Macrophages Expressing Heme Oxygenase-1 Improve Renal Function in Ischemia/Reperfusion Injury

Abstract

Acute kidney injury has a high mortality and lacks specific therapies, with ischemia/reperfusion injury (IRI) being the predominant cause. Macrophages (Mφ) have been used successfully in cell therapy to deliver targeted therapeutic genes in models of inflammatory kidney disease. Heme oxygenase-1 (HO-1) catalyzes heme breakdown and has important cytoprotective functions. We hypothesized that administration of Mφ modified to overexpress HO-1 would protect from renal IRI. Using an adenoviral construct (Ad-HO-1), HO-1 was overexpressed in primary bone marrow–derived Mφ (BMDM). In vitro Ad-HO-1 Mφ showed an anti-inflammatory phenotype with increased phagocytosis of apoptotic cells (ACs) and increased interleukin (IL)-10 but reduced TNF-α and nitric oxide (NO) following lipopolysaccharide/interferon-γ (IFNγ) stimulation compared to control transduced or unmodified Mφ. In vivo, intravenously (IV) injected Mφ homed preferentially to the post-IRI kidney compared to uninjured control following experimental IRI. At 24 hours postinjury, despite equivalent levels of tubular necrosis, apoptosis, and capillary density between groups, the injection of Ad-HO-1 Mφ resulted in preserved renal function (serum creatinine reduced by 46%), and reduced microvascular platelet deposition. These data demonstrate that genetically modified Mφ improve the outcomes in IRI when administered after the establishment of structural injury, raising the prospect of targeted cell therapy to support the function of the acutely injured kidney.

Introduction

Acute kidney failure remains a devastating clinical problem with a 90-day mortality exceeding 50%.¹ In the majority of cases, the etiology relates to a toxic or ischemia/reperfusion injury (IRI) to the renal medulla, resulting in acute tubular necrosis (ATN).¹ There is an increasing appreciation of the role of the immune system in modulating severity of injury in experimental models of renal IRI,² with studies demonstrating the potential involvement of Mφ,³ neutrophils,⁴ B lymphocytes,⁵ and T lymphocytes⁶ in determining disease outcome.

The findings of lymphocyte involvement were striking as these cells are neither resident in significant numbers nor infiltrate during the peak injury of renal IRI.⁷ Such contributions to the injury phenotype are likely mediated in part by the intrinsic leukocytes of the kidney—the predominant cell type being the Mφ. There is extensive evidence implicating Mφ in the pathogenesis of human renal disease in both the glomerulus and tubulo-interstitium.⁸ Recent studies have demonstrated that ablation of renal Mφ can improve outcome in models of renal IRI,³ glomerulonephritis,⁹ and fibrosis.¹⁰ Further study of Mφ phenotype has demonstrated that they represent key contributors to the successful resolution of renal inflammation (reviewed in ref. 11).

Monocytes and Mφ are chemotactic to sites of inflammation so are attractive candidates as a cell-based therapy for inflammatory renal disease, after therapeutic manipulation to adopt an “anti-inflammatory” or “pro-repair” phenotype.¹² Modulation of Mφ function can have beneficial effects in a number of renal disease models. Mφ expressing interleukin (IL)-4 or IL-10 ameliorate glomerulonephritis,^13,14 IL-1 receptor antagonist expressing Mφ reduce fibrosis in unilateral ureteric obstruction¹⁵ and injection of IL-4 stimulated Mφ reduce proteinuria in murine adriamycin nephropathy.¹⁶

In addition to the initiation of inflammation, the reperfusion phase of IRI exposes tissues to oxidative stress, generating free heme that potentiates oxidative damage¹⁷ unless promptly conjugated or metabolized. The key enzyme catalyzing heme degradation is heme oxygenase-1 (HO-1), itself induced by hypoxia. In addition to removing a source of oxidative stress, the products of heme metabolism—free iron (Fe²⁺), biliverdin, and carbon monoxide—are all immunomodulatory. HO-1 expression in Mφ opposes inflammatory activation and autoimmunity,^18,19 and improves outcome in both hepatic and renal IRI;^20,21 HO-1 inducing agents have therefore attracted considerable interest as renal cytoprotectants.²² However, in the majority of cases of acute kidney failure, there is no clear “therapeutic window” for pharmacological prophylaxis, and such agents have shown a spectrum of effects including worsened injury in experimental IRI,²³ limiting translational relevance.

This study examined the effect of overexpressing HO-1 within Mφ, on both in vitro phenotype and ability in vivo to improve outcome in established renal IRI.

Results

In vitro

Transduction of Mφ with Ad-HO-1 results in HO-1 overexpression and increased total heme oxygnase enzyme activity. Primary bone marrow–derived Mφ (BMDM) were treated with HO-1-expressing adenovirus (Ad-HO-1). Maximal protein expression without cytotoxicity was seen at a multiplicity of infection of 100 (Figure 1a). HO-1 transduction was associated with increased bioactivity as shown by degradation of heme to bilirubin (Figure 1b).

Figure 1.

Adenoviral transduction results in expression of HO-1 protein in Mφ. (a) Western blotting demonstrates potent induction of HO-1 protein expression in BMDM following increasing multiplicity of infection with Ad-HO-1. (b) HO-1 transduction was associated with increased bioactivity as shown by degradation of heme to bilirubin (n = 8/group). BMDM, bone marrow–derived Mφ.

HO-1 transduction attenuates Mφ inflammatory responses. To confirm the effects of viral transduction with or without HO-1 overexpression on BMDM phenotype, cells were plated and stimulated with lipopolysaccharide and interferon-γ (IFNγ). After 24 hours, production of nitric oxide (NO), TNFα, and IL10 was assessed. Ad-HO-1 treatment resulted in reduced NO release compared to untransduced and Adβgal-transduced cells (Figure 2a). There was also marked suppression of TNFα release in response to stimulation (Figure 2b) with augmented IL-10 secretion (Figure 2c).

Figure 2.

Transduction of HO-1 in Mφ results in altered responses to classical activating stimuli. Transduction with Ad-HO-1 results in significantly reduced (a) Mφ NO and (b) TNFα production, with augmented (c) IL-10 in response to stimulation with IFNγ+LPS (n = 9/group; all P < 0.001 versus all groups by analysis of variance).

HO-1 transduction augments Mφ uptake of ACs. Assays were undertaken to characterize the capacity of the transduced Mφ to phagocytose apoptotic cells (ACs). Ad-HO-1 transduction resulted in augmented phagocytic ability with an increased proportion of Mφ-ingesting ACs (Figure 3a,c; 34.8 ± 2.9% versus 14.1 ± 1.6% Ad-HO-1 versus untransduced control; P = 0.0032). Furthermore, the cells which ingested ACs demonstrated an increased phagocytic index—the mean number of ACs eaten by each phagocytosing Mφ (Figure 3b–d; 1.84 ± 0.14 versus 1.18 ± 0.04 ACs/Mφ Ad-HO-1 versus control; P = 0.0093).

Figure 3.

Transduction of HO-1 in Mφ results in increased phagocytosis of apoptotic cells in vitro. Transduction with Ad-HO-1 results in (a) augmented phagocytosis of apoptotic cells (ACs) (n = 9/group; P = 0.0032) and (b) increased phagocytic index (n = 9/group; P = 0.0093). Phagocytosis was quantified visually using colocalization of Lysotracker Red–labeled Mφ and CM-Green-labeled apoptotic cells to confirm presence of the apoptotic cells within a phagolysosome. Photomicrographs of Mφ following a 30-minute interaction with apoptotic cells [(c) control unmodified Mφ, (d) Ad-HO-1 Mφ phagocytosed ACs shown with white arrows]. BMDM, bone marrow–derived Mφ.

In vivo

Injected Mφ localize preferentially and rapidly to the injured kidney. To examine whether the injected Mφ exhibited the ability to home selectively to the injured kidney in vivo, experiments were conducted with a unilateral clamp to the renal vessels and the contralateral kidney left in situ as an uninjured control. PKH-labeled Mφ were injected intravenously (IV) 20 minutes after unilateral ischemic insult. Localization was assessed on frozen sections at 24 hours postinjury. Greater numbers of PKH⁺ Mφ were seen within the medulla of the injured kidney compared to the contralateral control [Figure 4a; 2.05 ± 0.18 versus 0.90 ± 0.12 PKH⁺ cells/high-power field (hpf); ischemic versus control kidney; P = 0.0048 by paired t-test]. To assess the kinetics of Mφ localization, 5 × 10⁶ HO-1 expressing or control PKH-labeled Mφ were administered IV following 20 minutes warm ischemia of the left kidney with right nephrectomy. Mφ localization was assessed at 1 and 24 hours post-IRI. This demonstrated rapid infiltration 1 hour after administration, persisting at reduced numbers at 24 hours (Figure 4b–d).

Figure 4.

Intravenously administered Mφ home to the injured kidney after IRI. (a) IV administered Mφ localize selectively to the injured kidney compared to the contralateral uninjured control (P = 0.0048, n = 6/group). (b) Maximal localization is seen at 1 hour after injection. PKH⁺ Mφ can be visualized within the interstitium of the injured renal medulla at (c) 1 and (d) 24 hours (original magnification: ×200, blue: DAPI nuclear stain, green: tissue autofluorescence, red: PKH26⁺ Mφ). Medullary F4/80⁺ cell counts (mean per ×400 field) are augmented in animals receiving IV Mφ. (e) Exogenously delivered Mφ were identified by PKH⁺ fluorescence, and comprise between 35 and 58% of total medullary Mφ post-IRI. IRI, ischemia/reperfusion injury.

IV injected Mφ localize within solid organs 24 hours postadministration. Liver, spleen, lung, and kidney were harvested at 24 hours after IRI/cell administration and cell localization quantified on frozen sections (Supplementary Figure S1). Labeled cells were found in all organs examined, with no differences in localization between transduced and nontransduced cells. The greatest cell densities were identified in the spleen and liver (32.4 ± 3.9 versus 29.2 ± 3.5 versus 2.4 ± 0.5 versus 1.1 ± 0.1 PKH⁺ cells/hpf; spleen versus liver versus lung versus kidney).

Administered PKH⁺ labeled Mφ augment renal medullary Mφ number in IRI. Given the low absolute numbers of Mφ localizing within the kidney in the aftermath of IRI, the total number of F4/80⁺ Mφ at 24 hours postinjury was quantified by immunohistochemistry. Medullary Mφ count increased by 40% in cell-treated animals (2.6 ± 0.5 versus 1.5 ± 0.4 F4/80⁺ cells/hpf), corresponding in number to the PKH⁺ cells seen on fluorescent microscopy (Figure 4e).

IV administration of HO-1 overexpressing Mφ results in improved renal function. Histological injury was quantified in the outer medulla of the kidney 24 hours following IRI. This demonstrated equivalent levels of ATN present in all groups (Figure 5a, 54.0 ± 3.6% versus 59.0 ± 3.5% versus 57.9 ± 3.2% versus 59.4 ± 3.7% necrotic tubules; Ad-HO-1 versus Adβgal versus control Mφ versus saline; P = ns). Furthermore, recruitment of Gr1⁺ neutrophils to the injured medulla was comparable in all groups (Supplementary Table S1). Despite this, animals treated with Ad-HO-1 Mφ had significant preservation of renal function after IRI (Figure 5b, 0.70 ± 0.05 versus 1.02 ± 0.15 versus 1.05 ± 0.24 versus 1.30 ± 0.17 mg/dl creatinine; Ad-HO-1 versus Adβgal versus control Mφ versus saline P < 0.05).

Figure 5.

Treatment with Ad-HO-1 transduced Mφ results in functional protection after IRI at comparable levels of acute tubular necrosis. (a) Cell therapy with Ad-HO-1 transduced Mφ after experimental IRI has no impact on severity of tubular necrosis but (b) results in significant preservation of renal function 24 hours after injury (all groups n = 9–12; P < 0.05). IRI, ischemia/reperfusion injury.

IV administration of HO-1 overexpressing Mφ had no effect on endothelial cell integrity and levels of tubular apoptosis post-IRI. As administration HO-1 expressing Mφ resulted in a functional preservation despite comparable levels of ATN, the integrity of the vascular network was assessed using the CD31 endothelial marker. This demonstrated a slight reduction in the amount of CD31⁺ cells present within the outer stripe of the outer medulla 24 hours after IRI, comparable across all experimental groups (Supplementary Table S1). Likewise, there was no difference in the expression of activated caspase-3, a marker of apoptosis within the injured outer stripe of the outer medulla at the same time point (Supplementary Table S1).

IV Administration of HO-1 overexpressing Mφ does not alter systemic cytokine levels. To assess any impact of systemic delivery of Mφ on circulating levels of pro- and anti-inflammatory cytokine release, serum was assayed for levels of IL-6, IL-10, monocyte chemoattractant protein-1, IFN-γ, TNFα, and IL-12p70. No alteration of cytokine profile was seen in any Mφ-treated group compared to injured control animals (Supplemental Table S2, all P = ns).

IV administration of HO-1 overexpressing Mφ results in reduced deposition of platelets and fibrin within the outer medulla after IRI. Given the potential effects of HO-1 activity upon the renal microcirculation, staining was undertaken to quantify platelet and fibrin deposition within the injured kidney (Figure 6a–d). IRI resulted in an 8.9-fold increase in platelet deposition in untreated injured kidney compared to the contralateral nephrectomy specimen 1 hour after injury (P < 0.05). Ad-HO-1 expressing Mφ treatment was associated with striking reductions in medullary platelet deposition 24 hour postinjury (0.17 ± 0.10% versus 4.08 ± 1.55% hpf positive for CD41 staining; Ad-HO-1 versus Adβgal; P = 0.03; Figure 6e). There was also a trend toward reduced fibrin deposition within the medulla, (1.155 ± 0.32% versus 5.65 ± 2.11% hpf positive for fibrin; Ad-HO-1 versus Adβgal; P = 0.08). In order to clarify whether the effects of Ad-HO-1 expressing Mφ reflected reduced deposition or increased resolution, further experiments examined early time points after IRI and cell administration, demonstrating that immediate deposition of platelets is equivalent between groups (P = 0.73).

Figure 6.

Animals treated with Ad-HO-1 transduced Mφ exhibit increased clearance of renal platelet deposition. Treatment with HO-1 expressing Mφ has no effect on immediate levels of platelet deposition compared with control Mφ at 1 hour post-IRI, while resulting in significant reduction in platelet deposition at 24 hours post-IRI [(a) Adβgal Mφ at 1 hour; (b) Ad-HO-1 Mφ at 1 hour, (c) Adβgal Mφ at 24 hours; (d) Ad-HO-1 Mφ at 24 hours] demonstrated in frozen sections stained with CD41 mAb (original magnification: ×200). (e) Mean area of platelet staining was quantified by image analysis and expressed as % hpf positive for CD41. HO-1, heme oxygenase-1; IRI, ischemia/reperfusion injury.

Discussion

These data provide the first demonstration of Mφ-based cell therapy as an intervention for acute kidney failure. There are two key novel findings from this work. First, cell delivered gene therapy for acute kidney failure is practical and can be administered after the initiating injury—a finding of major translational importance. Second, it is demonstrated that despite the presence of established ATN renal function can be improved, likely through enhanced resolution of platelet deposition within the medullary microvasculature.

The in vitro findings suggest a reduction in potential Mφ cytotoxicity through reduced NO and TNFα production. Inducible NO synthase is the predominant source of Mφ NO and has been implicated as deleterious on outcome in renal IRI,²⁴ whereas TNFα release from resident renal F4/80⁺ cells has been identified as an early factor in the pathogenesis of IRI.²⁵ Ad-HO-1-transduced Mφ also had increased IL-10 production, a cytokine reported to improve both histological injury and renal function in IRI when delivered exogenously.²⁶ This must be balanced against the in vivo findings of comparable levels of ATN and activated caspase-3 within the outer stripe of the outer medulla at 24 hours postinjury in all experimental groups—strongly suggesting that the mechanism of functional protection did not reflect any modulation of cell death within the kidney.

The reductions shown in NO and TNFα would indicate that adenoviral HO-1 overexpression opposes the development of the “M1” or classically activated inflammatory Mφ phenotype, echoing recent in vitro work demonstrating augmented M1 responses in murine Mφ from HO-1^−/− animals or after treatment with chemical HO inhibitors.²⁰ Recent work suggests that macrophage phenotypic polarization is a more plastic process than previously realized, with further subdivision of M2 Mφ into YM-1 expressing “wound-healing” and IL-10 secreting “regulatory” Mφ subsets being proposed.²⁷ The large augmentation in IL-10 production seen in the Ad-HO-1-transduced Mφ suggests that HO-1 expression promotes a “regulatory” M2 phenotype.

Ad-HO-1-transduced Mφ phagocytosed more ACs, suggesting a role for HO-1 in determining phagocytic capacity. This builds on work in murine peritonitis, where tHO-1 overexpression resulted in increased phagocytosis of Enterococcus faecalis, with comparable levels of neutrophil recruitment.²⁸ The phagocytic uptake of ACs by Mφ is well recognized to oppose the development of proinflammatory activation,²⁹ and given the high levels of renal cell death present represents an additional mechanism by which HO-1 induction may promote a prorepair phenotype.

In addition to the in vitro phenotype, the studies undertaken in unilateral IRI demonstrated that the manipulated Mφ home selectively to sites of inflammation following systemic administration. This trophic ability has previously been demonstrated in models of experimental glomerular disease.¹⁶ A number of factors may be contributing—renal IRI is characterized by early release of the cytokine TNFα,²⁵ increasing vascular permeability to facilitate leukocyte recruitment. Several chemokines including CCL1, CCL2/monocyte chemoattractant protein-1, CXCL1/KC and CX3CL1 are also known to be released in IRI, and are chemoattractant for monocytes/ Mφ.^30,31,32 In contrast to published studies showing improved outcome with blockade of Mφ recruitment, retaining normal cues for Mφ homing to the injured kidney is critical for the successful application of Mφ cell therapy.

It is of note that the numbers of administered Mφ were maximal in the kidney 1 hour following IRI with a reduction in counts seen at 24 hours. It has previously been demonstrated that resident renal phagocytes leave the kidney for the regional lymph nodes in the aftermath of IRI.³³ Given the augmented phagocytic behavior of the injected cells, it remains possible that a proportion of the introduced Ad-HO-1 Mφ have already taken up apoptotic/necrotic material and left via the lymphatics. Previous studies using Mφ cell therapy targeted specifically to the kidney via renal artery injection have demonstrated significant amelioration of disease with relatively low numbers of labeled cells detectable within the diseased organ.^13,14,15 Furthermore, the protection afforded by renal Mφ ablation in experimental IRI³ indicates that the low numbers of Mφ present within the normal kidney play a key role in the evolution of injury.

The experiments as designed do not allow full dissection of possible paracrine effects from factors secreted by administered Mφ within other organs such as the liver or spleen. Studies inducing systemic overexpression of IL-10 via an adeno-associated virus in rats demonstrated improvement in a remnant kidney model of kidney disease, despite no increase in IL-10 production within the kidney.³⁴ Similarly, systemic treatment of rats with a carbon monoxide-releasing compound before IRI resulted in improved renal function and histology.³⁵ There was, however, no increase in serum IL-10 demonstrated in any Mφ-treated animal compared to injured or uninjured controls—suggesting that systemic IL-10 release is unlikely to be impacting on the kidney by 24 hours postinjury.

These data provide the demonstration of dissociation between structural renal injury and preserved renal excretory function in IRI. This is in striking contrast to the literature in experimental IRI, for example in animals protected through constitutive immunodeficiency,^5,6 where preserved renal function was associated with a large reductions in ATN. The timing of intervention is a key difference from these studies, as cells were administered after injury—an important distinction which could permit translational application in settings such as organ transplantation and critical care.

The survival of the tubular cell has often been considered the key event in determining the onset of renal dysfunction in IRI. Although the fate of the tubular epithelium is clearly important, the aftermath of IRI is also recognized to impact dramatically on the function of the renal microvasculature,³⁶ with paradoxical vasoconstriction, endothelial dysfunction, and persistently reduced renal blood flow following reperfusion.³⁷

Our data demonstrates early platelet deposition after IRI, a phenomenon recognized as an adverse factor in renal transplantation.³⁸ In elegant studies in experimental mesenteric IRI endothelial P-selectin was shown to be required for platelet adhesion.³⁹ In similar experiments using P-selectin KO chimeras, platelet-derived P-selectin was demonstrated as a key neutrophil chemoattractant after renal IRI,⁴⁰ with blockade preserving renal function.⁴¹ Furthermore, activated platelets secrete a litany of proimflammatory chemokines, including CCL1/MIP-1α, CCL5/RANTES, CCL7, and CXCL4.⁴²

With early platelet deposition equivalent across all groups, it seems probable that increased clearance by phagocytic HO-1 overexpressing Mφ results in enhanced clearance and restoration of microvascular patency. Indeed, the Mφ is known to be an effector of platelet clearance in vitro.⁴³ Intriguingly, both monocyte chemoattractants and the administration of exogenous Mφ have been shown to promote the resolution of experimental thrombus formation within blood vessels.^44,45 Carbon monoxide can inhibit platelet aggregation in vitro even in the presence of activating thrombin,⁴⁶ and it remains possible that inhibition of further platelet deposition within the injured organ favors a net loss of platelet microaggregates and thrombus burden. Our findings resonate with recent work in experimental malaria, where increased HO-1 availability resulted in lowered intercellular cell adhesion molecule-1/vascular cell adhesion molecule-1 expression and reduced microvascular congestion.⁴⁷

In the light of these findings, it is proposed that highly phagocytic Mφ overexpressing HO-1 have a beneficial effect through their action within the microvasculature. Modified Mφ promote resolution of platelet deposition and that both local and systemic production of carbon monoxide have the potential to inhibit further platelet aggregation. By restoring perfusion to viable but otherwise hypofunctioning medulla, renal function can be maintained despite equivalent severe levels of structural tubular injury (Figure 7). This work also extends the spectrum of renal diseases for which Mφ therapy has been used successfully from the glomerulus^13,16 to the interstitium. Given the efficacy of Mφ administration even after the initiating renal insult, these data raise the prospect of translational therapies to support the function of the acutely injured kidney.

Figure 7.

Proposed schema of renal dysfunction in the aftermath of IRI. Dotted arrows indicate pathways potentially ameliorated by actions of HO-1 expressing Mφ. IRI, ischemia/reperfusion injury.

Materials and Methods

Materials and reagents. Tissue culture reagents were purchased from Life Technologies (Paisley, UK). Tissue culture plastics were obtained from Costar (Loughborough, UK) and Falcon (Runcorn, UK). All other reagents were from Sigma-Aldrich (Poole, UK) unless otherwise stated.

Preparation of primary BMDM. BMDM were prepared from FVB/nj mice as previously described.⁴⁸ Bone marrow was isolated from femurs using aseptic technique and cultured for 7 days in Teflon-coated pots in Dulbecco's modified Eagle's medium/F12 with 10% heat inactivated fetal calf serum, penicillin (100 U/ml), streptomycin (100 µg/ml) and 20% L929 cell-conditioned medium containing macrophage-colony-stimulating factor.

Recombinant adenoviruses and transfection. Ad-HO-1 was constructed, amplified, and purified on cesium chloride gradient as previously described⁴⁹ with control adenovirus provided by β-galactosidase expressing vector (Adβgal). The absence of replication competent virus was confirmed by plaque assay on HeLa cells.

Measurement of Mφ nitrite and cytokine production. Following transfection, Mφ were plated and selected groups activated with lipopolysaccharide (1 µg/ml) and murine interferon-γ (IFN-γ 100 U/ml). Media was collected and analyzed by specific enzyme-linked immunosorbent assay for IL-10 and TNF-α (both R&D Systems, Abingdon, UK), and by Greiss Assay for nitrite (Promega, Southampton, UK).

Phagocytosis assays. Strain matched thymi were homogenized and the liberated thymocytes stained with Cell Tracker Green CMFDA dye (Invitrogen, Paisley, UK). Cells were suspended in 100 ml RPMI, and rendered apoptotic by overnight culture with the addition of 1 µmol/l dexamethasone. Apoptosis was confirmed by staining with the dyes Trypan Blue and Hoescht 33342 (both Invitrogen). BMDM fed apoptotic thymocytes at a ratio of 10:1 ACs to BMDM. After 1 hour, the wells were washed three times with ice-cold phosphate-buffered saline, and Lysotracker Red (Invitrogen) was added to identify phagolysosomes. Fluorescent images were taken and colocalization between the red phagolysosome and green ACs taken to represent phagocytosis. Results were expressed as number of BMDM ingesting ACs as a proportion of total BMDMs (% phagocytosis), and the mean number of ACs ingested by each phagocytic BMDM (phagocytic index).

Western blotting of HO-1 protein. Mφ were suspended in lysis buffer [20 mmol/l Tris–HCl, 1 mmol/l EDTA (pH 8.0), 150 mmol/l NaCl and 1% (vol/vol) Triton-X] supplemented with commercially available antiprotease tablets (Amersham, Biosciences, Little Chalfont, UK) and spun at 18,000g for 15 minutes, after which the liquid phase was mixed with equal volumes of 2× Laemmli buffer (4% sodium dodecyl sulfate, 20% glycerol, 10% 2-mercaptoethanol, 0.004% bromophenol blue, and 0.125 mol/l Tris–HCl) before heating to 95 °C for 5 minutes. Protein concentrations were estimated by the Bio-Rad protein assay reagent (Bio-Rad Laboratories, Hercules, CA). Western blotting was performed as previously described.⁵⁰ Following electrophoresis onto Hybond-P membranes (Amersham, Biosciences). The membranes were blocked with 5% dried milk then incubated overnight at 4 °C with polyclonal rabbit anti-rat antibody (1:5,000 dilution; Stressgen Biotechnologies, Vancouver, Canada). After washing in Tris-buffered saline+0.1% Tween, the blots were incubated with secondary antibody (horseradish peroxidase–conjugated goat anti-rabbit; Sigma-Aldrich) for 1 hour. Blots were visualized using the ECL Plus system (Amersham Biosciences). Protein loading was confirmed by reprobing the membranes with anti-β-actin antibody (Sigma-Aldrich).

Heme oxygenase bioactivity assay. At 24 hours after transfection, Mφ were harvested and HO activity assays performed as previously described.⁵¹ Briefly, cells were harvested into 100 mmol/l potassium phosphate and 2 mmol/l MgCl₂ containing buffer by scrapin, then pelleted and lysed. A reaction mixture was then created, containing 500 µg of cell protein; 1.5 mg liver cytosol; 0.8 mmol/l NADPH; 2 mmol/l glucose; 6 mmol/l phosphate; 0.2 units glucose 6 phosphate dehydrogenase; and 20 µg hemin, incubated at 37 °C for 1 hour then terminated by adding 400 µl of chloroform and vortexing for 20 seconds, followed by centrifugation at 10,000g for 1 minute. Bilirubin content was measured by spectrophotometry. Results were expressed as picomole of bilirubin formed per milligram protein per hour.

Murine model of ischemia/reperfusion injury. All mice were males aged 6–8 weeks old, on inbred FVB/nj strain background purchased from Harlan (Loughborough, UK). Procedures were performed under Home Office guidelines. Ketamine and metatomidine anesthesia was induced via the intraperitoneal route, with buprenorphine analgesia subcutaneously. A right nephrectomy was performed and the left renal pedicle identified and occluded by atraumatic clamp for 20 minutes. Body temperature was maintained at 35 °C using a homeostatically controlled blanket (Harvard Apparatus, Boston, MA). Following reperfusion the peritoneum was closed with 5/0 suture and the skin with clips, and anesthesia reversed using atipamezole. One milliliter of sterile saline was administered subcutaneously before and after surgery and the animals were maintained in an incubator overnight. Blood and tissue samples were obtained at 1 and 24 hours postsurgery under terminal anesthesia.

Administration and localization of Mφ. Following transfection, Mφ were resuspended and fluorescently labeled using the PKH26-GCL kit. The dye was quenched with fetal calf serum, before washing for three times with phosphate-buffered saline. The cells were then counted and kept on ice until IV injection into animals via the tail vein 20 minutes after restoration of the renal blood supply. Renal lung, liver, and splenic tissue was obtained under terminal anesthesia 24 hours after IRI, and snap frozen in liquid nitrogen before embedding in optimal cutting temperature fixative. The 5-µm frozen sections were examined by fluorescence microscopy to permit counting of PKH⁺ cells. Results were expressed as mean cells per high-power field (×400 magnification), averaged over 5 fields per organ.

Assessment of renal function. Plasma samples were prepared from whole blood, and analyzed by the Jaffe method (Alpha Laboratories, Eastleigh, UK) on a Cobas Fara Centrifugal Analyser (Roche Diagnostics, Burgess Hill, UK) according to the manufacturer's instructions.

Immunohistochemistry. Whole kidneys were cut longitudinally and either snap frozen in liquid nitrogen or fixed in methyl Carnoy's solution (60% methanol, 30% chloroform, and 10% acetic acid) before embedding in paraffin. The 4-µm tissue sections were cut and stained with hematoxylin and eosin for assessment of medullary tubular necrosis. Renal Mφ and dendritic cells were identified by immunostaining for the tissue Mφ/dendritic cell marker F4/80. Embedded tissue was deparaffinized in xylene, rehydrated, and blocked using 3% H₂O₂ before incubation with monoclonal rat anti-F4/80 (1/250 dilution; Caltag Laboratories, Northampton, UK). Neutrophils were identified by nuclear morphology combined with immunostaining for the Gr1 (Ly6c/g) antigen using monoclonal Rat anti-Gr1 (1/250 dilution; Cambridge BioScience, Cambridge, UK). Platelet deposition was identified by immunostaining for CD41, the Integrin-α IIb glycoprotein subunit of the IIb/IIIa complex using monoclonal Rat anti-CD41 (1/250 dilution; AbD Serotec, Oxford, UK). All rat anti-mouse Ab were incubated at 4 °C overnight with subsequent incubation with mouse-adsorbed biotinylated rabbit anti-rat IgG (1/300 dilution; Vector Laboratories, Peterborough, UK) at room temperature for 30 minutes. After washing sections were incubated with Vectastain ABC Elite reagent (Vector Laboratories) for 30 minutes at room temperature, before washing and staining with diaminobenzidine (Dako UK, Ely, UK). Hematoxylin counterstaining was performed before mounting. In all cases, appropriate isotype antibodies were used as negative controls. Mφ and neutrophil counts were expressed as mean cells per ×400 microscope field, with five fields being assessed per area. Tubules within the outer stripe of the outer medulla were photographed and tubules counted as viable or necrotic based on nuclear morphology and integrity of the epithelial cell layer using ImageJ software (Cell_counter plugin; ImageJ 1.36b; National Institutes of Health, Bethesda, MD). Platelet deposition was quantified on ×200 fields, with five replicates being captured before analysis of area of diaminobenzidine positive staining using Colour Range tool on Photoshop CS3 Extended (version 10.0.1; Adobe Systems Europe, Uxbridge, UK). Results were expressed as a proportion of the image positive for CD41.

Immunofluorescence. Frozen tissue was embedded in optimal cutting temperature mounting media and 5 µm sections cut, then fixed in ice-cold acetone. After protein blockade (Spring Bioscience, Pleasanton, CA) primary antibodies (polyclonal rabbit anti-mouse active caspase-3; Abcam, Cambridge, UK, rat anti-mouse CD31; BD Biosciences, Oxford, UK) were added at 1:250 dilution and incubated for 1 hour at room temperature. Following washing in phosphate-buffered saline, secondary antibodies (Alexa-488-conjugated goat anti-rabbit Ig and Alexa-594 donkey anti-rat Ig, both Invitrogen) were added at 1:500 dilution for 1 hour. Slides were coverslipped with Vectashield containing 4′,6-diamidino-2-phenylindole (Vector Laboratories) before microscopy and photography. Image analysis was performed as previously described.

Serum cytokine analysis. Measurement of inflammatory cytokines was undertaken on serum samples taken at the time of killing. Analysis was undertaken using the BD Mouse Inflammation CBA kit to measure IL-6, IL-10, monocyte chemoattractant protein-1, IFN-γ, TNFα, and IL-12p70 on 50 µl of serum as per manufacturers instructions. Sample acquisition was performed on a BD FACSArray instrument.

Statistical analysis. All data are expressed as mean ± SEM. The Student's Unpaired t-test was used to compare two groups. Where multiple conditions were compared one-way analysis of variance for repeated measurements was utilized. P values of <0.05 were taken as representing statistical significance. All statistical analysis was performed using GraphPad Prism, version 4.0c for Macintosh (GraphPad Software, San Diego, CA).

SUPPLEMENTARY MATERIAL Figure S1. Quantification of modified and control Mφ in kidney (A), liver (B), spleen (C) and lung (D). Table S1. Summary of characteristics of murine kidneys and renal function after IRI. Table S2. Measurement of levels of circulating inflammatory cytokines in naïve animals and after IRI ± Mφ administration demonstrates no significant alteration in levels of any measured parameter in animals receiving Mφ IV (all P = ns).

Supplementary Material

Figure S1.

Quantification of modified and control Mφ in kidney (A), liver (B), spleen (C) and lung (D).

Table S1.

Summary of characteristics of murine kidneys and renal function after IRI.

Table S2.

Measurement of levels of circulating inflammatory cytokines in naïve animals and after IRI ± Mφ administration demonstrates no significant alteration in levels of any measured parameter in animals receiving Mφ IV (all P = ns).