Objectives: Cytokine response modifier A protein is a caspase inhibitor that inhibits caspase activity and protects cells from apoptosis. Chronic cyclosporine nephropathy is a significant cause of chronic graft dysfunction. We explored cytokine response modifier A protein-alleviated chronic cyclosporine nephropathy for ways of improving chronic graft dysfunction.

Materials and Methods: Cytokine response modifier A protein-transferring HK-2 cells were cultured with different concentrations of cyclosporine. Cytokine response modifier A protein mRNA and proteins were detected by real-time polymerase chain reaction and Western blot, cell viability was detected by (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), and apoptosis was detected by flow cytometry.

Results: Cyclosporine caused a concentration-dependent and time-dependent loss of cell viability in HK-2 cells. Cytokine response modifier A protein mRNA was expressed at 48 and 72 hours (P < .05), while protein was detected at 72 hours. Cell viability in the cytokine response modifier A protein-transfected group was significantly greater than that of the control group when treated with 1 µg/mL, 10 µg/mL, or 20 µg/mL cyclosporine at 24 or 48 hours (P < .05). The apoptosis in cytokine response modifier A protein-transfected cells was significantly lower than that of controls (P < .05).

Conclusions: Cytokine response modifier A protein protects renal cells from cyclosporine injury by inhibiting activated caspases. Cytokine response modifier A protein transfection may improve chronic cyclosporine nephropathy and provide for improving chronic graft dysfunction.

Key words : CrmA gene, Cyclosporine, Chronic graft dysfunction, HK-2 cell lines, Chronic cyclosporine nephropathy

Introduction

Chronic cyclosporine nephropathy is an important cause of chronic graft dysfunction. Long-term use of cyclosporine can cause chronic cyclosporine nephropathy by directly injuring renal cells.^1-2 Cyclosporine could dramatically induce their apoptosis in time-dependent and dose-dependent manner when cocultured with different renal cells in vitro.^3-5 When cyclosporine was given to Wistar rats and imprinting control region mice for 4 weeks, it induced chronic cyclosporine nephropathy that increased tubular and interstitial cell apoptosis in cyclosporine-treated animals.^6-9

There were at least 6 apoptotic pathways that mediated renal cell apoptosis in vitro and in vivo. Cyclosporine induced renal cell apoptosis through Fas/FasL,^10-11 mitochondrial,^{2, 12} and the endoplasmic reticulum pathways.¹³ Cyclosporine also induced cell apoptosis through angiotensin 2,^14-15 nitric oxide,¹⁵ and the hypertonicity-related apoptosis pathways.¹⁶ Activated caspases might be the final common pathway of all these pathways in vivo and in vitro. There were 21 interventions that could inhibit renal cell apoptosis induced by cyclosporine, such as antioxidant drugs,^17-19 recombinant human erythropoietin,²⁰ hepatocyte growth factor,²¹ spironolactone,²² rosiglitazone,¹⁴ and colchicine.²³ However, these interventions have not been clinically proven.

In 1986, Pickup and associates found that cytokine response modifier A protein has a 38 Kda protein virulence factor, which leads to hemorrhagic damage in the Brighton Red strain of cowpox viral infection.^24-25 Cytokine response modifier A (CrmA) protein was a caspase inhibitor that broadly inhibits the activity of many caspases, such as caspase-3, caspase-4, caspase-5, caspase-8, caspase-9, and caspase-10, and then protects target cells from apoptosis induced by many factors such as immune injury, anoxia, and chemical factors.^26-28

We hypothesized that CrmA-protected cyclosporine induced cell apoptosis and offer ideas for protecting graft kidney from injury by cyclosporine and improving chronic graft dysfunction.²⁹ In this article, the CrmA gene was transfected into HK-2 cells, and the CrmA protein inhibited activated caspases (their common pathway) and protected HK-2 cells from cyclosporine-induced apoptosis.

Materials and Methods

All protocols and experimental studies were approved by the ethics committee of the institution before the study began, and the protocol conforms to the ethical guidelines of the 1975 Helsinki Declaration.

Reagents
The human proximal tubular cell line HK-2 cells were obtained from American Type Culture Collection (American Type Culture Collection [ATCC], Manassas, VA USA), grown in Dulbecco Modified Eagle medium (Invitrogen Corporation, Carlsbad, CA, USA) supplemented with 10% fetal bovine serum (Invitrogen) and HEPES free acid 15 mM, at 37°C, 95% air, and 5% CO₂. p-CAGGS-CrmA (LMBP3830) and p-CAGGS (LMBP-2453) were purchased from BCCM/LMBP plasmid collection MOSAICC (Micro-Organisms Sustainable use and Access regulation International Code of Conduct, Belgium). Lipofectamine 2000 was purchased from Invitrogen. Cowpox virus CrmA-Ab was purchased from Santa Cruz Biotechnology, Inc (Santa Cruz, CA, USA) Cytokine response modifier A-Ab was purchased from Santa Cruz Biotechnology, Inc. A Catrimo-14TM RNA Isolation Kit, a PrimeScript RT Reagent Kit (Perfect Real Time), and a SYBR Premix Ex Taq II were from Takara Biotechnology ([Dalian], Shiga, Japan). An Annexin V-FITC apoptosis detection kit was purchased from Calbiochem (San Diego, CA, USA). All other chemicals were of analytic reagent grade.

Cell culture
Human killer (HK)-2 cells were grown in 25-mm Falcon T flasks using Dulbecco's Modified Eagle's Medium (DMEM) supplemented with 10% fetal calf serum. Transfection of the CrmA gene into HK-2 cells was achieved using Lipofectamine 2000 transfection agent; after its protocol, cell colonies were picked up and analyzed for expression of the CrmA protein and mRNA. Human killer-2 cells were harvested 72 hours after transfection and cultured with cyclosporine. Finally, their viability and apoptosis were detected by MTT (3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) and Annexin V/propidium iodide (PI) staining assay.

Real-time polymerase chain reaction
We quantified mRNA levels of CrmA in HK-2 cells at 24, 48, and 72 hours after transfection (1 × 10⁵) using a real-time fluorescence detection. RNAs were extracted from a single-cell suspension using the Catrimox-14 MIRNA Isolation Kit (Takara, Tokyo) according to the manufacturer’s instructions. Real-time quantitative polymerase chain reaction experiments were performed with an multicolor real-time polymerase chain reaction detection system (Applied Biosystems, Foster City, CA, USA), using a SYBR Premix Ex Taq II according to the manufacturer’s protocol. The primer sequences were as follows: for β-actin (sense, 5'-ATGGAGCCACCGATCCACA-3', and antisense, 5'-CATCCGTAAAGACCTCTATGCCAAC-3') and for CrmA (sense, 5'-TTCTCCACCGTCAATCTCGTC-3', and antisense 5'-CCTTATTCTTGTCCGCCTCCT-3'). Each sample was normalized based on its β-actin content. Thermal cycling conditions were as follows: 10 seconds at 95°C, followed by 40 cycles of 95°C for 5 seconds, and 60°C for 30 seconds.

Western blot analysis
At 72 hours after transfection, HK-2 cells were harvested and washed with cold phosphate buffered saline and solubilized with lysis buffer (50 mM Tris-HCl, 150 mM NaCl, 5 mM ethylenediaminetetraacetic acid [EDTA], 1 mM phenylmethanesulfonyl fluoride [PMSF], 1% Triton X-100, 1% sodium deoxycholate, 0.1% sodium dodecyl sulfate [SDS]) at 4°C for 1 hour. Lysates were centrifuged at 14 000 revolutions per minute for 10 minutes, and the supernatant was collected as sample. Samples were separated on 14% SDS-polyacrylamide gel, and transferred to nitrocellulose membranes using standard electroblotting procedures. Membranes were blocked with 5% skim milk in Tris-Cl buffered saline (TBS-T, 0.1% Tween-20). Blots were incubated with primary antibodies (1:800) of CrmA at 4°C overnight. Immunoblots were washed, and then they were incubated with horseradish peroxidase conjugated secondary antibody (ZSbio, China) at room temperature for 1 hour and subsequently processed for enhanced chemiluminescence detection using BCIP/NBT alkaline phosphatase substrate solution protocol (Roche Pharmaceuticals, Basel, Switzerland). Signals were detected by a chemiluminescence detection system (Bio-Rad Laboratories, Clinical Diagnostics Group, Hercules, CA, USA).

MTT assay
Four hours before the end of the incubation, 20 µL of 5 mg/mL MTT was added to each well. MTT was removed and replaced with 100 µL of dimethyl sulfoxide (DMSO) for further incubation for 10 minutes at 37°C until the crystals were dissolved. The optical density value of each well was measured using a KCjunior Microplate Spectrophotometer (BioTek US, Winooski, VT, USA) with a test wavelength of 570 nm.

Human killer-2 cells were harvested, and the cells were divided into 24-hour and 48-hour groups. Concentrations of cyclosporine were 0.01 and 1000 µg/mL. Each group had 5 repeated wells. Each well of a 96-well plate was seeded with 0.5 × 104 cells/200 µL. Cells were incubated at 37°C under an atmosphere of 5% CO₂ for 24 or 48 hours. Cells viability was detected by MTT.

Human killer-2 cells were harvested 72 hours after transfection. Cells were divided into the following 3 groups: the p-CAGGS-CrmA group, the p-CAGGS group, and the nontransfected group. Each group had 5 repeated wells. Each well of a 96-well plate was seeded with 0.5 × 10⁴ cells/200 µL. Concentrations of cyclosporine were 1 µg/mL, 10 µg/mL, and 20 µg/mL. Cells were incubated at 37°C under an atmosphere of 5% CO2 for 24 or 48 hours. Cells viability was detected by MTT.

Annexin V/propidium iodide staining assay
HK-2 cells were harvested 72 hours after transfection. Cells were divided into the following 3 groups: the p-CAGGS-CrmA transfected group, the p-CAGGS transfected group, and the nontransfected group. Each group had 5 repeated wells. Each well of a 6-well plate was seeded with 1 × 10⁵ cells/500 µL. Concentrations of cyclosporine were 1 µg/mL, 10 µg/mL, or 20 µg/mL. Apoptosis ratio was measured by BD FACSCalibur multicolor flow cytometer (Becton Dickinson, Sparks, MD, USA) according to the instruction provided by the annexin V/propidium iodide kit (BD Biosciences, San Jose, California, USA). Briefly, after treatment with cyclosporine 1 µg/mL, 10 µg/mL, and 20 µg/mL for 24 hours, HK-2 cells were harvested and washed twice with precold phosphate buffered saline and resuspended in a binding buffer containing FITC-conjugated annexin V antibody (1.25 µL) and propidium iodide, 10 µL. After incubation for 30 minutes in the dark, cells were analyzed by flow cytometry.

Statistical Analyses
Statistical analyses were performed with SPSS software for Windows (Statistical Product and Service Solutions, version 13.0, SSPS Inc, Chicago, IL, USA). Differences in the means between each group were detected by the t test. Values for P < .05 were considered statistically significant.

Results

Quantitative analysis of cytokine response modifier A protein gene expression
Real-time quantitative polymerase chain reaction was used to quantify CrmA gene expression in the p-CAGGS-CrmA transfected HK-2 cells, p-CAGGS transfected HK-2 cells, and nontransfected HK-2 cells. Amplification specificity for CrmA and β-actin was determined by analyzing the dissociation curves. Cytokine response modifier A protein expression was detected in CrmA transfected HK-2 cells (P < .05), not in p-CAGGS transfected or nontransfected HK-2 cells (P < .05). The highest level of CrmA expression was detected in CrmA transfected HK-2 cells at 72 hours (P < .05) (Figure 1).

Expression of cytokine response modifier A protein in human killer-2 cell lines
To test expression of CrmA protein in transfected HK-2 cells, we detected cell transfection after 72 hours. In Figure 2, Western blot analysis demonstrated that CrmA expression (34-48 Kd) was seen in CrmA transfected HK-2 cell lines. Cytokine response modifier A protein expression was not found in the p-CAGGS transfected group or nontransfected group.

Drug toxicity of cyclosporine to human killer-2 cell
Human killer-2 cells were cocultured with different doses of cyclosporine. In Figure 3, cyclosporine directly damaged HK-2 cells; its drug toxicity also was increased with dosage and time, and the corresponding cell activity was decreased. Cell viability of HK-2 cocultured with cyclosporine (0.01-10 µg/mL) at 48 hours was higher than it was at 24 hours. When HK-2 cells were cocultured with cyclosporine (> 200 µg/mL) at 24 or 48 hours, their activity was completely lost.

Protection of human killer-2 cell viability by cytokine response modifier A protein
At 72 hours after transfection, each cell group (the CrmA-transfected group, the p-CAGGS transfected, and the nontransfected group) was treated with 1 µg/mL, 10 µg/mL, or 20 µg/ml of cyclosporine for 24 or 48 hours. Then the MTT assay was done to quantify protection of HK-2 cells by CrmA. Optical density value was used to represent cell viability. As shown in Figure 4, the cell viability in the p-CAGGS-CrmA transfected group was significantly higher than that of the other groups treated with 1 µg/mL, 10 µg/mL, or 20 µg/mL of cyclosporine at 24 or 48 hours (P < .05). Cyclosporine could significantly decrease HK-2 cell viability in a concentration-dependent manner in controls (p-CAGGS transfected and nontransfected group) (P < .05). There was no significant difference in cell viability in either the p-CAGGS transfected group or the nontransfected group (P > .05).

Protection of HK-2 cell apoptosis by cytokine response modifier A protein
Detection of apoptosis of HK-2 treated with cyclosporine is usually characterized by the phosphatidylserine exposure at the outer leaflet of the plasma membrane. Based on their affinity for annexin V, apoptotic cells can be distinguished from annexin V-negative living cells by cytometric procedures.^30-31 Furthermore, the double labeling assay (annexin V combined with propidium iodide) allows further distinction of necrotic or late apoptotic (annexin V+/PI+) and early apoptotic cells (annexin V+/PI-).

As shown in Figure 5, flow counting cells were 18 605.7 ± 2461.11. Cyclosporine could induce apoptosis in cells of varying degrees (including early and late apoptosis). The apoptosis rate in CrmA transfected cells was significantly lower than it was in controls (the p-CAGGS transfected and the nontransfected group) (P < .05). It induced HK-2 cells apoptosis in a concentration-dependent manner in the control groups (P < .05). The rate of apoptosis showed no significant difference in the p-CAGGS transfected group or the nontransfected group (P > .05).

Discussion

Cyclosporine is widely used in organ transplant and autoimmune diseases,³² but it induces kidney cell apoptosis though many apoptotic pathways, causes chronic cyclosporine nephrotoxicity, and has become an important cause of chronic graft dysfunction.^33-38 We hypothesized that CrmA might protect cyclosporine-induced cells from dying and provide new ideas for protecting graft kidney from injury by cyclosporine and improving chronic graft dysfunction. Cytokine response modifier-A protein gene was transfected into HK-2 cells, and the CrmA protein inhibited the activated caspases (their common pathway) and protected the HK-2 cells from apoptosis induced by cyclosporine.

Previous studies have shown that when cyclosporine of different concentrations is cocultured with renal tubular epithelial cells of human, pig, dog, and mouse, it could significantly induce apoptosis in a dose-dependent and time-dependent manner.^{5, 10, 39-41} Here, we confirmed that cyclosporine caused a concentration-dependent and time-dependent loss of cell viability in HK-2 cells. Cell viability was recovery at 0.01 to 10 µg/mL of cyclosporine when the coculture time was extended from 24 to 48 hours. When HK-2 cells cocultured with cyclosporine (> 200 µg/mL) at 24 or 48 hours, their activity was completely lost. Therefore, we speculate that the injury was reversible under 10 µg/mL of cyclosporine, though it was not at 10 µg/mL of cyclosporine and more.

Regarding clinical use of drugs, plasma concentration must be controlled in safe and effective dosage range. There have been studies that cyclosporine nephrotoxicity can be restored after withdrawal of cyclosporine.^42-43 However, this does not explain the reversible phenomenon of low-dose cyclosporine nephrotoxicity.

The p-CAGGS-CrmA was transfected into the HK-2 cells. Cytokine response modifier A protein mRNA was detected at 48 and 72 hours and reached its peak at 72 hours. Cytokine response modifier A protein was detected by Western blotting at 72 hours. This result shows that p-CAGGS-CrmA is transfected and expressed in HK-2 cells by Lipofectamine 2000. Cytokine response modifier A protein was a caspase inhibitor and broadly inhibited the activity of many caspases.

In this study, each group of cells (the p-CAGGS-CrmA group, the p-CAGGS group, and the nontransfected group) at 72 hours after transfection was treated with 1 µg/mL, 10 µg/mL, or 20 µg/mL of cyclosporine for 24 or 48 hours. We found that the HK-2 activity in CrmA-transfected group was significantly higher than in controls by MTT3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide) assay (P<.05). The apoptosis rate in CrmA-transfected group was significantly lower than it was in the control groups by fetal calf serum (P<.05). Cell activity and the rate of apoptosis were no different in the p-CAGGS-transfected group and nontransfected group (P < .05).

Our results indicate that CrmA has different degrees of protecting HK-2 cells from injury induced by cyclosporine at levels of 1 µg/mL, 10 µg/mL, and 20 µg/mL. Related studies have indicated that high expression of CrmA proteins in target cells inhibit cell apoptosis and protect them from injury induced by many chemicals, for example, cisplatin,⁴⁴ paclitaxel,⁴⁵ parthenolide,⁴⁶ and resveratrol.⁴⁷ Other studies show that target organ cells with high expression of CrmA are resistant to specific cytotoxic T lymphocyte,⁴⁸ inflammatory cytokines,⁴⁹ and hypoxic injury.⁵⁰ Adachi and associates found that CrmA gene transfection could significantly prolong the survival of a rat liver graft by DA-LEW orthotopic liver transplant.^51-52

In our study, we speculated that CrmA shows high expression in graft transplant and also can inhibit graft rejection, ischemia reperfusion injury, and cyclosporine nephrotoxicity; it can induce comprehensive tolerance of target organs and improve chronic graft dysfunction. These results will be confirmed in further studies.

References:

1. Healy E, Dempsey M, Lally C, Ryan MP. Apoptosis and necrosis: mechanisms of cell death induced by cyclosporine A in a renal proximal tubular cell line. Kidney Int. 1998;54(6):1955-1966.
2. De Hornedo JP, De Arriba G, Calvino M, Benito S, Parra T. Cyclosporin A causes oxidative stress and mitochondrial dysfunction in renal tubular cells. Nefrologia. 2007;27(5):565-573.
3. Daly PJ, Docherty NG, Healy DA, McGuire BB, Fitzpatrick JM, Watson RW. The single insult of hypoxic preconditioning induces an antiapoptotic response in human proximal tubular cells, in vitro, across cold storage. BJU Int. 2009;103(2):254-259.
4. Amore A, Emancipator SN, Cirina P, et al. Nitric oxide mediates cyclosporine-induced apoptosis in cultured renal cells. Kidney Int. 2000;57(4):1549-1559.
5. Cheng CH, Hsieh CL, Shu KH, Chen YL, Chen HC. Effect of calcium channel antagonist diltiazem and calcium ionophore A23187 on cyclosporine A-induced apoptosis of renal tubular cells. FEBS Lett. 2002;516(1-3):191-196.
6. Ghee JY, Han DH, Song HK, et al. The role of macrophage in the pathogenesis of chronic cyclosporine-induced nephropathy. Nephrol Dial Transplant. 2008;23(12):4061-4069.
7. Lim SW, Li C, Sun BK, et al. Long-term treatment with cyclosporine decreases aquaporins and urea transporters in the rat kidney. Am J Physiol Renal Physiol. 2004;287(1 56-1):F139-F151.
8. Kasap B, Soylu A, Kuralay F, et al. Protective effect of Epo on oxidative renal injury in rats with cyclosporine nephrotoxicity. Pediatr Nephrol. 2008;23(11):1991-1999.
9. Yang CW, Faulkner GR, Wahba IM, et al. Expression of apoptosis related genes in chronic cyclosporine nephrotoxicity in mice. Am J Transplant. 2002;2(5):391-399.
10. Jo SK, Lee SY, Han SY, et al. alpha-Melanocyte stimulating hormone (MSH) decreases cyclosporine a induced apoptosis in cultured human proximal tubular cells. J Korean Med Sci. 2001;16(5):603-609.
11. Shihab FS, Andoh TF, Tanner AM, Yi H. Expression of apoptosis regulatory genes in chronic cyclosporine nephrotoxicity favors apoptosis. Kidney Int. 1999;56(6):2147-2159.
12. Lee SH, Li C, Lim SW, et al. Attenuation of interstitial inflammation and fibrosis by recombinant human erythropoietin in chronic cyclosporine nephropathy. Am J Nephrol. 2005;25(1):64-76.
13. Pallet N, Bouvier N, Bendjallabah A, et al. Cyclosporine induced endoplasmic reticulum stress triggers tubular phenotypic changes and death. Am J Transplantation. 2008;8(11):2283-2296.
14. Chung BH, Li C, Sun BK, et al. Rosiglitazone protects against cyclosporine-induced pancreatic and renal injury in rats. Am J Transplant. 2005;5(8):1856-1867.
15. Thomas SE, Andoh TF, Pichler RH, et al. Accelerated apoptosis characterizes cyclosporine-associated interstitial fibrosis. Kidney Int. 1998;53(4):897-908.
16. Sheikh-Hamad D, Nadkarni V, Choi YJ, et al. Cyclosporine A inhibits the adaptive responses to hypertonicity a potential mechanism of nephrotoxicity. J Am Soc Nephrol. 2001;12(12):2732-2741.
17. Josephine A, Amudha G, Veena CK, Preetha SP, Rajeswari A, Varalakshmi P. Beneficial effects of sulfated polysaccharides from Sargassum wightii against mitochondrial alterations induced by Cyclosporine A in rat kidney. Mol Nutr Food Res. 2007;51(11):1413-1422.
18. Khan M, Shobha JC, Mohan IK, Naidu MUR, Prayag A, Kutala VKDAS-ODOj. Spirulina attenuates cyclosporine-induced nephrotoxicity in rats. J Appl Toxicol. 2006;26(5):444-451.
19. Chong FW, Chakravarthi S, Nagaraja HS, Thanikachalam PM, Lee N. Expression of transforming growth factor-beta and determination of apoptotic index in histopathological sections for assessment of the effects of Apigenin (4', 5', 7'- Trihydroxyflavone) on Cyclosporine A induced renal damage. Malays J Pathol. 2009;31(1):35-43.
20. Pallet N, Bouvier N, Legendre C, et al. Antiapoptotic properties of recombinant human erythropoietin protects against tubular cyclosporine toxicity. Pharmacological Research. 2010;61(1):71-75.
21. Fornoni A, Li H, Foschi A, Striker GE, Striker LJ. Hepatocyte growth factor but not insulin like growth factor I protects podocytes against cyclosporin A induced apoptosis. Am J Pathol. 2001;158(1):275-280.
22. Perez-Rojas J, Blanco JA, Cruz C, et al. Mineralocorticoid receptor blockade confers renoprotection in preexisting chronic cyclosporine nephrotoxicity. Am J Physiol Renal Physiol. 2007;292(1):F131-F139.
23. Disel U, Paydas S, Dogan A, Gulfiliz G, Yavuz S. Effect of colchicine on cyclosporine nephrotoxicity reduction of TGF beta overexpression apoptosis and oxidative damage an experimental animal study. Transplant Proc. 2004;36(5):1372-1376.
24. Palumbo GJ, Pickup DJ, Fredrickson TN, McIntyre LJ, Buller RM. Inhibition of an inflammatory response is mediated by a 38-kDa protein of cowpox virus. Virology. 1989;172(1):262-273.
25. Pickup DJ, Ink BS, Hu W, Ray CA, Joklik WK. Hemorrhage in lesions caused by cowpox virus is induced by a viral protein that is related to plasma protein inhibitors of serine proteases. Proc Natl Acad Sci U S A. 1986;83(20):7698-7702.
26. Garcia-Calvo M, Peterson EP, Leiting B, Ruel R, Nicholson DW, Thornberry NA. Inhibition of human caspases by peptide-based and macromolecular inhibitors. J Biol Chem. 1998;273(49):32608-32613.
27. Dobo J, Swanson R, Salvesen GS, Olson ST, Gettins PG. Cytokine response modifier a inhibition of initiator caspases results in covalent complex formation and dissociation of the caspase tetramer. J Biol Chem. 2006;281(50):38781-38790.
28. Atkinson EA, Barry M, Darmon AJ, et al. Cytotoxic T lymphocyte-assisted suicide. caspase 3 activation is primarily the result of the direct action of granzyme B. J Biol Chem. 1998;273(33):21261-21266.
29. Xiao Z, Shan J, Li C, et al. CrmA gene transfer rescued cyclosporine-induced renal cell apoptosis in graft kidney. Cell Immunol. 2010;265(1):6-8.
30. Darzynkiewicz Z, Juan G, Li X, Gorczyca W, Murakami T, Traganos F. Cytometry in cell necrobiology: analysis of apoptosis and accidental cell death (necrosis). Cytometry. 1997;27(1):1-20.
31. Frey T. Correlated flow cytometric analysis of terminal events in apoptosis reveals the absence of some changes in some model systems. Cytometry. 1997;28(3):253-63.
32. Ponticelli C. Cyclosporine: from renal transplantation to autoimmune diseases. Ann N Y Acad Sci. 2005;1051:551-558.
33. Xiao Z, Li C, Shan J, et al. Mechanisms of renal cell apoptosis induced by cyclosporine a: a systematic review of in vitro studies. Am J Nephrol. 2011;33(6):558-566.
34. Magnasco A, Rossi A, Catarsi P, et al. Cyclosporin and organ specific toxicity: clinical aspects, pharmacogenetics and perspectives. Curr Clin Pharmacol. 2008;3(3):166-173.
35. Benigni A, Bruzzi I, Mister M, et al. Nature and mediators of renal lesions in kidney transplant patients given cyclosporine for more than one year. Kidney Int. 1999;55(2):674-685.
36. Takeda A, Uchida K, Haba T, et al. Chronic cyclosporin nephropathy: long-term effects of cyclosporin on renal allografts. Clin Transplant 2001;15(suppl 5):22-29.
37. Seron D, Moreso F, Grinyo JM. Prevention and management of late renal allograft dysfunction. J Nephrol. 2001;14(2):71-79.
38. Marcen R, Pascual J, Teruel JL, et al. Outcome of cadaveric renal transplant patients treated for 10 years with cyclosporine: is chronic allograft nephropathy the major cause of late graft loss? Transplantation. 2001;72(1):57-62.
39. Esposito C, Fornoni A, Cornacchia F, et al. Cyclosporine induces different responses in human epithelial, endothelial and fibroblast cell cultures. Kidney Int. 2000;58(1):123-130.
40. Justo P, Lorz C, Sanz A, Egido J, Ortiz A. Intracellular mechanisms of cyclosporin A-induced tubular cell apoptosis. J Am Soc Nephrol. 2003;14(12):3072-3080.
41. Conti G, Amore A, Cirina P, Gianoglio B, Peruzzi L, Coppo R. Cyclosporin induces apoptosis of renal cells by enhancing nitric oxide synthesis modulating effect of angiotensin II inhibitors. Transplant Proc. 2001;33(1-2):276-277.
42. Li C, Lim SW, Sun BK, et al. Expression of apoptosis related factors in chronic cyclosporine nephrotoxicity after cyclosporine withdrawal. Acta Pharmacologica Sinica. 2004;25(4):401-411.
43. Martin M, Krichbaum M, Kaever V, Goppelt-Strube M, Resch K. Cyclosporin A suppresses proliferation of renal mesangial cells in culture. Biochem Pharmacol. 1988;37(6):1083-1088.
44. Takeda M, Kobayashi M, Shirato I, Osaki T, Endou H. Cisplatin-induced apoptosis of immortalized mouse proximal tubule cells is mediated by interleukin-1 beta converting enzyme (ICE) family of proteases but inhibited by overexpression of Bcl-2. Arch Toxicol. 1997;71(10):612-621.
45. Wang S, Wang Z, Dent P, Grant S. Induction of tumor necrosis factor by bryostatin 1 is involved in synergistic interactions with paclitaxel in human myeloid leukemia cells. Blood. 2003;101(9):3648-3657.
46. Zhang S, Ong CN, Shen HM. Involvement of proapoptotic Bcl-2 family members in parthenolide-induced mitochondrial dysfunction and apoptosis. Cancer Lett. 2004;211(2):175-188.
47. Byun HS, Song JK, Kim YR, et al. caspase-8 has an essential role in resveratrol-induced apoptosis of rheumatoid fibroblast-like synoviocytes. Rheumatology (Oxford). 2008;47(3):301-308.
48. Fujino M, Kawasaki M, Funeshima N, et al. CrmA gene expression protects mice against concanavalin-A-induced hepatitis by inhibiting IL-18 secretion and hepatocyte apoptosis. Gene Ther. 2003;10(20):1781-1790.
49. Millet I, Wong FS, Gurr W, et al. Targeted expression of the anti-apoptotic gene CrmA to NOD pancreatic islets protects from autoimmune diabetes. J Autoimmun. 2006;26(1):7-15.
50. Gurevich RM, Regula KM, Kirshenbaum LA. Serpin protein CrmA suppresses hypoxia-mediated apoptosis of ventricular myocytes. Circulation. 2001;103(15):1984-1991.
51. Adachi K, Li X, Kimura H, Amemiya H, Suzuki S. Prolonged survival of rat liver allografts with CrmA gene transfer. Transplant Proc. 2001;33(1-2):605.
52. Adachi K, Fujino M, Kitazawa Y, et al. Exogenous expression of Fas-ligand or CrmA prolongs the survival in rat liver transplantation. Transplant Proc. 2006;38(8):2710-2713.