Murine renal ischaemia-reperfusion injury (Methods ...

NEPHROLOGY 2008; 13, 390–396

doi:10.1111/j.1440-1797.2008.00979.x

Methods in Renal Research Paper

Murine renal ischaemia-reperfusion injury SEAN E KENNEDY1,2 and JONATHAN H ERLICH2,3 1

Department of Nephrology, Sydney Children’s Hospital, Randwick, 2Prince of Wales Clinical School, Faculty of Medicine, University of New South Wales, Sydney, 3Department of Nephrology, Prince of Wales Hospital, Randwick, New South Wales, Australia SUMMARY: Ischaemia/reperfusion is a major cause of acute kidney injury in native and transplant kidneys and is associated with significant morbidity and mortality. Murine models of renal ischaemia/reperfusion injury have great potential to improve understanding of the underlying processes and are an important focus of ongoing research into therapeutic and preventative strategies. Like all experimental models, murine models of renal ischaemia/reperfusion are prone to significant variability and results may be influenced by a number of technical and design factors. In this article we review the factors that may influence experimental results and provide a guide to conducting reproducible experiments in murine renal ischaemia/reperfusion. KEY WORDS:

acute kidney failure, anaesthetic agents, temperature control, strain variations.

Ischaemia/reperfusion (I/R) injury of native and transplant kidneys is a major cause of acute kidney injury and an important determinant of long-term kidney dysfunction. In recent years there has been a dramatic expansion of knowledge about the pathophysiology of renal I/R injury, much of this derived from studies in animal models. However, despite this growing body of research, the means by which clinicians can detect, prevent or treat I/R injury remain limited. Sound reproducible models of renal I/R are an important step in helping us to further understand the pathophysiology of this condition. The mouse model of warm renal ischaemia and reperfusion shares important similarities with human I/R injury and, through the availability of genetically modified strains, is particularly useful in exploring the molecular and cellular biology of injury. In this article we provide a brief overview of the pathobiology of murine renal I/R injury and describe in detail a step-by-step approach to replicating the model (Boxes 1,2). PATHOBIOLOGY Renal I/R injury is initiated by the cellular depletion of energy substrates during a discrete ischaemic interval. The re-establishment of blood flow (reperfusion) then leads to a Correspondence: Dr Jonathan Erlich, Department of Nephrology, Level 3, High Street Building, Prince of Wales Hospital, Barker Street, Randwick, NSW 2031, Australia. Email: [email protected] Accepted for publication 7 April 2008. © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

Box 1 Instruments and materials Surgical instruments (All instruments should be cleaned and sterilized prior to each procedure.) Scissors Toothed forceps Blunt iris forceps Suture holders Atraumatic microaneurysm clamps Clamp forceps Disposables Sutures (we use 6/0 nylon) Scalpel blades Syringes and fine bore needles Operating bench equipment Thermostatically regulated heating mat Dissecting microscope Overhead lamps Rectal temperature probe and monitor

complex series of events including cytoskeletal disruption, alteration of cellular ionic homeostasis with subsequent oncosis and induction of proteolytic and phospholipolytic pathways, acidosis, production of reactive oxygen and nitrogen species, generation of inflammatory mediators, leucocyte infiltration, microvascular reactivity, and, depending on the severity of the insult, ultimately cell death by either necrosis or apoptosis [Reviewed by Bonventre & Weinberg1].

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Murine renal ischaemia-reperfusion injury

Box 2 Step-by-step guide 1. The anaesthetized mouse is placed supine upon the heated operating surface. 2. The skin of the abdomen is washed with 70% alcohol solution. 3. 300 mL of 0.9% saline is given subcutaneously in the upper thigh. 4. A midline incision is made through the skin and then through the peritoneum along the linea alba. 5. The descending colon and spleen are carefully moved to the right of midline to allow exposure of the left kidney. 6. The left renal pedicle is bluntly dissected from underlying adipose tissue and, using forceps, an atraumatic microaneurysm clamp is applied. The kidney is briefly observed for colour change. 7. The spleen is returned to its bed. The transverse and descending colon, plus small bowel as required, are moved to the left to allow exposure of the right kidney. The right lobe of the liver may need to be gently lifted to expose the renal pedicle. 8. The right renal pedicle is dissected and clamped as per the left (Step 6). 9. All abdominal contents are returned to the abdomen and the wound is sutured close during the ischaemic interval. With proper preparation and experience both renal pedicles can be clamped within 3 minutes of each other. 10. The mouse is observed for the ischaemic interval. Stable body temperature is crucial during ischaemia. 11. At the end of the ischaemic interval the wound is reopened and the left and right clamps are removed in turn. Both kidneys are observed for return of perfusion. 12. The abdomen is sutured closed. 13. 800 mL of 0.9% saline is given subcutaneously in the other upper thigh.

Studies in mice have mainly focused on acute kidney injury after bilateral warm ischaemia. This results in a characteristic appearance of necrosis of tubular epithelial cells and an interstitial inflammatory cell infiltrate, both developing within the first 24-hours after reperfusion. The tubular necrosis, which in mice is more marked than is usually seen in human biopsy specimens,2 is most evident in the outer medulla where hypoxia is greatest. The interstitial inflammatory cells are predominately polymorphonuclear cells; however, numerous studies have suggested an important role for lymphocytes and macrophages in I/R injury.3,4 The severity of injury in mice is primarily influenced by the duration of ischaemia. Most published studies describe a period of acute reduction in glomerular filtration as shown by a 7- to 10-fold increase in serum creatinine peaking at 24 to 48 hours after reperfusion and return to baseline 1 to 2 weeks after non-lethal injury.5 Prolonged bilateral ischaemia, generally greater than 45 minutes in mice, is © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

likely to cause frank necrosis and a high death rate from complications of renal failure.6 Studies in rats have shown that after weeks or months, surviving animals may develop chronic kidney injury marked by impaired urinary concentrating ability and a gradual increase in tubulo-interstitial fibrosis.7,8 Nephron reduction and hyperfiltration after unilateral nephrectomy appear to accelerate the long-term injury.9 The long-term effects of renal I/R in mice have not been extensively characterized, in one study from Park, Bonventre and co-workers. There was evidence of persistent tubular injury and early fibrotic changes in kidneys of mice 3–12 weeks after 30 minutes of bilateral ischaemia.5 In order to accelerate injury other investigators have inflicted severe (60 minutes of ischaemia) unilateral injury leading to marked fibrosis and albuminuria at 30 days after ischaemia and beyond.10,11 In general it seems that the degree of initial injury, the presence of any other previous injury and the genetic background of the animal will be important for determining whether any long-term renal injury will develop. Studies of renal I/R injury in mice have generally been limited to warm ischaemia. The more challenging technique of orthotopic transplantation has also been established.12 Caution must be employed when translating results in mice to other species as experience has shown that successful therapies in rodent studies have not been beneficial in clinical trials.13 There are clear structural, haemodynamic and molecular differences between kidneys of rodents, larger animals and humans which may determine how a given species responds to I/R.2 For example, tissue factor, a proinflamatory molecule that has been shown to contribute to renal I/R injury,14 is normally expressed in the tubules but not glomeruli of mice; in contrast, rabbits and humans express tissue factor in their glomeruli but not in tubules.15,16 However, following inflammatory stimuli mice and rabbits both upregulate tissue factor expression and both animal species have provided useful models to help understand inflammatory renal disease in humans. The pig kidney may be more similar to the human kidney than that of mice, rats and dogs and, according to several criteria, may provide a better model of human disease.2 However, there are several limitations of larger animal models in terms of expense, convenience and availability and they do not currently have the same degree of genetic modification offered by mice. Taking account of the known differences one can still potentially learn important lessons from small animals about relevant cellular and molecular pathways that lead to disease. It is likely that mice and rats will remain for some time as the basic building block of in-vivo studies and that researchers will seek to corroborate findings in rodents with experiments in pigs.17

GENERAL METHODOLOGICAL CONSIDERATIONS Over the last decade there have been over 150 original studies indexed in Medline of renal I/R employing murine models. Several different centres have described models

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with slight modifications in methodology. At the outset it is important to realize that subtle alterations in surgical approach and perioperative care of animals may lead to divergent results. Thus, as with any laboratory technique, it is imperative that individual researchers validate the model under their own conditions. Further conclusions drawn are valid for the particular model with a particular set of conditions and reagents. Great care needs to be applied in generalizing findings. We will provide a discussion of issues that may influence the outcome of the model and present a step-by-step guide based on our established methodology.14

SE Kennedy and JH Erlich

Total vascular occlusion can be readily confirmed by the kidney becoming engorged and taking on a mottled then purplish hue within 5 minutes. After 15 to 20 minutes the kidney will be deeply cyanosed (Fig. 1). Failure of the kidney to become uniformly purple suggests incomplete occlusion. In our experience, relative sparing of a portion of kidney will result in a significant lessening of injury, and such animals should be excluded from analysis. Rapid return of pink colouration is seen within minutes of clamp removal (Fig. 1). Failure of this to occur suggests thrombosis of the artery or vein or renal infarction. In our experience using atraumatic clamps and limiting ischaemia time to 25 to 30 minutes, kidneys rarely fail to reperfuse fully.

SURGICAL APPROACH The renal arteries can be approached by either a midline laparotomy or by flank incisions. Both techniques have been well described and are equally valid; however, they may lead to slightly different results and caution should be exercised when comparing outcomes of different studies. The midline approach is relatively simple and allows rapid exposure of both renal pedicles, but it is potentially associated with greater fluid loss via evaporation from abdominal contents and the handling of bowel may predispose to a more marked systemic inflammatory response. Despite these risks, we have found that, with gentle handling and careful attention to asepsis, temperature control and fluid replacement (as described below), the model based on a midline approach leads to reproducible injury with low perioperative mortality. A further issue with surgery is whether a unilateral nephrectomy is performed prior to initiating ischaemia in the contralateral kidney. Overall there seems little difference in the pattern of outcomes in the first 24 to 48 hours. Unilateral nephrectomy may alter results of longer-term studies when nephron reduction becomes more relevant.

CLAMPING OF PEDICLES In larger animals the renal artery is often dissected out from the vein and surrounding tissue before occlusion. Such a dissection in mice is technically challenging and most models describe clamping of the entire renal pedicle including artery, vein and nerves. Attempts to isolate the renal artery may well result in sympathetic nerve damage or functional sympathectomy. As sympathectomy in itself may influence outcome,18 anything that inconsistently effects sympathetic nerve function may provide an extra degree of variability. To most effectively and reproducibly place clamps we have found that it is necessary to dissect away some of the perihilar adipose tissue before the pedicles are clamped using atraumatic microaneurysm clamps. These clamps exert enough pressure to fully occlude the artery and vein but do not cause significant endovascular trauma. Their attachment and removal is facilitated by use of specially designed forceps. Vascular ties/sutures are more likely to lead to endovascular damage and potentially impair reperfusion.

DURATION OF ISCHAEMIA Most published studies of bilateral renal I/R in mice have used between 22 and 35 minutes of ischaemia. We have found in C57/BL6 mice that 25 minutes of ischaemia produces reproducible renal injury within the first 24 hours. Shorter ischaemic intervals were associated with a more varied response while more severe ischaemia increased the risk of death. As discussed below, some strains of mice are less susceptible to I/R injury and may therefore require longer ischaemia to attain equivalent degrees of injury.19 THE OPERATING ENVIRONMENT Adequate illumination is imperative for the comfort of the surgeon as well as for the sake of precision. Overhead lights may also be a source of heating. We have found that an operating microscope is invaluable for providing magnification. TEMPERATURE CONTROL Under general anaesthesia mice loose the ability to thermoregulate and may rapidly become hypothermic. In a recent study of ketamine induced anaesthesia for laparotomy, the body temperature of mice that were not warmed fell by 5°C (from 36°C to 31°C) 10 minutes after induction of anaesthesia.20 We have seen a similar degree of cooling in mice under inhalational anaesthesia. The benefits of hypothermia in slowing metabolic processes and reducing ATP depletion are well recognized and form the basis of cold preservation of kidney transplants.21,22 It is therefore not surprising that hypothermia during experimental I/R is associated with some protection from renal injury.23 Conversely, hyperthermia may predispose to cytokine release, more severe kidney failure and increased mortality, as is seen in heat stress.23 For these reasons it is critically important that body temperature of mice is monitored and regulated during I/R. We, like most authors in the field, use a thermostatically controlled, heated operating table and overhead lamps to maintain rectal temperature about 37°C throughout surgery, the ischaemic interval and during the early stages of recovery. Maintaining the ambient room temperature between © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

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21°C and 27°C assists in preserving perioperative body temperature and allowing subsequent recovery. In general, we have found that if the body temperature falls below 35°C during surgery the results become unreliable and if it rises above 38°C there is a higher mortality at 24 hours. We therefore don’t analyse animals where this has happened. It should be noted, however, that some researchers maintain the body temperature of mice at 39°C during surgery.13 Such variations need to be taken into consideration when comparing the experimental results from different laboratories.

FLUID MANAGEMENT Fluid shifts and loss of body water may occur during general anaesthesia and abdominal surgery in mice. Coupled with reduced postoperative drinking and the potential for a postischaemic diuresis, experimental mice are at risk for intravascular volume depletion, which in turn, may worsen kidney injury. It is for this reason that parenteral fluid is given perioperatively. We routinely give 1100 mL of normal saline subcutaneously during the surgical procedure. Even with this support we have seen that C57/BL6 mice loose on average 9% of their body weight 24 hours after reperfusion. To avoid cooling, any fluid or drug administered should be warmed, at least to room temperature and to body temperature if large volumes are infused. Weighing of mice before and after surgery should be considered for the purposes of monitoring fluid status and wellbeing. Again it is important to note that seemingly minor variations in technique may alter the effects of I/R injury. For instance, the volume and route of fluid administration may influence injury; preliminary experiments from our laboratory have shown that giving the same volume of normal saline intravenously leads to significantly improved renal function at 24 hours compared with the subcutaneous route.

OXYGEN THERAPY As we use an inhalational anaesthetic agent, we mix the agent with 100% oxygen at 2 L/min. By using pulse oximetry we have seen that this approach maintains an oxygen saturation of 98% to 100% during surgery. This is done to ensure that we do not add systemic hypoxia to either the pre surgical set up or during the procedure. The precise relevance of this to injury is unclear; however, ischaemic preconditioning has been suggested as a mechanism to protect organs. Certainly this is one aspect of this model that researchers should consider carefully. 䉴

Fig. 1 Predictable colour change of murine kidneys during ischaemia and reperfusion. (A). Left kidney and renal pedicle (arrow) exposed via a midline laparotomy. (B). Same kidney with atraumatic clamp (*) on pedicle showing deep cyanosis after 15 minutes of total ischaemia. (C). Same kidney showing return of uniform pink colour 5 minutes after clamp removal. © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

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ANAESTHESIA AND PAIN CONTROL All procedures on laboratory animals should be as humane and pain-free as possible. General anaesthesia is required for renal I/R whether by the flank or abdominal approach. Further it is often recommended that local anaesthetic be infiltrated into an area where an incision is to be located. If this is done great care is needed to avoid toxicity as most local anaesthetics have increased toxicity in the presence of renal failure. The choice of anaesthetic agent is wide. Many groups use thiopentone, usually administered by the intraperitoneal route. Ketamine and xylesine are also used. We have chosen to use the inhalational agent isoflurane. This allows induction of anaesthesia prior to any painful procedure and by using an anaesthetic machine, we can titrate anaesthesia during the procedure. Recovery from anaesthesia is also relatively rapid and this facilitates post op monitoring. We also administer intraperitoneal midazolam to support anaesthesia. A recently published study showed that volatile anaesthetics are associated with less renal I/R injury and inflammation in mice than barbiturates, an effect that is dependent upon sphingosine kinase.24,25 Within the group of barbiturates there may also be significant differences in IR injury.26 Propofol, another injectable anaesthetic agent, has been shown to have beneficial effects in renal I/R in rats and pigs but there are no studies in mice.27 Studies of I/R in kidneys and other organs suggest propofol may induce heme oxygenase production, stabilize mitochondrial function and reduce apoptosis and lipid peroxidation, all of which may contribute to organ protection following reperfusion injury.27–30 POSTOPERATIVE CARE Following surgery mice should be cared for and closely observed in a quiet, warm environment with free access to food and water. Any animals that are obviously suffering should receive analgesia (buprenorphine) or be euthanized. As animals may have renal failure care in dosing of analgesics is important so not to cause undue sedation or other toxicity. The use of non-steroidal anti-inflammatory drugs should be avoided as they are likely to exacerbate renal failure in animals undergoing renal I/R. EUTHANASIA In general, renal I/R experiments in mice are terminated by humane euthanasia to allow collection of plasma, kidneys and any other samples. MEASUREMENT OF INJURY The two primary indicators of renal injury used in animal I/R experiments are increased serum creatinine and histopathological evidence of tubular injury. Serum or plasma creatinine is the most frequently used indicator of reduced glomerular filtration rate in murine renal I/R injury and within the first 48 hours post injury is a reasonably reliable


indicator of injury. The creatinine levels of normal adult mice are in the vicinity of 10–30 mmol/L. The method of assessment of creatinine is important. Colorimetric methods are the most convenient but tend to give higher readings. High performance liquid chromatography (HPLC) methods are probably most accurate around the normal range reading 8–10 mmol/L while colorimetric methods for the same mice may read higher values 15–25 mmol/L. Most investigators use colorimetric methods and it is important to appreciate small increases will be less reliably measured than with HPLC. However, larger increase is easily detected. As long as consistent and similar measurement techniques are employed, results can be potentially compared. Histopathological evidence is usually quantified by scoring severity of injury seen on haematoxylin and eosin (H&E) sections after paraffin embedding. A variation of a grading system whereby injury is scored on bivalved renal sections in a blinded fashion according to a semiquantitative scale is most often used.19 Higher scores represent more severe damage, with the maximum score being four or five. Cell accumulations have also been assessed by techniques such as immunostaining or tissue digestion and subsequent flow cytometry (usually looking for T cells).31,32 Other techniques have included measurement of tissue myeloperoxidase (MPO) activity as a marker of neutrophil infiltration. There is considerable MPO activity detectable in normal kidneys; however, MPO has been widely used as a marker of tissue neutrophil infiltration and in our experience correlates well with other markers of tissue neutrophil infiltration including counting of neutrophils identified by morphological criteria on H&E stained sections and quantification of immunostained tissue sections with anti-neutrophil antibodies. In all cases the technique is only as good as the tissue sampling and the evenness of the original IR injury. When collecting tissue for cellular or biochemical analysis it is important to note where it is from and consistently compare like areas. The pattern of cellular infiltration and injury varies by region in the kidney so inconsistent sampling or incorrect comparisons could lead to erroneous conclusions.

ETHICAL CONSIDERATIONS As with all animal experiments there are important ethical considerations. This model addresses important medical questions and cannot easily be reproduced in vitro necessitating the use of animals. Renal I/R involves major surgery and hence care and attention should be paid to the choice of operative technique, anaesthesia and analgesia. It is important mice are not stressed prior to surgery, are allowed to acclimatize to their surroundings prior to surgery and are carefully maintained postoperatively. This is both an important part of ethical practice and good scientific practice to ensure reliable reproducible results. Animal group sizes should always be planned such that there is sufficient statistical power to determine likely differences. © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

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SELECTION OF MICE Non-technical issues may influence the degree of renal injury observed after I/R. Perhaps the two most important of these issues are to do with the choice of experimental mice, specifically genetic background and gender. Strain differences in response to renal I/R injury are well established. Of the commonly used strains, C57/BL6 mice are more prone to injury after I/R while NIH Swiss mice are relatively protected and BALB/c mice have an intermediate phenotype.33,34 Our understanding of the basis of these differences is incomplete. Recent research has shown that factors such as the Th-phenotype can influence I/R injury.34 Broadly speaking, renal I/R is a Th-1 response, which may explain the preponderance to injury of C57/BL6 mice as they demonstrate a Th-1 phenotype.35 Given these genetic differences, the choice of control mice is especially important in studies of genetically modified animals. Renal I/R injury also tends to be more severe in male compared with female mice.36,37 Work from Park and Bonventre suggest that this is due to testosterone, possibly via inhibition of protective mechanisms such as Heat Shock Proteins and manganese superoxide dismutase.36,37 The age of experimental animals may also influence the severity of injury with younger animals being potentially less severely affected.38 A further issue is the source and health status of mice used in studies. Mice should be free of any obvious external signs such as cuts, scratches and areas of hair loss that might indicate underlying pathology. Another potential consideration is the housing of the mice; specific pathogen free versus conventional animal handling. Underlying asymptomatic infections or differences in intestinal flora might effect the outcome of reperfusion models.39 One should thus take care when comparing mice from different suppliers and ensure an adequate acclimatization period. CONCLUSION Models of murine renal I/R will continue to be of great importance in gaining further understanding of the major cause of acute kidney injury in humans. The availability of gene knockout and transgenic mice provides a powerful tool for exploring complex pathways with the long-term goal of developing treatments. The basic methods of I/R are fairly straightforward, requiring uncomplicated surgical skills and relatively inexpensive equipment. However, the model is prone to marked variation if attention is not paid to such details as temperature control and fluid therapy. Furthermore, careful experimental design is necessary to allow selection of the most appropriate experimental and control groups, as well as such critical procedural aspects as ischaemic time. ACKNOWLEDGEMENTS The authors would like to thank their colleagues Dr Jacob Sevastos, Melissa Sam and Professor Margaret Rose for their © 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology

invaluable work in establishing and refining the experimental model. Dr Kennedy is a recipient of an NHMRC biomedical scholarship. REFERENCES 1. Bonventre JV, Weinberg JM. Recent advances in the pathophysiology of ischemic acute renal failure. J. Am. Soc. Nephrol. 2003; 14: 2199–210. 2. Lieberthal W, Nigam SK. Acute renal failure. II. Experimental models of acute renal failure: Imperfect but indispensable. Am. J. Physiol Renal Physiol. 2000; 278: F1–12. 3. Friedewald JJ, Rabb H. Inflammatory cells in ischemic acute renal failure. Kidney Int. 2004; 66: 486–91. 4. Day Y, Huang L, Ye H, Linden J, Okusa M. Renal ischemiareperfusion injury and adenosine 2A receptor-mediated tissue protection: Role of macrophages. Am. J. Physiol. Renal Physiol. 2005; 288: F722–31. 5. Park KM, Byun JY, Kramers C, Kim JI, Huang PL, Bonventre JV. Inducible nitric-oxide synthase is an important contributor to prolonged protective effects of ischemic preconditioning in the mouse kidney. J. Biol. Chem. 2003; 278: 27256–66. 6. Heemann U, Szabo A, Hamar P et al. Lipopolysaccharide pretreatment protects from renal ischemia/reperfusion injury: possible connection to an interleukin-6-dependent pathway. Am. J. Pathol. 2000; 156: 287–93. 7. Forbes JM, Hewitson TD, Becker GJ, Jones CL. Ischemic acute renal failure: Long-term histology of cell and matrix changes in the rat. Kidney Int. 2000; 57: 2375–85. 8. Basile DP, Donohoe D, Roethe K, Osborn JL. Renal ischemic injury results in permanent damage to peritubular capillaries and influences long-term function. Am. J. Physiol. Renal Fluid & Electrolyte Physiol. 2001; 281: 887–99. 9. Azuma H, Nadeau K, Takada M, Mackenzie HS, Tilney NL. Cellular and molecular predictors of chronic renal dysfunction after initial ischemia/reperfusion injury of a single kidney. Transplantation 1997; 64: 190–97. 10. Burne-Taney M, Yokota N, Rabb H. Persistent renal and extrarenal immune changes after severe ischemic injury. Kidney Int. 2005; 67: 1002–9. 11. Shimoda N, Fukazawa N, Nonomura K, Fairchild R. Cathepsin g is required for sustained inflammation and tissue injury after reperfusion of ischemic kidneys. Am. J. Pathol. 2007; 170: 930–40. 12. Han W, Murray-Segal L, Mottram P. Modified technique for kidney transplantation in mice. Microsurgery 1999; 19: 272–4. 13. Daemen M, van’t Veer C, Denecker G et al. Inhibition of apoptosis induced by ischemia-reperfusion prevents inflammation. J. Clin. Invest. 1999; 104: 541–9. 14. Sevastos J, Kennedy SE, Davis DR et al. Tissue factor deficiency and PAR-1 deficiency are protective against renal ischemia reperfusion injury. Blood 2007; 109: 577–83. 15. Luther T, Flossel C, Mackman N, et al. Tissue factor expression during human and mouse development. Am. J. Pathol. 1996; 149: 101–13. 16. Erlich J, Fearns C, Mathison J, Ulevitch R, Mackman N. Lipopolysaccharide induction of tissue factor expression in rabbits. Infect. Immun. 1999; 67: 2540–46. 17. Jayle C, Milinkevitch S, Favreau F et al. Protective role of selectin ligand inhibition in a large animal model of kidney ischemiareperfusion injury. Kidney Int. 2006; 69: 1749–55. 18. Ogawa T, Mimura Y, Kaminishi M. Renal denervation abolishes the protective effects of ischaemic preconditioning on function and haemodynamics in ischaemia-reperfused rat kidneys. Acta Physiol. Scand. 2002; 174: 291–7.

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© 2008 The Authors Journal compilation © 2008 Asian Pacific Society of Nephrology