reperfusion injury

Free Radical Biology and Medicine 74 (2014) 263–273

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Original Contribution

Complement-dependent NADPH oxidase enzyme activation in renal ischemia/reperfusion injury S. Simone a,1, F. Rascio b,n,1, G. Castellano a, C. Divella a, A. Chieti a, P. Ditonno c, M. Battaglia c, A. Crovace d, F. Staffieri d, B. Oortwijn e, G. Stallone b, L. Gesualdo a, G. Pertosa a,2, G. Grandaliano b,2 a

Nephrology, Dialysis, and Transplantation Unit, Department of Emergency and Organ Transplantation, University of Bari “Aldo Moro,” 70121 Bari, Italy Nephrology, Dialysis, and Transplantation Unit, Department of Medical and Surgical Sciences, University of Foggia, 71100 Foggia, Italy c Urology, Andrology, and Renal Transplantation Unit, and Department of Emergency and Organ Transplantation, University of Bari “Aldo Moro,” 70121 Bari, Italy d Veterinary Surgery Unit, Department of Emergency and Organ Transplantation, University of Bari “Aldo Moro,” 70121 Bari, Italy e Pharming BV, Leiden, The Netherlands b

art ic l e i nf o

a b s t r a c t

Article history: Received 11 March 2014 Received in revised form 2 July 2014 Accepted 3 July 2014 Available online 10 July 2014

NADPH oxidase plays a central role in mediating oxidative stress during heart, liver, and lung ischemia/ reperfusion injury, but limited information is available about NADPH oxidase in renal ischemia/ reperfusion injury. Our aim was to investigate the activation of NADPH oxidase in a swine model of renal ischemia/reperfusion damage. We induced renal ischemia/reperfusion in 10 pigs, treating 5 of them with human recombinant C1 inhibitor, and we collected kidney biopsies before ischemia and 15, 30, and 60 min after reperfusion. Ischemia/reperfusion induced a significant increase in NADPH oxidase 4 (NOX-4) expression at the tubular level, an upregulation of NOX-2 expression in infiltrating monocytes and myeloid dendritic cells, and 8-oxo-7,8-dihydro-20 -deoxyguanosine synthesis along with a marked upregulation of NADPH-dependent superoxide generation. This burden of oxidative stress was associated with an increase in tubular and interstitial expression of the myofibroblast marker α-smooth muscle actin (α-SMA). Interestingly, NOX-4 and NOX-2 expression and the overall NADPH oxidase activity as well as α-SMA expression and 8-oxo-7,8-dihydro-20 -deoxyguanosine synthesis were strongly reduced in C1-inhibitor-treated animals. In vitro, when we incubated tubular cells with the anaphylotoxin C3a, we observed an enhanced NADPH oxidase activity and α-SMA protein expression, which were both abolished by NOX-4 silencing. In conclusion, our findings suggest that NADPH oxidase is activated during ischemia/reperfusion in a complement-dependent manner and may play a potential role in the pathogenesis of progressive renal damage in this setting. & 2014 Elsevier Inc. All rights reserved.

Keywords: C1 inhibitor Complement system NOX-2 NOX-4 Oxidative stress Renal ischemia–reperfusion injury Renal transplantation Free radicals

Kidney transplantation represents the gold standard therapy for end-stage renal disease. Given the shortage of organs from deceased donors available for transplantation, a great effort has been made to expand the donor pool. From this perspective, uncontrolled deceased (after cardiac death) donors are becoming an increasing source of organs [1]. These grafts are prone to severe

Abbreviations: α-SMA, α-smooth muscle actin; C1-INH, C1 inhibitor; DGF, delayed graft function; DPI, diphenyleneiodonium; NADPH, nicotinamide adenine dinucleotide phosphate; NOX-2, NADPH oxidase 2; NOX-4, NADPH oxidase 4; 8-oxodG, 8-oxo-7,8-dihydro-20 -deoxyguanosine; ROS, reactive oxygen species; ZO1, zonula occludens 1 n Corresponding author. Fax: þ 39 0881 587123. E-mail address: [email protected] (F. Rascio). 1 These authors equally contributed to this study. 2 These authors contributed equally to this study as senior investigators. http://dx.doi.org/10.1016/j.freeradbiomed.2014.07.003 0891-5849/& 2014 Elsevier Inc. All rights reserved.

ischemia/reperfusion-induced renal damage and are characterized by a high rate of delayed graft function (DGF)3, leading to worse long-term function [2]. Indeed, also in this setting, the major risk factor for DGF is prolonged cold/warm ischemia time, suggesting that renal ischemia/reperfusion injury may play a significant role in its pathogenesis [2]. Hypoxia, featuring the ischemic phase, induces ATP depletion and hypoxanthine accumulation. During the reperfusion phase, the restoration of oxygen flow causes the generation of xanthine and reactive oxygen species (ROS) [3] leading to tubular necrosis/apoptosis and tissue inflammation [4]. The activation of the complement system has been shown to exacerbate this inflammatory process, playing a central role in the pathogenesis of ischemia/reperfusion-induced renal damage; the small complement fragments C3a and C5a contribute to local inflammation by acting on renal tubular epithelial cells and infiltrating cells [5,6]. As a result of the inflammatory

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and oxidative injury caused by ischemia/reperfusion, tubular epithelial cells may undergo profound functional alterations, losing tight junctions [7] and expressing myofibroblast markers [8,9]. Although the role of this event, also known as epithelial-tomesenchymal transition (EMT), in the pathogenesis of interstitial fibrosis is debated, in the transplant setting α-SMA expression at the tubular level is an early marker of progressive graft damage [10]. Hypoxia-induced ATP depletion does not completely explain the burden of oxidative stress after ischemia/reperfusion injury. Nicotinamide adenine dinucleotide phosphate (NADPH) oxidase represents the major source of superoxide production in several cell types [11] and its expression contributes to oxidative stress by increasing superoxide production [12,13]. The NADPH oxidase (NOX) complex consists of several subunits: NOX-2 (gp91phox) and p22phox represent the core region of the enzyme; p47phox, the “organizer subunit”; p67phox, the “activator subunit”; p40phox, the “adaptor subunit”; and the GTPase RAC is the cytosolic factor of the complex. Members of the NOX family participate in cellular functions related to inflammation, signal transduction, proliferation, and apoptosis. NOX-1, NOX-2, and NOX-4 are the three NOX isoforms expressed in the kidney [14–17]. Recent evidence suggests that NOX-4, originally identified as an NADPH oxidase homolog highly expressed in the kidney, plays a key role in modulating Toll-like receptor 4-mediated ROS production in posthypoxic renal tubular epithelial cells [18]. Although a role for NADPH oxidase has been demonstrated in the pathogenesis of ischemia/reperfusion-induced tissue injury in the brain, liver, and heart [19–21], existing literature is less exhaustive about the role of this enzyme in renal ischemia/reperfusion and the mechanisms responsible for ischemia/reperfusion-induced NADPH activation. Thus, the aim of this study was to investigate the expression and activation of NOX-4 in a swine model of renal ischemia/ reperfusion injury. Because we already demonstrated the pathogenic role of the complement cascade in the same swine model of ischemia/reperfusion [22], we examined the possible link between oxidative stress/NOX activity, complement priming, and tubulointerstitial α-SMA expression. Materials and methods Renal ischemia/reperfusion injury model After approval by the ethics committee of the Italian Ministry of Health, 10 4-month-old female Large White pigs weighing 40 kg underwent experimental open surgical procedures under general anesthesia. The animals were treated as previously described [22]. Briefly, they were fasted for 24 h before surgery. Vital signs were continuously monitored during anesthesia. The left renal artery and vein were isolated and a vessel loop was placed around the renal artery with a right-angle clamp. A basal renal biopsy was performed (T0) and ischemia was induced for 30 min by pulling on the vessel loop. Five minutes before the end of ischemia, recombinant human C1 inhibitor (Pharming), diluted in saline solution, was injected into the ear vein of 5 animals (500 U/kg); in another group of 5 animals an equal volume of vehicle was infused at the same time point (control group). Multiple biopsies were then performed at 15, 30, and 60 min after reperfusion. The animals were sacrificed 24 h after this surgical procedure. Immunohistochemical analysis Two-micrometer-thick sections of paraffin-embedded renal tissue were rehydrated through a series of xylene and graded alcohol washes. After antigen retrieval, endogenous peroxidase

was inhibited by incubation with 3% H2O2 for 7 min. The sections were blocked with serum-free protein block (Dako, Glostrup, Denmark) for 10 min at room temperature, incubated with the primary antibodies (NOX-4, 1:200, ab60940 rabbit polyclonal; 8-oxo-7,8-dihydro-20 -deoxyguanosine (8-oxodG), 1:100, ab26842 mouse monoclonal, both from Abcam, Cambridge, UK) and detected by the Dako EnVision G/2 system, according to the manufacturer’s instructions (Dako). The sections were counterstained with Mayers hematoxylin (blue) and mounted with glycerol (DakoCytomation, Carpinteria, CA, USA). Negative controls were obtained by incubating serial sections with the blocking solution and substituting the primary antibodies with an irrelevant IgG of the same species (mouse or rabbit). Specific NOX-4 immunostaining was quantified and expressed as pixel/area fraction using ImageScope software. Tissue immunofluorescence and confocal laser scanning microscopy NOX-2 and p22phox protein expression was evaluated by indirect immunofluorescence and confocal microscopy on paraffin-embedded swine kidney sections. After antigen unmasking in citrate buffer, sections were washed in phosphate-buffered saline (PBS) and blocked with 5% goat serum in PBS for 1 h at room temperature. Sections were incubated overnight at 4 1C with a primary antibody against NOX-2 (mouse IgG1 anti-53/gp91phox, BD Biosciences, Franklin Lakes, NJ, USA; 1:10) and p22phox (rabbit polyclonal, Santa Cruz Biotechnology, Santa Cruz, CA, USA; 1:100), followed by incubation for 1 h with the appropriate secondary antibody (Alexa Fluor 488 goat anti-mouse 1:200, Alexa Fluor 555 goat anti-rabbit 1:200, Molecular Probes, Eugene, OR, USA). The sections were counterstained with To-Pro-3 (Molecular Probes), mounted in Gel/Mount (Biomeda Corp., Foster City, CA, USA), and sealed with nail varnish. Negative controls were obtained by substituting the primary antibody with a control irrelevant IgG. Specific fluorescence was acquired by a Leica TCS SP2 (Leica, Wetzlar, Germany) confocal laser-scanning microscope using a 63  objective lens. Two independent observers (C.D. and F.R.), blinded to the origin of the slides, counted the number of tubular NOX-2 þ cells, p22phox þ cells, and NOX-2 þ /p22phox þ cells in at least 15 consecutive high-power (630  ) fields for each sample. The values were then averaged. The final reported count was the mean of the two observers’ measures. In no case was the interobserver variability greater than 20%. Paraffin-embedded swine kidney sections were double stained for NOX-2 and CD163 or SWC3a. After antigen unmasking, sections were permeabilized in PBS with 0.05% Tween 20 for 5 min, washed in PBS, and then blocked with 5% goat serum in PBS for 1 h at room temperature. Sections were incubated overnight at 4 1C with a primary antibody against NOX-2 (1:10 in 2% goat serum) followed by incubation for 1 h with Alexa Fluor 488 goat anti-mouse antibody (Molecular Probes, 1:200). Sections were washed in PBS and then blocked with 5% goat serum in PBS, incubated overnight at 4 1C with primary antibodies against SWC3a (dendritic cells, 74-22-15 A, BD Biosciences; 1:200 in 2% goat serum) or CD163 (monocytes/macrophages, US Biological, Swampscott, MA, USA; 1:50 in 2% goat serum) followed by incubation for 1 h with the appropriate secondary antibody. The sections were counterstained with To-Pro-3 (Molecular Probes), mounted in Gel/Mount (Biomeda), and sealed with nail varnish. Paraffin-embedded sections (3–4 mm) from kidney biopsies removed at various time points were used to evaluate α-SMA protein expression by single immunofluorescence and double immunofluorescence (α-SMA/ZO-1). After antigen unmasking in citrate buffer, sections were washed in PBS and incubated with 5% goat serum in PBS for 1 h at room temperature. For single staining, sections were incubated overnight at 4 1C with a primary antibody against α-SMA

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(1:100, sc-32251 mouse monoclonal anti-α-SMA, Santa Cruz Biotechnology) followed by incubation for 1 h with the appropriate secondary antibody (1:200, Alexa Fluor 488 goat anti-mouse, Molecular Probes). The sections were counterstained with To-Pro-3 (1:3000, Molecular Probes), mounted in Gel/Mount (Biomeda), and sealed with nail varnish. Negative controls were obtained by omitting the primary antibodies. For double staining, α-SMA/ZO-1, sections were incubated overnight at 4 1C with a mix of primary antibodies against α-SMA and ZO-1 (1:50, sc-10804 rabbit polyclonal anti-ZO-1, Santa Cruz Biotechnology) followed by incubation for 1 h with a mix of appropriate secondary antibodies (1:200 Alexa Fluor 488 goat anti-mouse, Molecular Probes, for α-SMA, and 1:200 Alexa Fluor 555 goat anti-rabbit, Molecular Probes, for ZO-1). After being washed in PBS, the sections were counterstained with To-Pro-3 (1:3000, Molecular Probes), mounted in Gel/Mount (Biomeda), and sealed with nail varnish. Negative controls were obtained by omitting the primary antibodies. Specific fluorescence was acquired by a Leica TCS SP2 (Leica) confocal laser-scanning microscope. The number of α-SMA þ cells was measured in at least 10 high-power (630  ) fields/section by two independent observers (C.D. and F.R.) blinded to the origin of the slides. The final counts were the means of the two measures and interobserver variability was lower than 20%. Cell cultures and small interfering RNA transfection Immortalized human proximal tubular epithelial cells (HK2) were purchased from American Type Culture Collection (Rockville, MD, USA) and grown in Dulbecco’s modified Eagle’s–F12 medium (Sigma–Aldrich) supplemented with 1% antibiotics and 10% fetal bovine serum (Sigma–Aldrich). For the experiments HK-2 cells were grown to confluence, starved for 24 h in serum-free culture medium, and stimulated with C3a (5  10  7 M; Calbiochem, Darmstadt, Germany) for the indicated time periods. NOX-4 small interfering RNA (siRNA) and scrambled siRNA were purchased from Qiagen (Valencia, CA, USA). For in vitro delivery, siRNA (25 nM) was incubated with 5 ml TransIT-TKO transfection reagent (Mirus, Madison, WI, USA) for 20 min at room temperature and added to cells in culture at 80% confluence in six-well plates. All procedures were performed according to the manufacturer’s instructions. Western blotting The Western blotting was performed as previously described [25]. Antibody directed toward α-SMA was from Santa Cruz Biotechnology (1:500, sc-32251 mouse monoclonal anti-human α-SMA) and the antibody against NOX-4 was from Abcam (1:1000, ab41886 rabbit polyclonal anti-human). Anti-β-actin antibody was from Sigma–Aldrich (1:10,000 mouse monoclonal anti-human β-actin, clone AC-15, Sigma–Aldrich). Intensities of protein bands were evaluated and quantified using ImageJ (NIH, Bethesda, MD, USA). NADPH oxidase assay Kidney cortex NADPH oxidase activity was measured by the lucigeninenhanced chemiluminescence method in portions of renal tissue (n ¼5) obtained by needle biopsies before ischemia (T0) and after reperfusion (30 and 60 min; T300 and T600 ) from control and C1INH-treated animals. We also analyzed one portion of T600 control biopsy after treatment (30 min) with DPI (10 mM), an inhibitor of NADPH oxidase. Tissue fragments were homogenized on ice in 1 ml of lysis buffer (20 mM KH2PO4, pH 7.0, 1 mM EGTA, 1 mM phenylmethanesulfonyl fluoride, 8 mg/ml leupeptin). Homogenates were subjected to a centrifugation at 800 g for 10 min at 4 1C.

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To start the assay, 100 μl of supernatant was added to 900 μl of 50 mM phosphate buffer, pH 7.0, containing 1 mM EGTA, 150 mM sucrose, 5 μM lucigenin, and 100 μM NADPH. Photon emission was measured as relative light units (RLU) every 30 s for 5 min. No measurable activity was detected in the absence of NADPH. Superoxide generation was expressed as RLU/min mg protein. Cultured tubular cells NADPH oxidase activity was measured as above described. Briefly, to evaluate NADPH oxidase activity in cultured tubular cells under basal conditions, in response to C3a and after NOX-4 silencing (24 h), both C3a-stimulated and unstimulated HK-2 cells were grown in serum-free medium containing 5 mM glucose for 24 h. HK-2 cells were stimulated with C3a (5  10  7 M) and treated with siRNA for NOX-4 and then washed in PBS, harvested in the same solution, and centrifuged at 800 g for 10 min at 4 1C. The cell pellets were resuspended in lysis buffer and homogenized on ice. Aliquots of the homogenates were used for the assay as previously described. Statistical analysis Data are expressed as the mean 7standard deviation. Differences between groups were analyzed by unpaired t-test analysis. A p value o 0.05 was considered statistically significant. Statistical analysis was performed using the StatView software package (version 5.0; SAS, Inc., Cary, NC, USA).

Results NOX-4 protein expression increases after reperfusion We first investigated whether ischemia/reperfusion might modulate NOX-4 protein expression in our swine model. Immunohistochemistry, performed on renal biopsies from control animals, showed that NOX-4 protein expression, hardly detectable at baseline (Fig. 1A), increased in a time-dependent manner with a peak 60 min after reperfusion and was predominantly localized at the tubular level (Figs. 1B–D). Fig. E shows negative control performed on T600 kidney biopsies. Quantification of NOX-4 þ tubules demonstrated a statistically significant increase in the NOX-4-expressing tubules at T600 compared to basal condition (T0) (Fig. 1F). These data were confirmed by Western blotting on tissue homogenates from T0, T150 , T300 , and T600 kidney biopsies (Figs. 1G and H). NOX-2 and p22phox protein expression increases after reperfusion We then evaluated, by confocal microscopy, whether ischemia/ reperfusion might modulate the expression of NOX-2, the main NADPH oxidase isoform expressed by inflammatory cells. NOX-2 (green) protein expression increased in a time-dependent manner with a peak at 60 min after reperfusion and was mainly localized within interstitial infiltrating cells (Fig. 2A–D). Considering that NOX-2 is unstable in the absence of the p22phox subunit and the association of the NOX-2/p22phox complex is required for activation [14], we investigated the modulation by ischemia/reperfusion of p22phox and its colocalization with NOX-2. The p22phox (red) protein expression, almost absent before ischemia, increased within the interstitial infiltrating cells after 15 and 30 min of reperfusion, reaching the peak after 60 min (Fig. 2E–H). Interestingly, NOX-2 (green) colocalized with p22phox (red) at all time points after reperfusion, but not before ischemia (Fig. 2I–L). Quantification of NOX-2 þ cells and of p22phox þ cells at the various times of reperfusion demonstrated a significant increase in NOX2 þ cells and p22phox þ cells after 60 min (Fig. 2M and N).

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Fig. 1. NOX-4 protein expression increases after ischemia/reperfusion. Paraffin-embedded kidney sections were examined for expression of NOX-4 by immunohistochemistry in control biopsies (n¼ 5). (A–D) Tubular NOX-4 expression increased along the time course. Immunohistochemistry for NOX-4 was carried out as described under Materials and methods. Specific staining was detected by DAB (brown) and nuclei were counterstained by hematoxylin (blue). Original magnification  200. Arrow indicates NOX4 þ tubules. (E) No measurable NOX-4 signal was detected in the absence of primary antibody (negative control). (F) Quantification of NOX-4 þ tubules was done at various times of reperfusion. Results are expressed as the mean7SD of NOX-4 þ tubules (T300 versus T0 p¼0.04; T600 versus T0 p¼0.03 (n¼ 5 for each group). (G) NOX-4 protein expression was also evaluated by direct immunoblotting of total lysates (40 μg) from T0, T150 , T300 , and T600 swine biopsies. (H) Histogram shows Western blotting analysis (n¼3); β-actin protein expression was used for normalization. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

Interestingly, quantification of double-positive NOX-2 þ /p22phox þ cells at the various times of reperfusion demonstrated a significant increase at the same time point (Fig. 2O). To identify the infiltrating cell populations expressing NOX-2 and probably involved in ROS generation in our model of ischemia/reperfusion-induced renal injury, we investigated whether NOX-2 expression colocalized with SWC3a, a marker of myeloid dendritic cells, or CD163, a marker of monocytes/macrophages. Our previous data suggested a significant influx in the interstitium of both cell types in our experimental model [22]. We observed the coexpression of NOX-2 with both SWC3a (Fig. 3A–C) and CD163 (Fig. 3D–F), suggesting the activation of the NOX-2 isoform in these two cell populations.

Ischemia/reperfusion-induced NOX-4 and NOX-2 expression and enzyme activity were reduced by C1-INH To investigate the pivotal role of this enzyme in ROS generation, we measured NADPH oxidase activity in renal biopsies before and after reperfusion. Ischemia/reperfusion caused a robust increase in NADPH-driven superoxide generation after 60 min of reperfusion compared to basal levels (p ¼ 0.01). The preincubation of tissue homogenates from T600 biopsies with DPI, an inhibitor of flavin-containing oxidases, restores the basal level of superoxide generation (p¼ 0.02 versus T600 ; Fig. 4A).

We then investigated the role of the complement cascade in modulating NADPH oxidase activation after renal ischemia/reperfusion. To this purpose, we evaluated NADPH oxidase activity in five animals undergoing renal ischemia/reperfusion and treated with recombinant C1-INH before stopping ischemia. NADPHdependent superoxide generation, expressed as RLU/min mg, was strongly reduced at T600 in cortical homogenates of C1-INHtreated animals (n ¼5) compared to the control group (n ¼5) at the same time point (p¼ 0.04; Figs. 4A and 4B). In agreement with this observation, immunohistochemistry clearly demonstrated a reduction in NOX-4 protein expression (Figs. 4C and 4D) and NOX2 protein expression (Figs. 4E and 4F) in C1-INH-treated animals compared to the control group. Quantification by ImageScope software showed a statistically significant reduction in NOX-4 þ tubules and NOX-2 þ cells in C1-INH-treated animals compared to the control group (Figs. 4G and 4H, respectively). Renal ischemia/reperfusion injury is associated with an increase in 8-oxodG To confirm the generation of oxidative stress after ischemia/ reperfusion, we evaluated the accumulation of a sensitive marker of oxidative damage, 8-oxodG, in our in vivo model of ischemia/ reperfusion injury. Consistent with NOX-4 overexpression in tubular areas and the increased NOX-2 þ cells at the interstitial level,

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Fig. 2. NOX-2 and p22phox protein expression increases postreperfusion. Paraffin-embedded swine kidney sections were examined for NOX-2 and p22phox protein expression by immunofluorescence and confocal microscopy. (A–D) NOX-2 protein expression was significantly upregulated at T600 . (E–H) The p22phox protein expression also increased in a time-dependent manner after reperfusion, with a peak at T600 within interstitium-infiltrating cells. (I–L) The merge of NOX-2 (green) with p22phox (red). Quantification (M) of NOX-2 þ cells, (N) of p22phox þ cells, and (O) of NOX-2 þ p22phox þ cells at various times of reperfusion. Results are expressed as the mean 7SD of NOX-2 þ cells/highpower field (hpf) *po 0.04, **p o 0.01.

ischemia/reperfusion injury also caused a marked increase in 8-oxodG synthesis in control pigs at T300 and T600 (Fig. 5A–C). Interestingly, 8-oxodG-specific staining was significantly reduced in C1-INH-treated animals (Fig. 5D–F). These data, confirmed by ImageScope quantification (Fig. 5G) further support our hypothesis of a pivotal role for the complement system in ischemia/reperfusion-induced oxidative stress.

α-SMA expression induced by ischemia/reperfusion is abrogated by complement inhibition

α-SMA þ myofibroblasts are the main actors in the pathogenesis of renal interstitial fibrosis. Because α-SMA production is

modulated by NOX-4-derived ROS in some cell lines [23,24], we investigated the expression of this myofibroblast marker in our swine model of ischemia/reperfusion injury. In the control group, only a few α-SMA þ cells could be detected within the interstitial space at baseline (Fig. 6A), whereas a significant increase in these cells was observed within the tubulointerstitial areas of renal biopsies at T300 and T600 (Figs. 6B–6D). In the same model we previously demonstrated that α-SMA expression is still high 24 h after reperfusion [26]. In contrast, the expression of this marker was almost totally abrogated by C1-INH treatment (Fig. 6E–H). Quantification of α-SMA þ cells by confocal microscopy confirmed a significant

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Fig. 3. NOX-2 is expressed by infiltrating SWC3a þ myeloid dendritic cells and CD163 þ monocytes/macrophages. Representative images of paraffin-embedded swine kidney sections showing that all NOX-2 þ cells (green) expressed SWC3a (red) (A–C) and all NOX-2 þ cells expressed CD163 (red) (D–F). Nuclei were stained with To-Pro (blue).

inhibition in C1-INH-treated animals compared to controls (Fig. 6I). Interestingly, at the most significant time point of our model, at T600 , we also observed a colocalization of α-SMA with a spatially deregulated ZO-1, an epithelial-specific marker, in several tubular sections (Fig. 6J–L). NOX-4 silencing abrogates C3a-induced NADPH oxidase activity and α-SMA protein expression in cultured tubular cells To further confirm the hypothesis suggested by our in vivo data, that ischemia/reperfusion-induced NADPH expression and activity are complement-mediated and that NOX-4 activation is important for α-SMA expression, we investigated the ability of C3a to modulate NOX-4 activity and α-SMA expression in cultured proximal tubular cells and evaluated whether NOX-4 knockout may play a role in C3a-induced α-SMA mRNA abundance. The anaphylotoxin C3a caused a statistically significant increase in NADPH-dependent superoxide generation compared to basal conditions (p¼ 0.01; Fig. 7A and B). To demonstrate that ROS production was NADPH-dependent, we also evaluated NADPH oxidase activity after preincubation with H2O2/C3a together with DPI treatment. Thirty minutes of DPI incubation was sufficient to reduce H2O2-induced or C3a-induced ROS generation below basal levels (p ¼0.009 and p ¼0.004, respectively; Fig. 7A and B). C3 also induced a rapid increase in NOX-4 protein expression with a peak at 15 min (Fig. 7C and D). To assess the effects of NOX-4-driven superoxide production on α-SMA expression, we knocked out NOX-4 by RNA interference in cultured proximal tubular cells [25]. After silencing of NOX-4 with a specific siRNA for 24 h, we evaluated NADPH oxidase activity in response to C3a, in both NOX-4 siRNA- and scramble siRNA-transfected cells. As expected, NOX-4 silencing caused a

reduction of enzyme activity in response to C3a stimulation (Fig. 8A and B). We, then, stimulated cultured tubular cells with C3a (24 h) and investigated α-SMA protein expression. Again, NOX-4 RNA silencing resulted in a limited reduction of α-SMA expression compared with basal conditions but caused a significant reduction in C3a-stimulated cells, suggesting that NOX-4 mediates C3a-induced α-SMA expression (p¼ 0.04; Fig. 8C and D).

Discussion DGF is an early posttransplant condition associated with an increased rate of acute rejection episodes and a worse long-term graft survival [27]. Renal ischemia/reperfusion injury has a central role in the pathogenesis of this form of acute renal failure [4]. Oxidative stress is a key pathogenic mechanism in tissue injury induced by ischemia/reperfusion. Several investigators in various models of tissue ischemia/reperfusion observed a burst of reactive oxygen metabolites after reestablishing blood flow to a previously ischemic zone [28–31]. The extreme reactivity of these oxidants results in irreversible damage to vital cell components, including membrane phospholipids, membrane ion transport proteins and other enzymatic proteins [32]. Experiments using ROS scavengers in several models of ischemia/reperfusion injury demonstrated significant reductions in the extent of tissue injury and emphasized the role of ROS in this scenario [13,33,34]. There are multiple potential sources of ROS after ischemia/reperfusion, including nitric oxide synthase, xanthine oxidase, and the mitochondrial respiratory chain. An increasing amount of evidence suggests that NADPH oxidase plays a role in mediating oxidative stress during heart, liver, and lung ischemia/reperfusion [35–38]. To date limited

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Fig. 4. C1-INH infusion modulates NADPH activity and NOX-4 and NOX-2 expression. (A and B) NADPH oxidase activity in homogenates from cortical biopsies collected at T0, T300 , and T600 after reperfusion was measured. Superoxide anion generation was determined by photoemission every 30 s for 5 min as described under Materials and methods. NADPH-dependent superoxide production was expressed as RLU/min mg protein and values are the means 7 SD of the activities from the kidney cortex of five animals/group. A significant increase in NADPH oxidase activity was observed at T600 in control animals (p ¼ 0.01 versus T0) and a strong reduction was observed after DPI incubation. Histogram shows that NADPH oxidase activity was inhibited by C1-INH infusion. Data represent the means 7 SD of the activities from homogenates of five animals for each group. Compared with (C and E) untreated pigs, C1-INH-treated animals showed a strong reduction of (D) NOX-4 and (F) NOX-2 protein expression in the kidney. (G) Quantification of NOX-4 expression by ImageScope software is shown as % pixels/area fraction (T600 C1-INH vs T600 CTRL p¼ 0.03). (H) Quantification of NOX-2 is shown as % positive cells/total area (T600 C1-INH vs T600 CTRL p ¼0.02).

information is available on NADPH oxidase activation in renal ischemia/reperfusion-induced injury. The first step in our investigation was, then, to examine NADPH-dependent superoxide generation along with NOX-4 protein expression in a swine model of ischemia/reperfusion injury. NADPH oxidase complex, expressed in phagocytes and vascular tissue, represents an important source of ROS. NOX-4 is the main NADPH oxidase isoform expressed in the kidney [14]. Despite the requirement of all regulatory subunits for NOX-1, NOX-2, and NOX-3 function, NOX-4 works independent of the presence of any of these components [14]. Previous studies indicated that this isoform plays a key role in Toll-like receptor 4-activated apoptosis in an in vitro model of ischemia/reperfusion [18]. Yang et al. [39] demonstrated that wild-type mice receiving a bone marrow transplant from p47phox / animals presented a significant reduction of lung dysfunction and injury after ischemia/reperfusion. In addition, the induction of several proinflammatory cytokines and chemokines, including TNF-α, IL-17, IL-6, RANTES, KC, MIP-2, and MCP-1, was significantly reduced after ischemia/reperfusion in p47phox / /wild type chimeras. On the basis of these results it is plausible that NADPH oxidase-generated ROS specifically from bone marrow-derived cells contribute importantly to lung ischemia/reperfusion injury. Because

renal ischemia/reperfusion is also characterized by an early recruitment of inflammatory cells, we investigated the expression and activation of NOX-2 in our model of oxidative damage. For NOX-2, the main phagocytic isoform of NADPH oxidase, to be activated it needs to interact with p22phox in the presence of other subunits, and we demonstrated the colocalization of NOX-2 with p22phox in infiltrating cells after reperfusion, supporting the hypothesis of NOX-2 activation in this model. Thus, the increased expression and activation of this NADPH oxidase might significantly participate in the total superoxide generation observed after ischemia/reperfusion. Our model was not designed to investigate the effects of NOX-4 or NOX-2 on the development of tissue damage. It is conceivable, on the basis of our previous data, that the protective effects of complement inhibition that we recently reported [22] might be, at least in part, mediated by the inhibition of complement-induced NOX-2 and -4 expression and activity. However, in the present study we demonstrated a close association in vivo between NOX-4 activation and α-SMA expression. α-SMA is a specific marker for myofibroblasts, the main cellular type involved in tissue fibrosis, and its expression by tubular cells has been suggested as an early marker of fibrogenesis [24]. Badid et al. [10] demonstrated that tubulointerstitial expression of α-SMA represents a reliable

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Fig. 5. Ischemia/reperfusion causes an increase in 8-oxodG protein expression in control pigs but not in C1-INH-treated pigs. Kidney sections from CTRL and C1-INH-treated pigs were stained with 8-oxodG antibody. (A) 8-OxodG levels, very low in preischemia biopsies, were higher in CTRL pigs after (B) 30 and (C) 60 min of reperfusion compared to C1-INH-treated pigs at the same time points (D–F). Original magnification  200. (G) Quantification of 8-oxodG area fraction (%).

biomarker for the progression of graft damage in kidney transplantation. There is an increasing body of evidence suggesting that ischemia/reperfusion injury, in addition to causing an acute tubular injury, may prime a progressive renal damage leading to end-stage renal disease [40]. Indeed, in the clinical setting we are well aware that DGF is a negative prognostic factor for long-term graft survival [4]. The presence of ZO-1/α-SMA coexpression may suggest a role for EMT in the origin of interstitial myofibroblasts in this setting. However, EMT does not fully explain the increase in interstitial myofibroblasts. Indeed, we recently reported in the same model the concomitant activation of endothelial-tomesenchymal transition [26] and we cannot definitely exclude a further pathogenic mechanism. Several reports assert that complement activation leads to the generation of C3a and C5a, two anaphylotoxins that can both increase the recruitment of inflammatory cells and induce oxidative stress through ROS generation [41]. Thus, we used C3a in our in vitro experiments. The observation of the effect of C3a in cultured tubular cells not only supports our in vivo findings, but also suggests a direct link between the priming of the complement cascade, NOX-4 activation, and tubular α-SMA expression. However, it is conceivable that the infiltrating cells we observed in our

model, in addition to directly contributing to superoxide generation, might also influence tubular NOX-4 expression through the release of cytokines and growth factors [17]. Our swine model closely resembles the warm ischemia observed in deceased (after cardiac death) donors, a source of kidney grafts continuously increasing in the past few years. However, we cannot exclude that this model may be also relevant for the classical scenario with transplantation under controlled conditions, which is mainly, but not exclusively, characterized by a cold ischemia. Our observation may then suggest a potential therapeutic approach to reduce the oxidative burden and the subsequent acute graft damage in the setting of posttransplant ischemia/reperfusion injury. Indeed, based on our data the inhibition of the complement cascade and/or of NOX-4 activation might influence the early and late outcomes of kidney transplantation. In conclusion, our data suggest that NADPH oxidase is activated during renal ischemia/reperfusion in a complement-dependent manner, playing a key role in ROS generation. Further investigations are warranted to confirm whether the NADPH oxidase pathway may, indeed, represent a reliable therapeutic target to prevent ischemia/ reperfusion damage in kidney transplantation.

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Fig. 6. α-SMA overexpression induced by ischemia/reperfusion is prevented by C1-INH infusion. In the swine model of ischemia/reperfusion injury α-SMA expression, (A) very limited before ischemia, was widely present within peritubular capillaries and interstitial spaces at (B) T300 and (C) T600 . (D) α-SMA þ cells were also present at the tubular level at T600 . (E) In C1-INH-treated pigs, very few α-SMA þ cells could be observed within the interstitial space under basal conditions. Infusion of C1-INH was able to abrogate interstitial α-SMA expression at (F) T300 and (G) T600 and (H) within the tubules. (I) The quantitative analysis of α-SMA was performed as described under Materials and methods and expressed as the mean 7 SD (T300 CTRL vs T300 C1-INH-treated p ¼0.04, T600 CTRL vs T600 C1-INH-treated p ¼0.02) of at least three pigs for each group. At T600 , (J) the expression of the epithelial marker ZO-1 (red) was limited to a few tubular areas of biopsies, whereas (K) α-SMA tubular expression was increased (green) and (L) the two signals colocalized in some tubules (yellow) as shown by the merge of double immunofluorescence for α-SMA/ZO-1. Original magnification  63. To-Pro-3 was used to counterstain nuclei (blue).

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Fig. 8. NOX-4 silencing reduces C3a-induced NADPH activity and α-SMA induction. (A and B) HK-2 cells were silenced with Qiagen NOX-4_5 siRNA (25 nM) for 24 h. NOX-4 siRNA- and scramble siRNA-transfected HK-2 cells were treated with C3a (5  10  7 M) and NADPH oxidase activity was measured by lucigenin-enhanced chemiluminescence. Scramble siRNA plus C3a vs scramble siRNA, p¼ 0.04; scramble siRNA plus C3a vs NOX-4 siRNA plus C3a, p ¼ 0.03. (C and D) NOX-4 siRNA- and scramble siRNAtransfected HK-2 cells were treated with C3a (5  10  7 M) for 24 h, and α-SMA protein expression was analyzed by immunoblotting.

Acknowledgments This study was supported by an unrestricted research grant from Pharming Group NV, Leiden, The Netherlands, and by a grant from the Ministero della Salute (Ricerca Finalizzata and Giovani Ricercatori 2009 to G.G., G.C., and L.G.). Pharming Group also provided the C1 inhibitor, and one of the authors, B. Oortwijn, is an employee of the company.

References [1] Brook, N. R.; Nicholson, M. L. Kidney transplantation from non heart-beating donors. Surgeon 1:311; 2003. [2] Asher, J.; Wilson, C.; Gok, M.; Balupuri, S.; Bhatti, A. A.; Soomro, N.; et al. Factors predicting duration of delayed graft function in non-heart beating donor kidney transplantation. Transplant. Proc. 37:348; 2005. [3] Devarajan, P. Update on mechanisms of ischemic acute kidney injury. J. Am. Soc. Nephrol. 17:1503–1520; 2006.

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[4] Perico, N.; Cattaneo, D.; Sayegh, M. H.; Remuzzi, G. Delayed graft function in kidney transplantation. Lancet 364:1814–1827; 2004. [5] Li, K.; Sacks, S. H.; Zhou, W. The relative importance of local and systemic complement production in ischemia, transplantation and other pathologies. Mol. Immunol. 44:3866–3874; 2007. [6] Peng, Q.; Li, K.; Smyth, L. A.; Xing, G.; Wang, N.; Meader, L.; et al. C3a and C5a promote renal ischemia–reperfusion injury. J. Am. Soc. Nephrol. 23:1474–1485; 2012. [7] Lee, D. B. N.; Huang, E.; Ward, H. J. Tight junction biology and kidney dysfunction. Am. J. Physiol. Renal Physiol 290:F20–F34; 2006. [8] Strutz, F.; Zeisberg, M.; Ziyadeh, F. N.; Yang, C. Q.; Kalluri, R.; Muller, G. A.; et al. Role of basic fibroblast growth factor-2 in epithelial–mesenchymal transformation. Kidney Int. 61:1714–1728; 2002. [9] Hay, E. D.; Zuk, A. Transformations between epithelium and mesenchyme: normal, pathological, and experimentally induced. Am. J. Kidney Dis. 26: 678–690; 1995. [10] Badid, C.; Desmouliere, A.; Babici, D.; Hadj-Aissa, A.; McGregor, B.; Lefroncois, N.; et al. Interstitial expression of alpha-SMA: an early marker of chronic renal allograft dysfunction. Nephrol. Dial. Transplant. 17:1993–1998; 2002. [11] Ray, R.; Shah, A. M. NADPH oxidase and endothelial cell function. Clin. Sci. 109:217–226; 2005. [12] Babior, B. M.; Lambeth, J. D.; Nauseef, W. The neutrophil NADPH oxidase. Arch. Biochem. Biophys. 397:342–344; 2002. [13] Werns, S. W.; Simpson, P. J.; Mickelson, J. K.; Shea, M. J.; Pitt, B.; Lucchesi, B. R. Sustained limitation by superoxide dismutase of canine myocardial injury due to regional ischemia followed by reperfusion. J. Cardiovasc. Pharmacol. 11:36–44; 1988. [14] Bedard, K.; Krause, K. H. The NOX family of ROS-generating NADPH oxidases: physiology and pathophysiology. Physiol. Rev. 87:245–313; 2007. [15] Groemping, Y.; Rittinger, K. Activation and assembly of the NADPH oxidase: a structural perspective. Biochem. J. 386:401–416; 2005. [16] Sumimoto, H. Structure, regulation and evolution of Nox-family NADPH oxidases that produce reactive oxygen species. FEBS J. 275:3249–3277; 2008. [17] Geiszt, M.; Kopp, J. B.; Varnai, P.; Leto, T. L. Identification of renox, an NAD(P)H oxidase in kidney. Proc. Natl. Acad. Sci. USA 97:8010–8014; 2000. [18] Mkaddem, S.; Pedruzzi, E.; Wertz, C.; Coant, N.; Bens, M.; Cluzeaud, F.; et al. Heat shock protein gp96 and NAD(P)H oxidase 4 play key roles in Toll-like receptor 4-activated apoptosis during renal ischemia/reperfusion injury. Cell Death Differ. 17:1474–1485; 2010. [19] Kahles, T.; Luedike, P.; Endres, M.; Galla, H. J.; Steinmetz, H.; Busse, R.; et al. NADPH oxidase plays a central role in blood–brain barrier damage in experimental stroke. Stroke 38:3000–3006; 2007. [20] Lehnert, M.; Arteel, G. E.; Smutney, O. M.; Conzelmann, L. O.; Zhong, G.; Thurman, R. G.; et al. Dependence of liver injury after hemorrhage/resuscitation in mice on NADPH oxidase-derived superoxide. Shock 19:345–351; 2003. [21] Zhang, Y. S.; He, L.; Liu, B.; Li, N. S.; Luo, X. J.; Hu, C. P.; et al. A novel pathway of NADPH oxidase/vascular peroxidase 1 in mediating oxidative injury following ischemia–reperfusion. Basic Res. Cardiol. 107:266–270; 2012. [22] Castellano, G.; Melchiorre, R.; Loverre, A.; Ditonno, P.; Montinaro, V.; Rossini, M.; et al. Therapeutic targeting of classical and lectin pathways of complement protects from ischemia–reperfusion-induced renal damage. Am. J. Pathol. 176:1648–1659; 2010. [23] Moon, Y. M.; Kang, H. J.; Cho, J. S.; Park, I. H.; Lee, H. M. Nox4 mediates hypoxia-stimulated myofibroblast differentiation in nasal polyp-derived fibroblasts. Int. Arch. Allergy Immunol. 159:399–409; 2012.

273

[24] Liu, Y. Epithelial to mesenchymal transition in renal fibrogenesis: pathologic significance, molecular mechanism, and therapeutic intervention. J. Am. Soc. Nephrol. 15:1–12; 2004. [25] Simone, S.; Cosola, C.; Loverre, A.; Cariello, M.; Sallustio, F.; Rascio, F.; et al. BMP-2 induces a profibrotic phenotype in adult renal progenitors cells through Nox4 activation. Am. J. Physiol. Renal Physiol 303:F23–F34; 2012. [26] Curci, C.; Castellano, G.; Stasi, A.; Divella, C.; Loverre, A.; Gigante, M. Endothelial-to-mesenchymal transition and renal fibrosis in ischaemia/reperfusion injury are mediated by complement anaphylatoxins and Akt pathway. Nephrol. Dial. Transplant. 29:799–808; 2014. [27] Peeters, P.; Vanholder, R. Therapeutic interventions favourably influencing delayed and slow graft function in kidney transplantation: mission impossible? Transplantation 85(Suppl. 7):S31–37; 2008. [28] Garlick, P. B.; Davies, M. J.; Hearse, D. J.; Slater, T. F. Direct detection of free radicals in the reperfused rat heart using electron spin resonance spectroscopy. Circ. Res. 61:757–760; 1987. [29] Henry, T. D.; Archer, S. L.; Nelson, D.; Weir, E. K.; From, A. H. Enhanced chemiluminescence as a measure of oxygen-derived free radical generation during ischemia and reperfusion. Circ. Res. 67:1453–1461; 1990. [30] Kramer, J. H.; Arroyo, C. M. Dickens BF, Weglicki WB. Spin-trapping evidence that graded myocardial ischemia alters post-ischemic superoxide production. Free Radic. Biol. Med. 3:153–159; 1987. [31] Zweier, J. L.; Kuppusamy, P.; Williams, R.; Rayburn, B. K.; Smith, D.; Weisfeldt, M. L.; et al. Measurement and characterization of postischemic free radical generation in the isolated perfused heart. J. Biol. Chem. 264:18890–18895; 1989. [32] Simpson, P. J.; Lucchesi, B. R. Free radicals and myocardial ischemia and reperfusion injury. J. Lab. Clin. Med. 110:13–30; 1987. [33] Ambrosio, G.; Becker, L. C.; Hutchins, G. M.; Weisman, H. F.; Weisfeldt, M. L. Reduction in experimental infarct size by recombinant human superoxide dismutase: insights into the pathophysiology of reperfusion injury. Circulation 74:1424–1433; 1986. [34] Zweier, J. L.; Rayburn, B. K.; Flaherty, J. T.; Weisfeldt, M. L. Recombinant superoxide dismutase reduces oxygen free radical concentrations in reperfused myocardium. J. Clin. Invest. 80:1728–1734; 1987. [35] Chen, J. X.; Zeng, H.; Tuo, Q. H.; Yu, H.; Meyrick, B.; Aschner, J. L. NADPH oxidase modulates myocardial Akt, ERK1/2 activation, and angiogenesis after hypoxia–reoxygenation. Am. J. Physiol. Heart Circ. Physiol 292:H1664–1674; 2007. [36] Dodd, O. J.; Pearse, D. B. Effect of the NADPH oxidase inhibitor apocynin on ischemia–reperfusion lung injury. Am. J. Physiol. Heart Circ. Physiol 279: H303–H312; 2000. [37] Harada, H.; Hines, I. N.; Flores, S.; Gao, B.; McCord, J.; Scheerens, H. Role of NADPH oxidase-derived superoxide in reduced size liver ischemia and reperfusion injury. Arch. Biochem. Biophys. 423:103–108; 2004. [38] Pearse, D. B.; Dodd, J. M. Ischemia–reperfusion lung injury is prevented by apocynin, a novel inhibitor of leukocyte NADPH oxidase. Chest 116(Suppl. 1):55S–56S; 1999. [39] Yang, Z.; Sharma, A. K.; Marshall, M.; Kron, I. L.; Laubach, V. E. NADPH oxidase in bone marrow-derived cells mediates pulmonary ischemia–reperfusion injury. Am. J. Respir. Cell Mol. Biol. 40:375–381; 2009. [40] Venkatachalam, M. A.; Griffin, K. A.; Lan, R.; Geng, H.; Saikumar, P.; Bidani, A. K. Acute kidney injury: a springboard for progression in chronic kidney disease. Am. J. Physiol. Renal Physiol 298:F1078–1094; 2010. [41] Ricklin, D.; Lambris, J. D. Complement-targeted therapeutics. Nat. Biotechnol. 25:1265–1275; 2007.