Secondary Logo

Journal Logo

Caspase Inhibition During Cold Storage Improves Graft Function and Histology in a Murine Kidney Transplant Model

Nydam, Trevor L., MD1; Plenter, Robert, BS1; Jain, Swati, MD1; Lucia, Scott, MD1; Jani, Alkesh, MD1,2

doi: 10.1097/TP.0000000000002218
Original Basic Science—General
Free
SDC

Background Prolonged cold ischemia is a risk factor for delayed graft function of kidney transplants, and is associated with caspase-3–mediated apoptotic tubular cell death. We hypothesized that treatment of tubular cells and donor kidneys during cold storage with a caspase inhibitor before transplant would reduce tubular cell apoptosis and improve kidney function after transplant.

Methods Mouse tubular cells were incubated with either dimethyl sulfoxide (DMSO) or Q-VD-OPh during cold storage in saline followed by rewarming in normal media. For in vivo studies, donor kidneys from C57BL/6 mice were perfused with cold saline, DMSO (vehicle), or QVD-OPh. Donor kidneys were then recovered, stored at 4°C for 60 minutes, and transplanted into syngeneic C57BL/6 recipients.

Results Tubular cells treated with a caspase inhibitor had significantly reduced capsase-3 protein expression, caspase-3 activity, and apoptotic cell death compared with saline or DMSO (vehicle) in a dose-dependent manner. Treatment of donor kidneys with a caspase inhibitor significantly reduced serum creatinine and resulted in significantly less tubular cell apoptosis, BBI, tubular injury, cast formation, and tubule lumen dilation compared with DMSO and saline-treated kidneys.

Conclusions Caspase inhibition resulted in decreased tubular cell apoptosis and improved renal function after transplantation. Caspase inhibition may be a useful strategy to prevent cold ischemic injury of donor renal grafts.

The authors investigate the efficacy of caspase inhibitor on improvement of graft function and attenuation of tubular cell apoptosis in a mouse kidney transplantation model (in vivo) as well as mouse tubular cells (in vitro).

1 Division of Renal diseases and Hypertension, University of Colorado Anschutz Medical Center, Aurora, CO.

2 Denver Veterans Affairs Medical Center Denver, CO.

Received 28 August 2017. Revision received 24 January 2018.

Accepted 30 January 2018.

T.L.N., R.P., and S.J. contributed equally to this work.

This work was supported by a T32 award 5T32DK007, in addition to a 135 VA Merit Award 1I01BX001737 to AJ and the 2014 American Society of Transplant Surgeons-Astellas Faculty Development Award to TN.

The authors declare no conflicts of interest.

T.N. participated in the performance of the research, research design, writing the article, and data analysis. R.P. participated in performance of the research, writing the article, and data analysis. S.J. participated in performance of the research, writing the article, and data analysis. S.L. participated in data analysis. A.J. participated in performance of the research, research design, writing the article, and data analysis.

Correspondence: Alkesh Jani, MD, Division of Renal Diseases and Hypertension, University of Colorado Denver, 12700 East 19th Ave, C281, Aurora, CO 80045. (Alkesh.jani@ucdenver.edu).

Supplemental digital content (SDC) is available for this article. Direct URL citations appear in the printed text, and links to the digital files are provided in the HTML text of this article on the journal’s Web site (www.transplantjournal.com).

Deceased donor kidneys are subjected to a period of cold storage (CS), referred to as cold ischemia (CI), before transplant. During CI, the donor kidney is cooled to approximately 4°C to delay ex vivo cell death. Cold storage (either by static CS or machine perfusion) is used in the majority of deceased donor kidney transplants performed at centers in Europe, North America, and Australia.1 Static CS is still the most common form of human kidney preservation, although machine perfusion has experienced a recent resurgence in use.2

The beneficial effects of CI are limited and deceased-donor kidneys cannot be maintained at 4 °C indefinitely. Several studies over many decades have identified prolonged CI time (CIT) as a risk factor for the development of delayed graft function (DGF).3-8 DGF refers to acute kidney injury (AKI) after transplantation and is typically defined as the need for dialysis within the first week posttransplant.9 For every 6-hour increment of CI, the risk of DGF increases by 23%8 and a CIT longer than 36 hours significantly increases the risk of DGF.10,11 The mechanism by which prolonged CI leads to DGF is not known.

Caspase-3 is regarded as the “executioner caspase” and is centrally important in apoptotic cell death.12 The rate of apoptosis in kidney tubular cells has been shown to correlate significantly with CIT in human deceased donor kidney transplants.13 Biopsies of human donor kidneys indicate that increased apoptosis in renal tubular epithelial cells (RTEC) predicts the development of DGF after transplantation.14 Our previous work demonstrates that prolonged CS results in caspase-3 activation, tubular injury and apoptosis in an ex-vivo model of kidney CS in mice.15 Treatment of cold stored kidneys with a pan-caspase inhibitor, Q-VD-OPh, prevented cold-storage associated tubular cell apoptosis and brush border injury (BBI). However, it is not known whether Q-VD-OPh protects cold stored donor kidneys from subsequent renal injury after transplantation.

We hypothesized that treatment of a donor organ with Q-VD-OPh before kidney transplantation would be associated with significantly reduced RTEC apoptosis, BBI and improved graft function in a mouse kidney transplant model.

Back to Top | Article Outline

MATERIALS AND METHODS

In vitro CS/rewarming model

M-1 (ATCC CRL-2038) RTECs were subjected to CS in cold saline for 24 hours at 4°C and rewarmed (REW) in normal media at 37°C for 24 hours as previously described.16-18 Renal tubular epithelial cells were incubated with either dimethyl sulfoxide (DMSO) or a caspase inhibitor Q-VD-OPh (R&D systems, OPH001) at a final concentration of 50 μM during CS/REW. Q-VD-OPh was reconstituted in DMSO. A dose-response analysis (5-50 μM) was performed to determine the optimal concentration of Q-VD-OPh to completely inhibit active caspase-3 (Figure 1).

FIGURE 1

FIGURE 1

Back to Top | Article Outline

Cell lysis and Immunoblot Analysis

Total cellular extracts were prepared by washing cells in ice-cold phosphate-buffered saline twice and then lysing cells in radioimmunoprecipitation assay buffer buffer containing a protease inhibitor cocktail as previously described.16-18

Back to Top | Article Outline

Flow Cytometry

Dead Cell Apoptosis Kit (Life Technologies, V13241) with Alexa Fluor 488 annexin V and PI was used to quantify cell death in M-1 RTECs, induced by CS and REW as previously described.16,18 Untreated cells were taken as negative controls. All experiments were independently performed at least 4 times.

Back to Top | Article Outline

Caspase-1 and Caspase-3 Activity

A colorimetric kit was used to determine caspase-3 activity (Biovision, K106) and caspase-1 activity (Biovision, K111) as previously described.16-18

Back to Top | Article Outline

Animals and Study Approval

Inbred male C57Bl/6J mice were purchased from The Jackson Laboratory (Bar Harbor, ME) and were housed under pathogen-free conditions at the University of Colorado Denver, Barbara Davis Center Animal Facility according to NIH guidelines and with approval of the University of Colorado Denver IACUC.

Back to Top | Article Outline

Kidney Transplant

We have previously demonstrated that recipients of grafts subjected only to a kidney transplant without CI have low SCr (0.4 mg/dL) and minimal histological injury in the form of mild BBI (acute tubular necrosis [ATN] score, 1; apoptosis score, 0).19 Kidney transplants were performed as previously described19,20 except that CI was now added as a component. Briefly, during graft recovery the donor was perfused with either: (1) 4°C saline alone, (2) DMSO in 4°C saline, or (3) DMSO and Q-VD-OPh (100 μM) in 4°C saline. DMSO was needed to ensure solubility of the Q-VD-OPh. Since DMSO has been shown to have anti-inflammatory effects,21,22 we included a saline-treated group to remove any confounding by the potentially protective effects of DMSO and to demonstrate the effect of pure CI on graft function and histology. The left kidney, vessels, and ureter were then removed and stored in either 4°C saline alone, DMSO in 4°C saline, or DMSO and Q-VD-OPh (100 μM) in 4°C saline for a total CI time of 60 minutes. Before the end of the CS period, the recipient mouse was prepared. After right recipient nephrectomy, the donor kidney was placed in the right flank and the arterial and venous cuffs were anastomosed to the recipient aorta and inferior vena cava, respectively. The recipient bladder was prepared, and the ureter was anastomosed to the bladder according the procedure developed by Han et al.23 Total warm ischemia time was approximately 35 to 38 minutes. To ensure survival of the animal, the left native kidney was left in place. Bilateral native nephrectomy at the time of transplant was not performed, because it leads to significant loss of animals.24,25 On posttransplant day 7, a left recipient nephrectomy was performed, leaving the recipient with only the transplanted kidney. After left native nephrectomy, the recipient mouse is dependent only on the transplant kidney and can survive longer than 30 days (data not shown). To determine the effects of Q-VD-OPh on renal function and histology in the early posttransplant period, serum creatinine was measured, and the transplanted kidney was removed on posttransplant day 8. The kidney transplant was then immediately sectioned for snap freezing in liquid nitrogen or fixation in 10% phosphate-buffered formalin.

Back to Top | Article Outline

Renal Function

Serum creatinine was measured using a creatinine enzymatic kit (Pointe Scientific, C7548).

Back to Top | Article Outline

Renal Histology

Renal histology was assessed as previously described.15,17,26-28 All histological parameters were assessed by a nephropathologist and observers blinded to the treatment modality. Periodic acid-SchiffPeriodic acid-Schiff–stained sections were examined for histological changes due to acute tubular injury and ATN and were quantitated by counting the percent of tubules that displayed loss of brush border, cast formation, tubular simplification, and tubule dilatation as previously described.15,17,26-28 Slides were scanned using the Aperio scanner. Each high power field (400×) was divided into 6 quadrants. For every parameter at least 150 to 200 tubules were examined per mouse section in both cortex and medulla.

Back to Top | Article Outline

BBI Score

The percent of tubules that displayed the loss of brush border was counted as previously described.15,18,26-28 Severity of BBI was assessed in each tubule as follows: a score of 1 indicates less than 25% of the circumference of the analyzed tubule had BBI, a score of 2 indicates 26% to 50% of the tubule had BBI, a score of 3 indicates 51% to 75% of the tubule had BBI, and a score of 4 indicates more than 76% of the circumference of the tubule had BBI. The total score was calculated as mean BBI score per number of tubules in each quadrant.

Back to Top | Article Outline

Cast formation, Tubular Simplification, and Tubular Dilation

Casts were counted in each tubule in every quadrant as previously described.15,17,26-28 Similarly each tubule demonstrating tubular injury (ie, the presence of flattened tubular cells with nuclei and minimal cytoplasm) was counted. To measure the tubule dilation, image processing and analysis with Aperio ImageScope (Leica Biosystems) were used to manually demarcate the borders of the whole tubule and the lumen. The software then quantitated the tubular and luminal areas. Ten tubules per sample were demarcated and used to quantify the data.

Back to Top | Article Outline

Morphological Quantification of Apoptosis Cells

Morphologic criteria were used to count apoptotic tubular epithelial cells on PAS staining as previously described.15,18,26,27 These characteristics included cellular rounding and shrinkage, nuclear chromatin compaction and formation of apoptotic bodies. Apoptotic tubular epithelial cells were quantitatively assessed per high-power field (400×) in a blinded fashion.

Back to Top | Article Outline

Apoptotic Cell Detection via TUNEL Staining

Apoptotic cells were also detected using the DeadEnd Colorimetric TUNEL assay kit (Promega, G7130), which measures nuclear DNA fragmentation, an important biochemical hallmark of apoptosis. All steps were followed according to the manufacturer's directions. The brown-colored TUNEL-positive cells were quantified in 12 randomly selected quadrant fields at 400× magnifications by an observer blinded to the treatment modality.

Back to Top | Article Outline

Statistics

All in vitro data presented were confirmed in at least 4 independent experiments. All mouse studies included 4 in the control group and at least 7 (7-11) animals per experimental group. All values are expressed as means ± SEM. Data were analyzed by analysis of variance (ANOVA) followed by Newman-Keuls Comparison Test using GraphPad prism software version 5.01 (San Diego, CA). A P value of less than 0.05 was considered significant.

Back to Top | Article Outline

RESULTS

Q-VD-OPh Significantly Reduces Caspase-3 Protein Expression and Activity In Vitro

Mouse RTECS were subjected to an in vitro model of CS/REW as previously described.16-18 Mouse RTECs cold stored in saline with or without vehicle (DMSO) demonstrated increased cleaved capsase-3 protein expression compared with cells stored with Q-VD-OPh. The effect of Q-VD-OPh on caspase-3 protein expression was dose-dependent, and the greatest inhibition of protein expression occurred at a concentration of 50 μM (Figure 1). We then determined whether Q-VD-OPh at a concentration of 50 μM prevented apoptosis during CS/REW. Cell death percentage, measured as annexin V-positive and/or PI-positive cells by flow cytometry was significantly reduced by Q-VD-OPh compared with saline or vehicle (DMSO) (Figure 2). Q-VD-OPh (50 μM)–treated cells also had significantly reduced cleaved-caspase-3 activity (Figure 3A), but no difference in caspase-1 activity (Figure 3B) compared with saline and vehicle (DMSO)-treated cells.

FIGURE 2

FIGURE 2

FIGURE 3

FIGURE 3

Back to Top | Article Outline

The Effect of Q-VD-OPh on Renal Function in a Syngeneic Mouse Model of Kidney Transplantation

We used a mouse model of kidney transplantation to determine whether the reduction in tubular cell death observed with Q-VD-OPh in vitro would translate into improved renal function and histology in vivo after kidney transplantation. In the current study we subjected mouse donor kidneys to CS of 60 minutes in saline, vehicle (DMSO), or Q-VD-OPh followed by syngeneic kidney transplantation. Nontransplanted, wild-type donors were used as controls. Cold ischemia times (60 ± 0.12 minutes for all transplanted groups) and warm ischemia times (34.5 ± 2.6 minutes for saline; 34.4 ± 2.8 minutes for DMSO; 35.1 ± 2.9 minutes for Q-VD-OPh) were recorded to ensure consistency of the mouse kidney transplant procedure and did not vary between the groups (Figure S1, SDC,http://links.lww.com/TP/B558). Transplantation of donor kidneys that had been cold stored in saline or DMSO resulted in significantly increased recipient serum creatinine (sCr in mg/dL) compared with controls (saline sCr, 2.71 ± 0.21 vs control sCr, 0.33 ± 0.02; P < 0.001) (DMSO sCr, 1.85 ± 0.29; P < 0.05 vs control sCr). In contrast, sCr was not significantly different in donor kidneys cold stored with Q-VD-OPh compared with controls (Q-VD-OPh sCr, 0.99 ± 0.41 vs control sCr, 0.33 ± 0.02; P = NS). Transplantation of donor kidneys cold stored for 60 minutes at 4 °C in saline resulted in significantly increased recipient sCr versus recipients of donor kidneys cold-stored in DMSO or Q-VD-OPh (saline sCr, 2.71 ± 0.21 vs DMSO sCr, 1.85 ± 0.29, *P < 0.05) (saline sCr, 2.71 ± 0.21 vs Q-VD-OPh sCr, 0.99 ± 0.41, *P < 0.001) (Figure 4).

FIGURE 4

FIGURE 4

Back to Top | Article Outline

The Effect of Q-VD-OPh on Histology in a Syngeneic Mouse Model of Kidney Transplantation

Widefield microscopic assessment (400×) of sham kidneys demonstrated normal renal architecture with well-preserved tubular brush border, normal columnar epithelium, and an absence of casts or tubular dilation. Transplantation of kidneys cold stored in saline or DMSO resulted in widespread flattened tubular cells, loss of brush border, cast formation, and distended tubular lumens in both cortex and medulla. In contrast, transplantation of kidneys cold stored in Q-VD-OPh resulted in partial preservation of renal architecture, with demonstrable brush border, fewer casts, and retention of the normal cuboidal appearance of tubular epithelial cells (Figure 5). Morphologic assessment of apoptosis in both cortex and medulla in Q-VD-OPh–treated kidney transplants showed significantly decreased apoptotic tubular epithelial cells compared with DMSO- or saline-treated kidney transplants (Figure 6). Morphological assessment of apoptosis was confirmed with TUNEL staining which also demonstrated significantly decreased apoptotic tubular epithelial cells in Q-VD-OPh treated kidney transplants compared with DMSO- or saline-treated kidney transplants (Figure 7).

FIGURE 5

FIGURE 5

FIGURE 6

FIGURE 6

FIGURE 7

FIGURE 7

The number of tubules demonstrating BBI in both cortex and medulla was significantly reduced in Q-VD-OPh kidney transplants versus DMSO- or saline-treated kidney transplants (Figure 8B), as was the BBI severity (Figure 8C). Kidneys cold stored in saline demonstrated significantly increased tubular injury (Figure 9) compared with DMSO- and Q-VD-OPh–treated kidneys. The number of casts observed within tubular lumens was significantly increased after transplantation in DMSO- and saline-treated kidneys versus Q-VD-OPh–treated kidneys (Figure 10). Using software analysis of lumen tubule size with the Aperio ImageScope imaging system, we observed significantly increased tubule lumen area in saline- and DMSO-treated kidneys compared with Q-VD-OPh–treated kidneys (Figure 11B). To determine whether the increased tubule lumen area observed in the saline- and DMSO-treated groups was due to dilation of the tubule itself or loss of brush border and tubular cell volume, we also measured the total tubule cross-sectional in over 290 tubules. No difference was observed in tubule cross-sectional area in saline-, DMSO-, and Q-VD-OPh–treated kidneys after transplantation, and no difference was observed when the 3 groups were compared with nontransplanted control kidneys (Figure 11C), indicating that the increased tubule lumen area in kidneys treated with saline and DMSO was due to the loss of brush border and tubule cell volume and not due to dilation of the tubule itself.

FIGURE 8

FIGURE 8

FIGURE 9

FIGURE 9

FIGURE 10

FIGURE 10

FIGURE 11

FIGURE 11

Back to Top | Article Outline

DISCUSSION

Delayed graft function is an extremely important problem in the field of kidney transplantation.8,29-32 Despite the importance of DGF, little advancement in understanding its pathogenesis has occurred in the last 50 years. A previous editorial33 highlighted not only the importance of DGF but also “the limitations of the state of the art” regarding techniques of organ preservation and the pathogenesis of DGF, and “the stagnation that this area has undergone.”

Delayed graft function is associated with a variety of potential causes in both the donor and the recipient, including brain death,34 circulatory death,35,36 allogenicity,9 donor age, donor weight, prolonged cold/warm ischemia time,3,31,37 recipient dialysis vintage,38 and recipient diabetes mellitus.39 The relative contribution of each of the aforementioned variables to the development of DGF is difficult to determine clinically. It is also difficult, in the clinical setting, to isolate each variable and demonstrate causality or the mechanism by which they may lead to DGF.

In our model, we have endeavored to isolate CI as a variable to determine its effect on graft histology and function after transplant. We have previously demonstrated that perfusion of donor kidneys with saline in the absence of CI results in minimal histological injury to the graft and good posttransplant renal function.19,20 In the current model, CS of donor kidneys with saline at 4°C resulted in significantly increased sCr, BBI, and apoptosis, indicating that CI itself can result in histological injury and renal dysfunction.

The observation that Q-VD-OPh significantly reduced apoptosis, BBI, cast formation, and improved renal function implicates caspases in the pathogenesis of injury after CI. The improvement in tubular cell apoptosis observed with Q-VD-OPh was likely due to a decrease in proapoptotic caspase-3 protein expression and activity as opposed to a decrease in inflammation, because Q-VD-OPh had no effect on proinflammatory caspase-1. In contrast, tubular injury was improved in both DMSO- and Q-VD-OPh–treated kidneys. Q-VD-OPh is reconstituted in DMSO (vehicle), which introduces a possible confounder because DMSO itself has been shown to have anti-inflammatory effects.21,22 However, treatment of cultured renal tubular epithelial cells with DMSO alone during CS/REW did not have any effect on caspase-3 protein expression or caspase-3 activity, suggesting that Q-VD-OPh was responsible for the inhibition of caspase-3.

There are several limitations to our study. Our model does not fully represent clinical DGF, because it does not involve renal replacement therapy. Indeed, renal support is provided by 1 remaining native kidney in the first postoperative week, after which a native nephrectomy is performed, and the mouse is left with only the kidney transplant. At the present time, renal replacement therapy of mice for an extended period is not possible.

Another departure from clinical DGF in our model is the CIT of 1 hour, which is shorter than CITs typically associated with DGF in humans. Our attempts to use CITs used in human transplantation (ranging from 4 to 48 hours) led to unacceptably high animal loss and produced the equivalent of primary nonfunction. We determined that 1-hour CIT was the maximum tolerable period that would allow graft survival and would also cause significant tubular injury, necrosis, and renal dysfunction, features that are typical of human DGF kidneys.40,41 We examined approximately 4700 tubules and observed widespread tubular injury affecting 60% to 70% of tubules, accompanied by widespread BBI, apoptosis, and cast formation in murine grafts cold stored in saline for 1 hour. Therefore, 1-hour CIT in our model induced diffuse injury and renal dysfunction that resembled the pathology of biopsies from human kidney grafts with DGF.40,41 Other investigators have also demonstrated renal dysfunction and tubular necrosis in approximately 60% of tubules of grafts cold stored in saline for 60 minutes before mouse kidney transplant.42 It is clear therefore, that mouse kidneys are more susceptible to injury than human kidneys, requiring shorter CIT to produce the same injury observed in clinical DGF. A similar parallel is observed in clamp models of warm ischemia-reperfusion injury, in which a large body of literature demonstrates that brief periods of warm ischemia (~18-30 minutes) produce significant ATN and high SCr.27,43-49

Another limitation of our study was that we did not use a preservation solution such as University of Wisconsin solution. Our goal was to isolate CI as a variable and determine whether CI-induced injury could be ameliorated by caspase inhibition. We therefore excluded other potentially injurious variables such as circulatory death and allogenicity. We also necessarily excluded potentially beneficial interventions, such as machine perfusion or the use of preservation solutions to prevent any confounding of a protective effect by Q-VD-OPh. Our study provides proof-of-principle evidence that CI alone results in significant tubular cell apoptosis, injury, and renal dysfunction, which can be ameliorated by caspase inhibition in a mouse model. Further studies including other variables, such as circulatory death, preservation solutions, and machine perfusion, as well as the use of large animal models will further determine the clinical applicability of this approach. In this regard, we have previously demonstrated substantial renal tubular epithelial cell apoptosis that occurs in a porcine DCD model.26

Caspase inhibition reduced but did not completely prevent histological injury and renal dysfunction, suggesting that other injurious pathways may still have been active. It is important to note that Q-VD-OPh does not inhibit the proinflammatory capsase-1. Several murine studies implicate caspase-1 in warm ischemia-reperfusion injury, and inhibition of capsase-1 reduces tubular injury and AKI.48-50 Recent studies have also highlighted the potential importance of programed necrosis and the RIP-kinase pathway in warm ischemia-reperfusion injury.51,52 It is likely that several pathways are involved in the pathogenesis of AKI after transplantation, and it may be necessary to combine a cocktail of agents that inhibit apoptosis and programed necrosis to completely prevent DGF.

A strength of our study is that it provides proof-of-principle evidence that pretreatment of a donor kidney during cold preservation will reduce subsequent renal dysfunction and histological injury posttransplant. We focused on the treatment of the donor kidney because it would be the easiest therapy to translate to human kidney transplantation. Treating the donors before organ retrieval or the recipient after transplantation may provide added benefit. Treatment of both donors and recipients is feasible clinically, but is complicated by issues of obtaining consent and the possible systemic side effects of a given therapy. Treatment of donor kidneys before transplantation is an attractive potential solution to the problem of CI-induced injury since the majority of deceased donor kidneys require cold preservation during transport. In addition, the risk of systemic side effects is minimized.

In conclusion, we have demonstrated that treatment of a donor kidney with Q-VD-OPh prevents histological injury and improves renal function in a mouse model of kidney transplantation. Inhibition of caspases before transplantation may be an effective therapy for CI-induced injury in DGF.

Back to Top | Article Outline

REFERENCES

1. Opelz G, Dohler B. Multicenter analysis of kidney preservation. Transplantation. 2007;83:247–253.
2. Hameed AM, Pleass HC, Wong G, et al. Maximizing kidneys for transplantation using machine perfusion: from the past to the future: a comprehensive systematic review and meta-analysis. Medicine (Baltimore). 2016;95:e5083.
3. Irish WD, Ilsley JN, Schnitzler MA, et al. A risk prediction model for delayed graft function in the current era of deceased donor renal transplantation. Am J Transplant. 2010;10:2279–2286.
4. Treat E, Chow EKH, Peipert JD, et al. Shipping living donor kidneys and transplant recipient outcomes. Am J Transplant. 2018;18:632–641.
5. Matos ACC, Requiao Moura LR, Borrelli M, et al. Impact of machine perfusion after long static cold storage on delayed graft function incidence and duration and time to hospital discharge. Clin Transplant. 2018;32.
6. Kyllonen LE, Salmela KT, Eklund BH, et al. Long-term results of 1047 cadaveric kidney transplantations with special emphasis on initial graft function and rejection. Transpl Int. 2000;13:122–128.
7. Hetzel GR, Klein B, Brause M, et al. Risk factors for delayed graft function after renal transplantation and their significance for long-term clinical outcome. Transpl Int. 2002;15:10–16.
8. Ojo AO, Wolfe RA, Held PJ, et al. Delayed graft function: risk factors and implications for renal allograft survival. Transplantation. 1997;63:968–974.
9. Shoskes DA, Cecka JM. Deleterious effects of delayed graft function in cadaveric renal transplant recipients independent of acute rejection. Transplantation. 1998;66:1697–1701.
10. Halloran PF, Hunsicker LG. Delayed graft function: state of the art, November 10–11, 2000. Summit meeting, Scottsdale, Arizona, USA. Am J Transplant. 2001;1:115–120.
11. Lee CM, Carter JT, Alfrey EJ, et al. Prolonged cold ischemia time obviates the benefits of 0 HLA mismatches in renal transplantation. Arch Surg. 2000;135:1016–1019.
12. Liu X, Zou H, Slaughter C, et al. DFF, a heterodimeric protein that functions downstream of caspase-3 to trigger DNA fragmentation during apoptosis. Cell. 1997;89:175–184.
13. Castaneda MP, Swiatecka-Urban A, Mitsnefes MM, et al. Activation of mitochondrial apoptotic pathways in human renal allografts after ischemiareperfusion injury. Transplantation. 2003;76:50–54.
14. Oberbauer R, Rohrmoser M, Regele H, et al. Apoptosis of tubular epithelial cells in donor kidney biopsies predicts early renal allograft function. J Am Soc Nephrol. 1999;10:2006–2013.
15. Jani A, Ljubanovic D, Faubel S, et al. Caspase inhibition prevents the increase in caspase-3, -2, -8 and -9 activity and apoptosis in the cold ischemic mouse kidney. Am J Transplant. 2004;4:1246–1254.
16. Jain S, Keys D, Martin S, et al. Protection from apoptotic cell death during cold storage followed by rewarming in 13-lined ground squirrel tubular cells: the role of prosurvival factors X-linked inhibitor of apoptosis and phosphoAkt. Transplantation. 2016;100:538–545.
17. Jain S, Keys D, Ljubanovic D, et al. Protection against cold storage-induced renal tubular cell apoptosis. Transplantation. 2015;99:2311–2316.
18. Jain S, Keys D, Nydam T, et al. Inhibition of autophagy increases apoptosis during re-warming after cold storage in renal tubular epithelial cells. Transpl Int. 2015;28:214–223.
19. Plenter RJ, Jain S, Nydam TL, et al. Revised arterial anastomosis for improving murine kidney transplant outcomes. J Invest Surg. 2015;28:208–214.
20. Plenter R, Jain S, Ruller CM, et al. Murine kidney transplant technique. J Vis Exp. 2015:e52848.
21. Smith G, Bertone AL, Kaeding C, et al. Anti-inflammatory effects of topically applied dimethyl sulfoxide gel on endotoxin-induced synovitis in horses. Am J Vet Res. 1998;59:1149–1152.
22. Repine JE, Eaton JW, Anders MW, et al. Generation of hydroxyl radical by enzymes, chemicals, and human phagocytes in vitro. Detection with the anti-inflammatory agent, dimethyl sulfoxide. J Clin Invest. 1979;64:1642–1651.
23. Han WR, Murray-Segal LJ, Mottram PL. Modified technique for kidney transplantation in mice. Microsurgery. 1999;19:272–274.
24. Ge F, Gong W. Strategies for successfully establishing a kidney transplant in a mouse model. Exp Clin Transplant. 2011;9:287–294.
25. Tse GH, Hughes J, Marson LP. Systematic review of mouse kidney transplantation. Transpl Int. 2013;26:1149–1160.
26. Jani A, Zimmerman M, Martin J, et al. Perfusion storage reduces apoptosis in a porcine kidney model of donation after cardiac death. Transplantation. 2011;91:169–175.
27. Akcay A, Nguyen Q, He Z, et al. IL-33 Exacerbates Acute Kidney Injury. J Am Soc Nephrol. 2011;22:2057–67.
28. Jani A, Epperson E, Martin J, et al. Renal protection from prolonged cold ischemia and warm reperfusion in hibernating squirrels. Transplantation. 2011;92:1215–1221.
29. Perico N, Cattaneo D, Sayegh MH, et al. Delayed graft function in kidney transplantation. Lancet. 2004;364:1814–1827.
30. Hariharan S, McBride MA, Cherikh WS, et al. Post-transplant renal function in the first year predicts long-term kidney transplant survival. Kidney Int. 2002;62:311–318.
31. Siedlecki A, Irish W, Brennan DC. Delayed graft function in the kidney transplant. Am J Transplant. 2011;11:2279–2296.
32. Yarlagadda SG, Coca SG, Formica RN Jr, et al. Association between delayed graft function and allograft and patient survival: a systematic review and meta-analysis. Nephrol Dial Transplant. 2009;24:1039–1047.
33. Shapiro R, Halloran PF. Organ preservation—can we do it better? Am J Transplant. 2008;8:479–480.
34. Powner DJ, Hendrich A, Nyhuis A, et al. Changes in serum catecholamine levels in patients who are brain dead. J Heart Lung Transplant. 1992;11:1046–1053.
35. Rojas-Pena A, Reoma JL, Krause E, et al. Extracorporeal support: improves donor renal graft function after cardiac death. Am J Transplant. 2010;10:1365–1374.
36. Summers DM, Johnson RJ, Allen J, et al. Analysis of factors that affect outcome after transplantation of kidneys donated after cardiac death in the UK: a cohort study. Lancet. 2010;376:1303–1311.
37. McLaren AJ, Jassem W, Gray DW, et al. Delayed graft function: risk factors and the relative effects of early function and acute rejection on long-term survival in cadaveric renal transplantation. Clin Transplant. 1999;13:266–272.
38. Doshi MD, Garg N, Reese PP, et al. Recipient risk factors associated with delayed graft function: a paired kidney analysis. Transplantation. 2011;91:666–671.
39. Parekh J, Bostrom A, Feng S. Diabetes mellitus: a risk factor for delayed graft function after deceased donor kidney transplantation. Am J Transplant. 2010;10:298–303.
40. Racusen LC, Fivush BA, Li YL, et al. Dissociation of tubular cell detachment and tubular cell death in clinical and experimental “acute tubular necrosis”. Lab Invest. 1991;64:546–556.
41. Solez K, Racusen LC, Marcussen N, et al. Morphology of ischemic acute renal failure, normal function, and cyclosporine toxicity in cyclosporine-treated renal allograft recipients. Kidney Int. 1993;43:1058–1067.
42. Wang L, Wei J, Jiang S, et al. The effects of different storage solutions on renal ischemia tolerance after kidney transplantation in mice. Am J Physiol Renal Physiol. 2017;314:F381–F387.
43. Vervaet BA, Moonen L, Godderis L, et al. Untargeted DNA-demethylation therapy neither prevents nor attenuates ischemia-reperfusion-induced renal fibrosis. Nephron. 2017;137:124–136.
44. Bongoni AK, Lu B, Salvaris EJ, et al. Overexpression of human CD55 and CD59 or treatment with human CD55 protects against renal ischemia-reperfusion injury in mice. J Immunol. 2017;198:4837–4845.
45. Ryan J, Kanellis J, Blease K, et al. Spleen tyrosine kinase signaling promotes myeloid cell recruitment and kidney damage after renal ischemia/reperfusion injury. Am J Pathol. 2016;186:2032–2042.
46. Yuan D, Collage RD, Huang H, et al. Blue light reduces organ injury from ischemia and reperfusion. Proc Natl Acad Sci U S A. 2016;113:5239–5244.
47. Ranganathan P, Jayakumar C, Tang Y, et al. MicroRNA-150 deletion in mice protects kidney from myocardial infarction-induced acute kidney injury. Am J Physiol Renal Physiol. 2015;309:F551–F558.
48. He Z, Lu L, Altmann C, et al. Interleukin-18 binding protein transgenic mice are protected against ischemic acute kidney injury. Am J Physiol Renal Physiol. 2008;295:F1414–F1421.
49. Melnikov VY, Faubel S, Siegmund B, et al. Neutrophil-independent mechanisms of caspase-1- and IL-18-mediated ischemic acute tubular necrosis in mice. J Clin Invest. 2002;110:1083–1091.
50. Kim HJ, Lee DW, Ravichandran K, et al. NLRP3 inflammasome knockout mice are protected against ischemic but not cisplatin-induced acute kidney injury. J Pharmacol Exp Ther. 2013;346:465–472.
51. Linkermann A, Brasen JH, Himmerkus N, et al. Rip1 (receptor-interacting protein kinase 1) mediates necroptosis and contributes to renal ischemia/reperfusion injury. Kidney Int. 2012;81:751–761.
52. Gunther C, Neumann H, Neurath MF, et al. Apoptosis, necrosis and necroptosis: cell death regulation in the intestinal epithelium. Gut. 2013;62:1062–1071.

Supplemental Digital Content

Back to Top | Article Outline
Copyright © 2018 Wolters Kluwer Health, Inc. All rights reserved.