplant rejection. We ruled out that uPAR deficiency protected kidney tissue from generation of ROS and consecutively from severe apoptosis in IR injury and acute kidney allograft rejection. In the transplantation model, the resistance of the allograft against hypoxia-induced apoptosis due to uPAR deficiency was linked to better renal function and increased allograft survival. The inadequate up-regulation of ICAM-1 in the blood vessels of uPAR−/− allografts resulted in attenuated monocyte/macrophage and neutrophil infiltration.

Allogenic transplant rejection is triggered by several stimuli such as allograft hypoxia and subsequent apoptosis and inflammatory response. Tubular cell apoptosis is considered to be an important pathway leading to tubular atrophy in progressive renal disease. Recently, it has been shown that uPAR may modify the rate of apoptotic renal cell death (44). In vitro, human glioma cells exposed to uPAR antisense have been reported to undergo more apoptotic cell death (45). In contrast to these observations, the major finding of this study was that uPAR deficiency strongly protected renal cells from apoptosis via reduced ROS formation in both models, IR injury and kidney transplantation. One possible explanation for this discrepancy might be that uPAR exposes pro- or anti-apoptotic action and modulates the cell survival/apoptosis ratio depending on the cell type and/or the apoptosis-triggering pathways. This hypothesis is supported, for example, by demonstration of the anti-apoptotic and uPA-independent action of uPAR in endothelial cells in which anti-uPAR Abs as well as soluble recombinant uPAR blocked the apoptotic effect of cleaved high molecular mass kininogen by preventing of the binding of high molecular mass kininogen to these cells (46).

Recently, it has been shown that uPA stimulates ROS production in VSMC in vitro (47). Because hypoxia triggers ROS generation and mediates apoptosis, we investigated the possible role of uPA/uPAR interaction in ROS generation and subsequent ROS-dependent apoptosis using both uPA- and uPAR-deficient mice. Our results clearly demonstrate that in both models uPAR deficiency protected renal tissue from ROS generation independently from uPA. The molecular mechanisms underlying the attenuated ROS formation in uPAR−/− allografts remain to be investigated. uPAR can interact laterally with a wide variety of membrane proteins including integrins, endocytic receptors, caveolin, the gp130 cytokine receptor, the epidermal growth factor receptor, chemoattractant receptors (1) and platelet-derived growth factor receptor (2). This interaction might activate both pro- and anti-apoptotic downstream signaling pathways. For example, recently ROS-triggered apoptosis in polymorphonuclear leukocytes (PMNs) cells has been shown to be β2 integrin-dependent (48). Therefore, uPAR may stimulate ROS production via its lateral interaction with other molecules.

The early inflammatory response during reperfusion of allografts is initiated by the infiltration of PMNs into the graft (38) followed by infiltration of monocytes/macrophages (MO). Macrophages constitute 38 to 60% of infiltrating cells during acute allograft rejection and increased influx of MO has been strongly correlated to complement activation and acute rejection in a human protocol biopsy study (49). Furthermore, it has been shown that MO infiltration 3 mo after transplantation correlated inversely with graft survival (50). It is well known that uPAR deficiency on the surface of leukocytes strongly reduces their migratory capacity both in vivo and in vitro (15, 51, 52). However, it should be noted that in our transplant experiments, the host leukocytes expressed uPAR normally. Despite the normal expression of uPAR on host leukocytes, we found strongly reduced PMN/MO infiltration in kidney allografts from uPAR−/− as compared with those from WT2 mice. This finding stresses the importance of the local uPAR expression on resident renal cells in the allograft for adequate cell adhesion and subsequent PMN/MO accumulation after kidney transplantation, independently from the uPAR expression on infiltrating cells. Recently it has been shown that inhibition of PMN infiltration into cardiac allografts may have a significant downstream impact on the efficacy of recipient T cell responses to the allograft (38). In contrast, in our model of acute renal allograft rejection we did not find any differences in CD4 and CD8 positive cell infiltration between WT2 and uPAR−/− allografts after transplantation. Moreover, the expression of T cell activation Ag CD25 used as a marker for T cell activation (53) was also similar in WT2 and uPAR−/− allografts. Furthermore, by investigating of priming of alloreactive T cells by MLR and IFN-γ ELISPOT assays we did not found differences of T cell function of WT2 and uPAR−/− allografts recipients. These results suggest that superior survival of uPAR−/− deficient allografts is not mediated by T cell-dependent alloimmune response.

The migration of inflammatory cells into an extravascular site requires a series of coordinated signals including the generation of a chemotactic gradient by the cells of the extravascular compartment and up-regulation of adhesion molecules on activated endothelium. The strong up-regulation of MCP-1 and MIP-2 has been demonstrated in animal models during renal ischemia as well as in renal biopsies from patients with acute and chronic renal rejection (54). In line with this report, we observed a strong up-regulation of these proinflammatory mediators which was similar in WT2 and uPAR−/− allografts. Therefore, it could be concluded that the level of these chemoattractants was not responsible for the diminished PMN/MO infiltration in uPAR−/− allografts.

Adhesion molecules are rapidly up-regulated early after transplantation (55) and after induction of IR injury (56). Therefore, we analyzed expression of ICAM-1 in WT2 and uPAR−/− allografts given the facts that, being the major counterreceptor for leukocyte β2-integrins, this adhesion molecule is rapidly up-regulated within the first 2–3 h after transplantation (57) and the blockade of ICAM-1 strongly attenuated PMN infiltration in some experimental models of renal ischemia (58, 59, 60). Indeed, we could demonstrate that the transplantation-induced up-regulation of ICAM-1 observed in WT2 allografts was strongly attenuated in uPAR−/− allografts. However, the reduced expression of ICAM-1 in uPAR−/− allografts was not sufficient to prevent T cell infiltration. This result coincides with observation of Zhang and coworkers (61) demonstrating that primed alloreative T cells do not require allograft expression of ICAM-1 to infiltrate heart allografts. The interaction of integrin leukocyte function Ag-1 with its alternative ligand ICAM-2, or interaction of other T cell homing receptors with their ligands on endothelium, such as CD44/E-selectin interaction, may be involved in lymphocyte adhesion to the vascular endothelium of the graft and its subsequent infiltration bypassing the requirement for ICAM-1 (62).

By performing in vitro experiments with primary aortic endothelial cells we could elucidate impaired TNF-α and C5a signaling in uPAR−/− cells leading to strongly decreased TNF-α-induced and completely abrogated C5a-induced up-regulation of ICAM-1. In line with a prior report that only the TNFR1 receptor is involved in TNF-α-induced ICAM-1 up-regulation (63) we could demonstrate that the up-regulation of ICAM-1 in MAEC after TNF-α stimulation was mediated predominantly via TNFR1 because the neutralizing anti-mouse TNFR1 Ab almost completely blocked this effect. These results coincide with strongly reduced adhesion of WT2 PBMC to isolated aortas from uPAR−/− mice compared with WT2 aortas after stimulation with TNF-α and C5a. This result is in line with a prior report that platelet uPAR is critical for the response to TNF-α (64) and coincides also with our previous observation that uPAR is necessary for C5a/C5aR-mediated responses in mouse alveolar macrophages (15) and in human mesangial cells (14). Collectively, these data suggest that uPAR plays a major role in mediating IR injury and subsequently influences early inflammation and allograft survival in renal transplantation rather than directly affecting T cell mediated alloimmune responses.

In summary, this study shows that the local expression of uPAR on resident renal cells of the allograft contributes to the development of acute allograft rejection via at least two different pathways, ischemia-induced apoptosis and the infiltration of host leukocytes. These results suggest that uPAR, whose role was presumed to be involved in the migratory behavior of infiltrating cells, has a broader critical function as an early regulator of ischemia-triggered initial generation of ROS, which, in turn, induces the apoptosis in intrinsic renal cells. uPAR is also necessary for adequate TNF-α and C5a signaling, leading to up-regulation of ICAM-1 on endothelial cells of the allograft which is a crucial step for adequate leukocyte recruitment to the inflamed tissue. These observations underscore a new role of uPAR in acute allograft rejection and highlights uPAR as a target for prevention of organ dysfunction and damage in IR injury.

We thank Yvonne Nikolai, Herle Chlebusch, and Kerstin Bankes for excellent technical assistance.

The authors have no financial conflict of interest.

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

3

Abbreviations used in this paper: uPA, urokinase-type plasminogen activator; uPAR, urokinase-type plasminogen activator receptor; IR, ischemia reperfusion; ROS, reactive oxygen species; RT, room temperature; DHE, dihydroethidium; PFA, paraformaldehyde; MAEC, mouse aortic endothelial cell; PBMC, peripheral blood mononuclear cell; MO, monocytes/macrophages; PMN, polymorphonuclear leukocyte; ATN, acute tubular necrosis; C5aR, receptor for C5a anaphylatoxin; MO, macrophage/monocyte; WT, wild type.

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