 |
||||
| 2007 JASN IMPACT FACTOR 7.111 | HOME AUTHOR INFO EDITORIAL BOARD SUBSCRIBE FEEDBACK ALERTS HELP | |||
| CURRENT ISSUE | ARCHIVES | JASN Express | ONLINE SUBMISSION | |
|
||||
| 2007 JASN IMPACT FACTOR 7.111 | HOME AUTHOR INFO EDITORIAL BOARD SUBSCRIBE FEEDBACK ALERTS HELP | |||
| CURRENT ISSUE | ARCHIVES | JASN Express | ONLINE SUBMISSION | |
BASIC SCIENCE |
*Department of Experimental Medicine and Nephrology, Anthony Raine Laboratory and William Harvey Research Institute, St. Bartholomew’s and the Royal London School of Medicine, Queen Mary College, University of London, London, United Kingdom; Department of Pathology, University Medical Buildings, Aberdeen, Scotland, United Kingdom; Laboratory of Pharmacology, University of Lisbon, Lisbon, Portugal; and Department of Pathology, Royal London Hospital, London, United Kingdom
Correspondence to Dr. Edward J. Sharples, Department of Experimental Medicine and Nephrology, William Harvey Research Institute, John Vane Science Block, Charterhouse Square, London. Phone: 02007377480; Fax: 02073777003; E-mail: edsharps{at}doctors.org.uk
 |
Abstract |
|---|
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
|
Introduction |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
and the constitutively expressed subunit HIF-1
. Low oxygen tension adverts enzymatic prolyl-residue hydroxylation by prolyl-4-hydroxylase, which, in normoxia, serves as a signal for polyubiquitination and proteosomal degradation, thereby preventing von-Hippel-Lindau (VHL)-dependent HIF degradation, leading to nuclear accumulation of HIF-1 (2). HIF-1 controls the expression of several cytokines that mediate the adaptive response to ischemia, including vascular endothelial growth factor and glucose metabolism. The prolyl-4-hydroxylase requires iron as a co-factor, and cobalt mimics the effect of hypoxia on HIF-1 activation (3). Cobalt administration to rats caused upregulation of HIF-dependent proteins, including EPO, and diminished the degree of renal injury caused by ischemia-reperfusion (I/R), suggesting the HIF/EPO pathway may play an important role in ischemic preconditioning (4). EPO is upregulated in the brain and spinal cord after hypoxic stimuli and protects neurones against ischemic or oxidative injury in vivo and in vitro (5,6). The neuroprotective effects of EPO are dependent on EPO receptor–mediated JAK2 phosphorylation and NF-
B–dependent transcription of antiapoptotic genes, including endogenous inhibitors of apoptosis XIAP and cIAP-2 (7). In the retina, EPO upregulation is essential for hypoxic preconditioning via HIF-1 stabilization. The systemic administration of recombinant EPO also reduces the degree of retinal apoptosis induced by high-intensity light insult (8). EPO receptor–mediated intracellular signaling may involve nuclear translocation of the transcription factor NF-B and phosphorylation of Akt (protein kinase B) by phosphatidylinositol 3-kinase (PI3K) and the extracellular signal–regulated kinases 1 and 2. Although the peritubular fibroblasts are the major adult site for EPO production, the presence of the EPO receptor in many cell types in the kidney was only recently reported. The EPO receptor is present on proximal tubule epithelial cells, mesangial cells, and the glomerulus (9). Renal ischemia, whether caused by shock or during surgery or transplantation, is a major cause of acute renal failure. Although reperfusion is essential for the survival of ischemic tissue, there is good evidence that reperfusion itself causes additional cellular injury. Reperfusion initiates a complex and interrelated sequence of events that results in injury and the eventual death of renal cells as a result of a combination of both apoptosis and necrosis (10,11). The degree of cellular ATP and GTP depletion plays a crucial role in the determination of mode of cell death. Moderate ATP and GTP depletion favors apoptotic cell death and is associated with DNA fragmentation, p53 induction, and increased cysteine protease activity, whereas profound ATP depletion results in necrosis (12). Apoptotic cell death has been documented in animal models and human biopsies after renal I/R (13), and inhibition of apoptotic signaling and cell death ameliorates the associated injury and inflammation in a murine model (14). Several studies have documented caspase-3 activation after I/R injury in the kidney and in posthypoxic isolated proximal tubules (15,16). Renal I/R in animal models causes mitochondrial swelling and loss of membrane potential, which contributes to both necrotic and apoptotic forms of cell death. In vivo, DNA fragmentation and terminal deoxynucleotidyl transferase-mediated nick end labeling (TUNEL) staining, although not wholly specific for apoptotic cell death, are significantly elevated after 2 h of reperfusion (17,18). Ischemic renal injury is associated with an increase in the expression of both the antiapoptotic Bcl-2 family of proteins, Bcl-2 and Bcl-XL, and the proapoptotic proteins Bad, p53, FADD, and Bak in the distal and proximal tubules during the first 24 h, with the net effect determining the severity of injury and dysfunction (19).
Here, we report that EPO protects the rat kidney in a model of severe I/R injury, with inhibition of caspase activation and reduced apoptotic cell death. Thus, the systemic administration of recombinant EPO may reduce the renal injury and dysfunction caused by I/R in humans.
|
Materials and Methods |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
At the end of the reperfusion period (6 h in all groups unless stated), blood (1 ml) samples were collected via the carotid artery into tubes that contained serum gel. The samples were centrifuged (6000 x g for 3 min) to separate serum from which biochemical parameters were measured. All plasma samples were analyzed within 24 h after collection (Vetlab Services, Sussex, UK). Urine was collected throughout the reperfusion period, and the volume was recorded. Activity of urinary N-acetyl--D-glucosaminidase (NAG), a specific indicator of tubular damage, was also measured (Clinica Medica e Diagnostico Dr Joaquim Chaves, Lisbon, Portugal).
Histologic Evaluation and TUNEL Staining
Renal sections were prepared as described previously and used for assessment of renal I/R injury (20). Briefly, kidneys were removed from rats at the end of the experimental period and were cut in a sagittal section into two halves, which were fixed in immersion in 10% (wt/vol) formaldehyde in PBS (0.01 M; pH 7.4) at room temperature for 1 d. After dehydration using graded ethanol, pieces of kidney were embedded in Paraplast (Sherwood Medical, Mahwah, NJ) and cut in fine (8 µm) sections and mounted on glass slides. Sections were then deparaffinized with xylene, counterstained with hematoxylin and eosin, and viewed under a light microscope (Dialux 22; Leitz, Milan, Italy). One hundred intersections were examined for each kidney, and a score from 0 to 3 was given for each tubular profile: 0, normal histology; 1, tubular cell swelling, brush border loss, and nuclear condensation with up to one third nuclear loss; 2, as for score 1 but greater than one third and less than two thirds tubular profiles showing nuclear loss; and 3, greater than two thirds tubular profile showing nuclear loss. The histologic score for each kidney was calculated by addition of all scores, with a maximum score of 300. Sections were assessed quantitatively for apoptotic nuclei and graded for severity and extent of nuclear changes. TUNEL was performed as described previously, with a modification of the detection system using the ApopTag Plus kit (R&D Systems, Abingdon, UK), which adds an amplification step using streptavidin-biotin-immunoperoxidase staining to increase the sensitivity of the technique (21).
Cell Culture and Reagents
All reagents were obtained from Sigma-Aldrich Co. (Poole, Dorset, UK) unless otherwise stated. HK-2 cells (American Type Culture Collection, Manassas, VA), an immortalized human proximal tubule epithelial cell line, were grown and passaged in 75-cm2 cell culture flasks that contained DMEM Ham’s F12 media (1:1) supplemented with 5% FCS and antibiotics (100 U/ml penicillin G, 100 µg/ml streptomycin, and 0.25 µg/ml amphotericin). Subconfluent (80%) HK-2 cells were harvested and seeded into six-well tissue culture plates in 2 ml of growth medium. The cells were allowed to adhere for 18 h in an incubator at 37°C with 5% CO2 in 95% air. Immediately before experimental treatments, the medium was replaced with fresh medium. EPO (1 to 100 U/ml) or inhibitors (± vehicle) were added for 60 min preincubation, unless otherwise stated.
Analysis of DNA Fragmentation and Lactate Dehydrogenase
Monolayers of HK-2 cells were disrupted with a Teflon cell scraper. Cells were centrifuged at 2000 x g for 5 min, and the pellet was incubated on ice for 30 min in 200 µl of lysis buffer (PBS [pH 7.4] containing 10 µM digitonin). After centrifugation at 16,000 x g for 15 min at 4°C, the supernatant was diluted 1:10 with lysis buffer. Diluted lysate (20 µl) was assayed in duplicate for microsomal DNA fragments using a commercial ELISA (Cell Death Detection ELISA Plus; Roche Diagnostics, Lewes, UK). Results were expressed as multiples of control corrected to amount of total protein in the lysate (Bradford method).
Lactate dehydrogenase (LDH) release into the supernatant was assayed using a commercial colorimetric method (Cytotoxicity Detection Kit; Roche Diagnostics). Results were expressed as a percentage of total cellular LDH per well measured from cells lysed in 1% Triton-X100 and corrected with appropriate media controls and total protein.
Western Blot Analysis
For in vivo experiments, frozen samples of harvested kidneys were washed in ice-cold PBS and homogenized in ice-cold lysis buffer that contained 25 mmol/L EDTA, 20 mmol/L EGTA, 63.2 mmol/L imidazole-HCl, and 10 mmol/L 2-mercaptoethanol (pH 7.3). After centrifugation at 16,000 x g, the supernatant was aspirated and protein was quantified by the Bradford method. In vitro, cell pellets were washed in ice-cold PBS and incubated for 15 min on ice in modified RIPA buffer (150 mM NaCl, 10 mM Tris, 0.1% SDS, 1.0% NP40, 1.0% sodium deoxycholate, 5 mM EDTA, and 100 µM sodium orthovanadate [pH 7.2]). Mitochondria-rich cytosolic fractions were obtained by homogenizing cell pellets in extraction buffer (50 mM HEPES [pH 7.5], 1 M mannitol, 350 mM sucrose, and 5 mM EGTA) followed by centrifugation at 600 x g. The resulting supernatant was centrifuged at 11,000 x g for 10 min. Twenty to 40 µg of protein per lane was electrophoresed on precast graduated (4 to 12%) NuPAGE SDS gel (Invitrogen, San Diego, CA). Proteins were transferred to polyvinylidene difluoride membranes, which were subsequently blocked with Tris-buffered saline/Tween-20 (TBS-T) containing 5% BSA and then incubated for 3 h with antibodies against caspase-3, Bcl-XL (Autogen Biotech, Calne, UK), and XIAP (R&D Systems Europe, Abingdon, UK). After washing, the membranes were incubated for 1 h with horseradish peroxidase–conjugated secondary antibodies (diluted 1:2000 in TBS-T/1% BSA) and washed, and then immunoreactivity was visualized using enhanced chemiluminescence.
Caspase Activity
The activity of caspase-3 was measured using the fluorometric substrate Ac-DEVD-AMC as described previously (22). Fifty micrograms of cellular protein was incubated with 50 µM substrate in caspase assay buffer (213.5 mM HEPES [pH 7.5], 31.25% sucrose, and 0.3125% CHAPS) for 1 h, and fluorescence was measured on a microplate reader (Fluostar Galaxy; BMG Laboratory Technologies, Aylesbury, UK), with excitation at 380 nm and emission at 460 nm. For each sample, four replicates were assayed with two replicates that contained 50 µM of the caspase-3 inhibitor (Ac-DEVD-CHO) and the remaining pair that contained vehicle (DMSO). The activities of caspase-8 and caspase-9 were determined in the same way, using Ac-IETD-AMC and Ac-LEHD-AMC as respective substrates with Ac-IETD-CHO and Ac-LEHD-CHO as inhibitors, respectively. Fluorescence readings from wells that contained inhibitor were subtracted from total fluorescence, and results were calculated as nmol AMC/min per mg protein (Bradford method).
Statistical Analyses
All values described in the text and figures are expressed as the mean ± SEM for the number of observations. Each data point representing biochemical measurements was obtained from up to 10 to 12 separate animals. For histologic scoring, caspase activity assays, and apoptosis scoring, each data point represents analysis of kidneys taken from six individual animals. Statistical analysis was performed using GraphPad Prism 3.01/Instat 1.1 (GraphPad Software, San Diego, CA). Data were analyzed using one-way ANOVA followed by Dunnett post hoc test or Kruskal-Wallis ANOVA for nonparametric data. P < 0.05 was considered significant.
|
Results |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
). Administration of EPO to sham-operated animals did not have any effect on markers of renal function despite an increase in urine flow rate (Figure 1). EPO administration did not have an effect on mean BP in either sham-operated or I/R animals (shown supplementary data, Appendix 1). EPO administration to sham-operated animals had no effect on markers of tubular or reperfusion injury (urinary NAG activity and serum AST; Figure 2), histologic changes (Figure 3), or the number of apoptotic proximal tubular epithelial cells (Figure 4).
|
|
|
|
Effect of EPO on Tubular and Reperfusion Injury Caused by Renal I/R
In comparison with values obtained from sham-operated animals, renal I/R produced a significant increase in urinary NAG activity (Figure 2A), consistent with tubular injury. Renal I/R also produced a significant increase in serum AST, a marker of tubular reperfusion injury (Figure 2B). Administration of EPO caused a significant reduction in urinary NAG activity and serum AST levels (Figure 2), suggesting a marked reduction in the tubular and reperfusion injury associated with renal I/R, respectively.
Effects of EPO on Histologic Alterations Caused by Renal I/R
Renal I/R caused marked alterations in renal histology compared with kidneys taken from sham-operated animals. Specifically, this included widespread degeneration of tubular architecture, tubular dilation, swelling and necrosis, luminal congestion with loss of brush border, and an infiltration of polymorphonuclear neutrophils (Figure 3, A and B). In contrast, renal sections obtained from animals that were treated with EPO preischemia or prereperfusion demonstrated marked reduction of the histologic features of renal injury (Figure 3C). EPO administration 30 min after the onset of reperfusion was still associated with a significant reduction in injury, but this reduction was less marked than in the other treatment groups (Figure 3D).
EPO Administration Reduced Tubular Apoptosis and Necrotic Cell Death
Proximal tubular epithelial cell apoptosis was quantified on renal sections and confirmed by TUNEL staining (data not shown). Kidneys from animals that were subjected to renal I/R showed extensive nuclear changes consistent with apoptotic cell death. In comparison, kidneys from sham-operated animals had no evidence of apoptosis and were negative for TUNEL-positive cells. EPO administration before ischemia and prereperfusion significantly reduced the extent of apoptotic cell death (P < 0.05; Figure 4A). EPO administration 30 min after the onset of reperfusion caused only a small reduction in the number of apoptotic cells, but there was a reduction in the degree of necrotic cell death (Figure 4B).
Effect of EPO on Caspase Activity and Protein In Vivo
Caspase-3 activity (nmol/min per mg protein at 37°C) was significantly increased in the homogenates of kidneys at 6 h in kidneys that were subjected to renal I/R (Figure 5A), when compared with sham-operated animals (P < 0.01). The elevation in caspase-3 activity was significantly reduced by EPO administration, either preischemia (P < 0.05) or just before reperfusion (P < 0.05). EPO treatment 30 min after reperfusion was associated with a nonsignificant reduction in caspase-3 activity, consistent with the higher levels of apoptosis observed in these kidneys (Figure 5A). Caspase-3 protein in kidney tissue, determined by Western blot analysis, showed that a 17-kD band, representing the active caspase-3 subunit, was markedly increased by renal I/R compared with sham-operated animals. The amount of this caspase-3 subunit was diminished in both early EPO treatment groups (Figure 5B). Immunohistochemical staining using a specific antibody to the cleaved active fragment of caspase-3 confirmed these observations. There was widespread positive cytoplasmic and perinuclear staining in both cortical and medullary tubules after I/R, and the severity and distribution of staining was greatly reduced in animals that received EPO either preischemia or prereperfusion but only partially when EPO was administered later in reperfusion (Figure 5C). EPO treatment was associated with significant reduction in the increase in both caspase-8 and caspase-9 activity observed after I/R (supplementary data, appendix 2).
|
|
|
|
|
Discussion |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
What, then, is the mechanism(s) by which EPO protects the kidney against I/R injury? One could argue that an increase in renal blood flow may contribute to or even account for the observed protective effects of EPO. This is unlikely, however, as we were unable to demonstrate any significant hemodynamic effect of acute administration of EPO. Although EPO increased urine flow rate in sham-operated rats, suggesting a possible effect on cortical perfusion and intraglomerular pressure, creatinine clearances were similar and there was no difference in tubular function. This observation is backed by the study by Huang et al. (23), who demonstrated no effect on cortical and papillary perfusion in normal rats after acute EPO administration.
We propose here that EPO acts directly on proximal tubule epithelial cells against I/R injury. This proposal is based on the following key findings: EPO protected human proximal tubule epithelial cells against the cell injury and death caused by serum starvation and oxidative stress (hydrogen peroxide), the latter of which is most likely to simulate reperfusion injury. We then investigated whether the protection by EPO of proximal tubule epithelial cells is secondary to the activation of EPO receptors and due to an antiapoptotic effect of EPO in these cells. We demonstrated that the protection of proximal tubule epithelial cells by EPO was attenuated by an inhibitor of JAK2 signaling. Thus, the activation of the EPO receptor is essential in the observed protective effect of EPO. Activation of JAK2 by EPO leads (in endothelial cells and neuronal cells) to the activation of PI3K and Akt phosphorylation (24). Once activated, Akt activates multiple targets with antiapoptotic effects, including phosphorylation of Bad, Bax, caspase-9, and GSK-3, maintenance of mitochondrial membrane potential and preservation of glycolysis and ATP synthesis (25). Inhibition of PI3K abolishes the protective effects of EPO in human proximal tubule epithelial cells and inhibited the specific serine-473 phosphorylation of AKT induced by EPO. Thus, we suggest that EPO protects proximal tubule cells against injury by activating the EPO receptor, resulting in activation of PI3K and ultimately Akt. We demonstrated that EPO inhibits apoptotic cell death in proximal tubule epithelial cells (as determined by DNA fragmentation) and in higher doses causes significant proliferation despite serum-free conditions. Most notably, EPO administration was associated with the upregulation of Bcl-XL and XIAP and reduction of capase-3 activation. XIAP may be an important mediator of EPO’s protective effects in I/R injury, because it acts on both death receptor (Fas-FasL) mediated and mitochondrial pathways of caspase activation, as well as directly inhibits the activation of caspase-3, -7, and -9 (28). XIAP may also have an indirect antiapoptotic effect by inducing p21cip1, leading to cell cycle arrest at G1/S transition, which has been shown to be protective in renal I/R (29,30).
Taken together, these in vitro findings support the conclusion that EPO directly protects proximal tubule epithelial cells by (1) activating EPO receptor/JAK-2 kinase, (2) activating PI3K leading to activation of Akt, (3) upregulation of Bcl-XL and XIAP, and (4) preventing the activation of caspase-3 and ultimately apoptosis. We confirm that EPO reduced the activation of caspase-3 caused by I/R of the kidney by inhibiting the activation of the caspase cascade both through the mitochondrial caspase-9 pathway and possibly to a lesser degree the death receptor–mediated activation of caspase-8. The death receptors Fas (CD95), TNF-R1, and CD27 have been implicated in the pathogenesis of renal I/R injury (31), and TNF- is induced after reperfusion by the upregulation of NF-B via p38. We saw significant activation in mean caspase-8 activity after I/R. The increase in caspase-8 activity was less than the increase in caspase-9 activity. This difference may be due to differences in the relative sensitivity of the assay but may also be attributed to changes in mitochondrial membrane potential leading to caspase-9 activation beginning during ischemia, whereas the death receptor pathway may be activated only at a later time point during reperfusion (30).
In summary, we demonstrate here that a single intravenous bolus injection of recombinant human EPO, either preischemia or just before the onset of reperfusion, attenuates renal I/R injury in rats, via the inhibition of proapoptotic caspase activation. We propose that the nonhemopoietic pleiotropic effects of EPO have major implications in the treatment of I/R injury to the kidney, in particular, and possibly other organs.
|
Acknowledgments |
|---|
|
References |
|---|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
|---|
B signalling cascades. Nature 412: 641–647, 2001[CrossRef][Medline]
B: New survival pathways disabled by caspase-mediated cleavage during apoptosis of human endothelial cells. Circ Res 88: 282–290, 2001This article has been cited by other articles:
 |
|
 |
|
  C. Rosenberger, S. Rosen, A. Shina, U. Frei, K.-U. Eckardt, L. A. Flippin, M. Arend, S. J. Klaus, and S. N. Heyman Activation of hypoxia-inducible factors ameliorates hypoxic distal tubular injury in the isolated perfused rat kidney Nephrol. Dial. Transplant., November 1, 2008; 23(11): 3472 - 3478. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 B. Bi, J. Guo, A. Marlier, S. R. Lin, and L. G. Cantley Erythropoietin expands a stromal cell population that can mediate renoprotection Am J Physiol Renal Physiol, October 1, 2008; 295(4): F1017 - F1022. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 A. B. Sanz, B. Santamaria, M. Ruiz-Ortega, J. Egido, and A. Ortiz Mechanisms of Renal Apoptosis in Health and Disease J. Am. Soc. Nephrol., September 1, 2008; 19(9): 1634 - 1642. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Y. Yokomaku, T. Sugimoto, S. Kume, S.-i. Araki, K. Isshiki, M. Chin-Kanasaki, M. Sakaguchi, N. Nitta, M. Haneda, D. Koya, et al. Asialoerythropoietin Prevents Contrast-Induced Nephropathy J. Am. Soc. Nephrol., February 1, 2008; 19(2): 321 - 328. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
V. Schwenger, C. Morath, and M. Zeier Use of Erythropoietin after solid organ transplantation Nephrol. Dial. Transplant., September 1, 2007; 22(suppl_8): viii47 - viii49. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
A. Mitra, S. Bansal, W. Wang, S. Falk, E. Zolty, and R. W. Schrier Erythropoietin ameliorates renal dysfunction during endotoxaemia Nephrol. Dial. Transplant., August 1, 2007; 22(8): 2349 - 2353. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Menne, J.-K. Park, N. Shushakova, M. Mengel, M. Meier, and D. Fliser The Continuous Erythropoietin Receptor Activator Affects Different Pathways of Diabetic Renal Injury J. Am. Soc. Nephrol., July 1, 2007; 18(7): 2046 - 2053. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
W. H. Horl, Y. Vanrenterghem, P. Aljama, P. Brunet, R. Brunkhorst, L. Gesualdo, I. Macdougall, C. Wanner, and B. Wikstrom OPTA: Optimal treatment of anaemia in patients with chronic kidney disease (CKD) Nephrol. Dial. Transplant., June 1, 2007; 22(suppl_3): iii20 - iii26. [Full Text] [PDF]
|
|
|
|
|
|
S.-H. Park, M.-J. Choi, I.-K. Song, S.-Y. Choi, J.-O. Nam, C.-D. Kim, B.-H. Lee, R.-W. Park, K. M. Park, Y.-J. Kim, et al. Erythropoietin Decreases Renal Fibrosis in Mice with Ureteral Obstruction: Role of Inhibiting TGF-beta-Induced Epithelial-to-Mesenchymal Transition J. Am. Soc. Nephrol., May 1, 2007; 18(5): 1497 - 1507. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
I. Kojima, T. Tanaka, R. Inagi, H. Kato, T. Yamashita, A. Sakiyama, O. Ohneda, N. Takeda, M. Sata, T. Miyata, et al. Protective Role of Hypoxia-Inducible Factor-2{alpha} against Ischemic Damage and Oxidative Stress in the Kidney J. Am. Soc. Nephrol., April 1, 2007; 18(4): 1218 - 1226. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 W. M. Bernhardt, M. S. Wiesener, A. Weidemann, R. Schmitt, W. Weichert, P. Lechler, V. Campean, A. C. M. Ong, C. Willam, N. Gretz, et al. Involvement of Hypoxia-Inducible Transcription Factors in Polycystic Kidney Disease Am. J. Pathol., March 1, 2007; 170(3): 830 - 842. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 S. K. Jo, M. H. Rosner, and M. D. Okusa Pharmacologic Treatment of Acute Kidney Injury: Why Drugs Haven't Worked and What Is on the Horizon Clin. J. Am. Soc. Nephrol., March 1, 2007; 2(2): 356 - 365. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Z. Aydin, A. J. van Zonneveld, J. W. de Fijter, and T. J. Rabelink New horizons in prevention and treatment of ischaemic injury to kidney transplants Nephrol. Dial. Transplant., February 1, 2007; 22(2): 342 - 346. [Full Text] [PDF]
|
|
|
|
|
|
A. Grenz, T. Eckle, H. Zhang, D. Y. Huang, M. Wehrmann, C. Kohle, K. Unertl, H. Osswald, and H. K. Eltzschig Use of a hanging-weight system for isolated renal artery occlusion during ischemic preconditioning in mice Am J Physiol Renal Physiol, January 1, 2007; 292(1): F475 - F485. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
Z. P. Shaik, E. K. Fifer, and G. Nowak Protein kinase B/Akt modulates nephrotoxicant-induced necrosis in renal cells Am J Physiol Renal Physiol, January 1, 2007; 292(1): F292 - F303. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 T. B. Drueke, F. Locatelli, N. Clyne, K.-U. Eckardt, I. C. Macdougall, D. Tsakiris, H.-U. Burger, A. Scherhag, and the CREATE Investigators Normalization of Hemoglobin Level in Patients with Chronic Kidney Disease and Anemia N. Engl. J. Med., November 16, 2006; 355(20): 2071 - 2084. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
K. E. Jie, M. C. Verhaar, M.-J. M. Cramer, K. van der Putten, C. A. J. M. Gaillard, P. A. Doevendans, H. A. Koomans, J. A. Joles, and B. Braam Erythropoietin and the cardiorenal syndrome: cellular mechanisms on the cardiorenal connectors Am J Physiol Renal Physiol, November 1, 2006; 291(5): F932 - F944. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
W. M. Bernhardt, V. Campean, S. Kany, J.-S. Jurgensen, A. Weidemann, C. Warnecke, M. Arend, S. Klaus, V. Gunzler, K. Amann, et al. Preconditional Activation of Hypoxia-Inducible Factors Ameliorates Ischemic Acute Renal Failure J. Am. Soc. Nephrol., July 1, 2006; 17(7): 1970 - 1978. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. Wang, M. P. Biju, M.-H. Wang, V. H. Haase, and Z. Dong Cytoprotective Effects of Hypoxia against Cisplatin-Induced Tubular Cell Apoptosis: Involvement of Mitochondrial Inhibition and p53 Suppression J. Am. Soc. Nephrol., July 1, 2006; 17(7): 1875 - 1885. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
E. Spandou, I. Tsouchnikas, G. Karkavelas, E. Dounousi, C. Simeonidou, O. Guiba-Tziampiri, and D. Tsakiris Erythropoietin attenuates renal injury in experimental acute renal failure ischaemic/reperfusion model Nephrol. Dial. Transplant., February 1, 2006; 21(2): 330 - 336. [Abstract] [Full Text] [PDF]
|
|
|
|
P < 0.05; 
