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-Catenin and LEF-1
Renal Section, Boston Medical Center, Boston University School of Medicine, Boston, Massachusetts.
Correspondence to: Dr. John H. Schwartz, Evans Biomedical Research Center, Renal Section, 5th Floor, 650 Albany Street, Boston, MA 02118. Phone: 617-638-7321; Fax: 617-638-7326; E-mail: jhsch{at}bu.edu
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Abstract |
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-catenin, and 
-catenin, probably as intact cadherin-catenin complexes from the basolateral membrane of tubular cells. The relative amounts of all three proteins increased in the Triton X-100–insoluble fraction of cell lysates and decreased in the Triton X-100–soluble pool. These changes were reversed with repletion of cell ATP. It is additionally shown that ATP depletion induces nuclear translocation of -catenin and T cell factor (TCF)/lymphoid enhancer factor–1 (LEF-1), a transcriptional factor with which -catenin associates. The redistribution of the ZA proteins as intact E-cadherin-catenin complexes from the plasma membrane facilitates the rapid recovery of the ZA after sublethal ischemic injury. The translocation of -catenin and TCF/LEF-1 to the nucleus indicates that ATP depletion may activate the wnt/wingless signal transduction pathway. Thus, entirely novel evidence is provided that both of the known roles of -catenin, as a structural part of the ZA and as a component of the wnt/wingless pathway, play a role after sublethal ischemic injury to tubular cells. It is also speculated that the nuclear translocation of -catenin and TCF/LEF-1 modulates gene expression after ischemic injury and may contribute to events necessary for renal regeneration and repair after ischemic injury. |
Introduction |
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The ZA of renal epithelial cells is composed of complexes of E-cadherin and catenins. E-cadherin is a transmembrane glycoprotein. The extracellular domain of E-cadherin from adjacent cells bind to each other in a homophilic, calcium-dependent manner (9). The intracellular domain of E-cadherin is connected to the actin cytoskeleton by the catenin family of proteins that include -, -, and 
-catenin. Catenins (either - and -catenin or - and -catenin) link E-cadherin to the actin cytoskeleton via -actinin, an intermediary actin-binding protein. These interactions are essential for structural stability of the ZA and the functional integrity of the ZO (5,6,10).
In addition to forming an integral part of the ZA, -catenin has a second discrete function as a component of the wnt/wingless signaling pathway (11–14). Activation of the wnt/wingless pathway leads to translocation of -catenin to the nucleus in association with members of the T cell factor (TCF)/lymphoid enhancer factor (LEF) DNA-binding transcription factor family (15). -catenin and TCF/LEF-1 form a complex that induces transcription and expression of specific TCF/LEF-1–responsive genes (11–13,16–18). Although initially described in Drosophila (15), homologues of the wnt/wingless pathway are also present in mammalian cells. The activity of the wnt pathway appears to be determined by the level of "transcriptionally competent" -catenin that accumulates within the cytosol (19). Although the mechanisms regulating the amount of transcriptionally competent -catenin in the cytosol available are complex and remain incompletely defined, the amount of E-cadherin appears to play a role. It is therefore possible that a decrease in E-cadherin–mediated adhesion of -catenin induced by ATP depletion could potentially activate the wnt signaling pathway by increasing availability of transcriptionally competent catenin (19).
This study had two interrelated goals. One was to determine the fate of components of the ZA, E-cadherin, and -, -, and -catenin after ATP depletion, and the other was to examine our hypothesis that ATP depletion activates the wnt-signaling pathway. We demonstrate that ATP depletion results in the reversible withdrawal from the ZA of E-cadherin and the catenins, probably as intact complexes, and an increase of these complexes in the Triton X-100–insoluble pool. We also provide novel evidence that ATP depletion is associated with the translocation of both -catenin and LEF-1 to the nucleus. Thus, ATP depletion not only leads to loss of the ZA and dysfunction of the tight junction, but it also appears to activate the wnt pathway. We hypothesize that activation of this signaling pathway by ATP depletion may contribute to the changes in gene expression that occur in tubular cells during the recovery phase after ischemic renal injury (20,21).
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Materials and Methods |
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-catenin, anti--catenin, and anti-actin (Sigma, St Louis, MO); goat anti–LEF-1 (Santa Cruz Biotech, Santa Cruz, CA); and mouse anti–E-cadherin (Transduction Laboratories, Lexington, KY). Secondary antibodies were rabbit or mouse IgG conjugated with horseradish peroxidase (Sigma) for immunoblotting and donkey anti-mouse or goat anti-rabbit IgG conjugated with indocarbocyanine (CY3) (Sigma) for immunohistochemistry.
ATP Depletion-Repletion
Opossum kidney (OK) cells were grown to confluence on glass coverslips and plastic 60-mm dishes in Dulbecco’s modified Eagle’s medium (DMEM) containing 5% fetal calf serum/5% Nu Serum (Collaborative Research, Bedford, MA) at 37°C/5% CO2. Monolayers were washed three times with phosphate-buffered saline (PBS) containing 1 mM Ca++ and 1 mM Mg++ to remove residual substrates in the medium. Cells were incubated at 37°C in either DMEM containing 10 mM glucose (control cells) or in glucose-free DMEM containing 10 mM sodium cyanide (NaCN) (ATP-depleted cells). After 70 min of ATP depletion, 10 mM glucose was added to the medium (ATP-repleted cells).
Cell ATP
Cell ATP levels were determined with a luciferase assay as described previously by our group (3).
Immunohistochemistry
OK cells, grown to confluence on glass coverslips, were prepared as control, ATP-depleted, or ATP-repleted samples and then fixed by incubation at 37°C for 15 min in 3% formaldehyde in PBS containing 1mM Ca2+ and 1mM Mg2+. The cells were then permeabilized by 0.1% Triton X-100 in PBS for a 5-min interval. Coverslips were incubated in primary antibody (1:100 dilution) for 1 h at room temperature or nonimmune serum before three 5-min washes in 1 ml of PBS and then incubation for 1 h at room temperature in secondary antibody (1:500 dilution). After two washes with PBS, coverslips were mounted onto glass slides with Elvanol (DuPont, Wilmington, DE). Images were examined under fluorescence microscopy with an Olympus (Tokyo, Japan) epifluorescence microscope (magnification, x400). In control studies, fixed monolayers of OK cells incubated with nonimmune rabbit or mouse serum and appropriate secondary antibody were fluorescence-negative.
Separation of Cell Lysate into Detergent-Soluble and -Insoluble Cell Fractions.
Triton X-100–soluble and –insoluble cell fractions were obtained by incubating OK cell monolayers for 30 min at 4°C in CSK buffer. CSK consists of 0.5% Triton X-100, 50 mM NaCl, 300 mM sucrose, 10 mM Pipes, 3 mM MgCl2, 1 mM Na3VO4, 10 mM Na4P2O7, 50 mM NaF, pH 7.4, to which 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml aprotinin, and 10 µg/ml pepstatin were added just before use. To separate Triton X-100–soluble from –insoluble fractions, cell lysates were centrifuged for 15 min at 4°C at 20,000 x g in a Sorval SS-34 rotor (Clearwater, MN). Supernatants (representing the soluble fractions) were aliquoted into separate eppendorfs. The pellets (insoluble fractions) were resuspended and homogenized in sodium dodecyl sulfate–immunoprecipitation (SDS-IP) buffer containing 10mM Tris HCl, 2mM ethylenediaminetetraacetic acid, 1% SDS, 1 mM Na3VO4, pH 7.4, 0.1 mM N-octylglucopyranoside, 10 µg/ml aprotinin, 10 µg/ml pepstatin, and 1 mM phenylmethylsulfonyl fluoride.
Immunoblotting
Triton X-100–extracted (soluble and insoluble) fractions containing equal amounts of protein (40 µg) were boiled for 5 min in sample buffer and loaded onto 7% polyacrylamide SDS gels. Proteins were electrophoretically transferred to nitrocellulose filters that were then washed in TBST (150 mM NaCl, 100 mM Tris HCl, pH 7.5, 0.05% Tween 20) and blocked for 1 h in TBST containing 5% wt/vol nonfat powdered milk (MTBST). Filters were incubated with primary antibody in a 1:1000 dilution at 4°C overnight and then washed 3 times with TBST for 15 min each. After reblocking with MTBST for 1 h, filters were incubated in a 1:2000 dilution of secondary antibody in 2% TBST for 2 h at room temperature. After three washes with TBST, bound antibody was detected using the ECL enhanced chemiluminescence system (Amersham Pharmacia Biotech, Piscataway, NJ). The relative abundance of the protein was quantified by densitometric analysis. Densitometry was performed from scanned images, using the NIH Images 1.61 program.
Immunoprecipitation
Samples were adjusted to protein concentrations of 300 µg/ml with lysis buffer. For preclearing, 5 µl of nonimmune rabbit or mouse serum and 20 µl of protein A agarose beads were added to 1-ml samples. These mixtures were incubated for 1 h at 4°C and centrifuged for 2 min at 13,000 rpm in an eppendorf centrifuge at room temperature. The supernatants were incubated with 10 µl of primary antibody at 4°C for 30 min. The amount of primary antibody used was shown in preliminary study to be saturating and completely removed the detectable antigen form the homogenate. Protein A agarose beads were pretreated by incubating 60 µl of beads in 1 ml of PBS/0.1% bovine serum albumin with 5 µl of rabbit anti-mouse antibody at 4°C for 60 min and then washed three times in PBS. Aliquots of 60 µl of pretreated beads were added to samples that were then incubated at 4°C overnight. The beads were pelleted by centrifugation (12,000 x g) and washed three times with high stringency buffers containing 0.1% SDS, 1% deoxycholic acid, Triton X-100, and various concentrations of NaCl and sucrose. Beads suspended in 60 µl of sample buffer were boiled for 5 min and analyzed by SDS–polyacrylamide gel electrophoresis (PAGE) immunoblot analysis. E-cadherin, catenin, and LEF-1 were not identified when beads from the initial preclearing procedure were subjected to immunoblot analysis for these epitopes.
Isolation of Nuclear Cell Fraction
The nuclear fraction of cultured OK cells was isolated by using previously described methods (22). The purity of the preparation was assessed at each step by phase-contrast microscopy. The final preparation had no observable intact cells and consisted of bare nuclei. The remaining non-nuclear faction contained cytosolic, cellular organelles and plasma membrane components.
Statistical Analyses
Densitometric values from scanned immunoblots were expressed as averages of percent distribution between soluble and insoluble fractions for each sample or averages of absolute values (nuclear isolates). Data were expressed as mean ± SE. Groups were compared using a paired two-tailed t test. P < 0.05 was defined as significant.
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Results |
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-catenin, or -catenin (Figure 2). In control cells, all three ZA proteins were localized predominantly at the basolateral membrane (Figure 2, upper panel). ATP depletion caused a marked decrease in the amount of all three ZA proteins at the basolateral membrane and an increase of these proteins within cytosolic aggregates (Figure 2, middle panels). In addition, ATP depletion induced a striking increase in the intensity of nuclear staining of -catenin (Figure 2B, middle panel) without any change in nuclear staining of either E-cadherin or -catenin (Figure 2, A and C, middle panel). After ATP repletion, the locality of E-cadherin, -catenin, and -catenin were largely restored to the basolateral membrane (Figure 2, A, B, and C, lower panels). However, substantial nuclear staining of -catenin persisted during ATP-repletion (Figure 2B, lower panel). Control, ATP-depleted, and ATP-repleted cell monolayers when reacted with nonimmune serum, and secondary antibody were essentially unstained (Figure 2D).
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- and -catenin. Aliquots of whole cell homogenates containing equal amounts of protein (60 µg) obtained from control MPT cell monolayers were subjected to SDS-PAGE and immunoblotted. As shown in Figure 3, all three antibodies are highly specific, and each identifies a single band of protein at the appropriate molecular weight (Figure 3). When nonimmune serum from mouse, rabbit, or goat was used instead of primary antibody no bands were detected when equivalent amounts of homogenate was subjected to Western blot analyses (Figure 3).
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-catenin and -catenin were seen in ATP-depleted monolayers (Figure 4). The Triton X-100 distribution of E-cadherin and -catenin returned to values comparable to control with ATP repletion (Figure 4). By contrast, the amount of -catenin in the Triton X-100–insoluble fraction remained somewhat elevated after ATP repletion (Figure 4). These data suggest that ATP depletion results in an increase in the relative amount of E-cadherin and catenins bound to the cytoskeleton.
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Co-Immunoprecipitation Studies
To evaluate the degree to which the catenins and E-cadherin remain complexed or dissociated from each other before and after ATP depletion, we analyzed the amount of - and -catenin that could be co-immunoprecipitated with E-cadherin or the amount of E-cadherin and -catenin that could be co-immunoprecipitated with -catenin from both the Triton X-100–soluble and –insoluble fractions. We had to use SDS to consistently clarify and solubilize the Triton X-100–insoluble pellet suspension for immunoprecipitation. Therefore, we initially had to determine the effect of the addition of SDS on complex stability in a Triton X-100–soluble fraction. SDS at concentrations up to 1.5% did not appreciably reduce the amount of -catenin bound to E-cadherin as assessed by co-immunoprecipitation of these two proteins. Only when the concentration of SDS exceeded 2% did the amount of -catenin co-immunoprecipitated by anti–E-cadherin antibody decrease (Figure 5A). On the basis of these preliminary experiments, we elected to use a 1% SDS to solubilize the Triton X-100–insoluble fractions.
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-catenin obtained from Triton X-100–soluble and –insoluble fractions of control and ATP-depleted lysates were separated by SDS-PAGE and probed with antibodies for either E-cadherin, -catenin, or -catenin. Intact E-cadherin complexes always contain -catenin and either - and or -catenin, but never both of these latter catenins (7). Therefore, the amount of -catenin immunopecipitated by E-cadherin or the amount of E-cadherin immunoprecipitated by -catenin represents the sum of all intact E-cadherin–catenin complexes. The effect of ATP depletion on the distribution of cadherin-catenin complexes within Triton X-100–insoluble and –soluble pools is shown by a representative immunoblot in Figure 5B. These changes were also quantified by densitometric analysis of six immunoblots (Figure 5C). The relative amount of E-cadherin or -catenin co-immunoprecipitated from ATP-depleted monolayers increased in the Triton X-100–insoluble and decreased in the Triton X-100–soluble fractions (Figure 5, B and C). However, the total amount of E-cadherin co-immunoprecipitated with -catenin, and the total amount of -catenin co-immunoprecipitated with E-cadherin (determined by adding the amounts present in insoluble + soluble fractions) was found to be comparable in control and ATP-depleted monolayers (Figure 5B). The change in the amount of -catenin co-immunoprecipitated with either -catenin or E-cadherin from Triton X-100–soluble and –insoluble pool with ATP depletion was similar to that observed above (Figure 5B). Also by densitometric analysis the total amount of -catenin co-immunoprecipitated with E-cadherin or -catenin after ATP depletion was not reduced compared with control (90.5 ± 3.4% and 91.7 ± 4.1% of control respectively; n = 5; P > 0.08).
Effects of ATP Depletion on Nuclear Content of -catenin and LEF-1
Immunohistochemical studies suggested that the amount of -catenin in the nucleus increased after ATP depletion and that this increase persisted to some extent after recovery (Figure 2). To confirm these morphologic observations, we assessed the amount of immunodetectable -catenin, -catenin, and E-cadherin in nuclear and non-nuclear cell extracts by immunoblotting and then quantifying the data by densitometric analyses of six immunoblots. After ATP depletion, the nuclear content of -catenin increased by 40 ± 3% and the non-nuclear fraction decreased by 24 ± 2% (Figure 6). In contrast to the change in nuclear -catenin content, we detected no change in -catenin or E-cadherin nuclear content with ATP depletion and repletion (Figure 6). The lack of accumulation of these latter two proteins within the nucleus documents that the increment of -catenin observed with ATP depletion is specific and cannot be ascribed to methodologic errors.
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-catenin and LEF-1 are complexed to each other within the nucleus. Nuclear extracts were subjected to immunoprecipitation with anti–-catenin antibody. These immunoprecipitates contained barely detectable amounts of LEF-1 in control cells. However, after ATP depletion, substantial amounts of LEF-1 could be co-immunoprecipitated with antibody to -catenin (Figure 7). In contrast, LEF-1 was not co-immunoprecipitated from nuclear extracts with antibody to -catenin (Figure 7). These data suggest that ATP depletion induces the translocation of -catenin to the nucleus and that -catenin is complexed to LEF-1 within the nucleus. These observations are consistent with our hypothesis that the wnt/wingless signal transduction pathway is activated by ATP depletion.
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Discussion |
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We have recently shown that ATP depletion results in the withdrawal of E-cadherin from the basolateral membrane of tubular cells (31). Similar alterations in E-cadherin distribution of tubular cells after ischemia have been shown by other investigators (1,4,32). However, our group has also shown that some of the other proteins that comprise the ZA, such as the catenins, become tyrosine phosphorylated after ATP depletion and that these signaling events are important in the loss of ZA integrity and of tight junction function (31). Our findings are consistent with a role for tyrosine phosphorylation as a regulator of the formation and disaggregation of the ZA (33,34).
One purpose of this study was to characterize the effects of ATP depletion and repletion on the structural components of the ZA in greater detail (31). Immunohistochemical studies demonstrate that E-cadherin, -catenin, and -catenin are withdrawn from their normal position at the basolateral membrane after ATP depletion, changes that are largely reversible when ATP levels are allowed to increase toward control levels after ATP depletion (Figures 1 and 2). We next examined the effect of ATP depletion on the relative amounts of three ZA proteins (E-cadherin, -catenin, and -catenin) within Triton X-100–soluble and –insoluble fractions of cell lysate. The Triton X-100–insoluble fraction of the cell is predominantly in a cytoskeletal-associated pool, whereas the Triton X-100–soluble pool contains proteins largely unbound to the cytoskeleton (35). This technique has been used for many years by many different investigators to assess the effects of ischemia and other interventions on the distribution of a number of cell proteins between the cytoskeletal and noncytoskeletal compartments of the cell, including ezrin (36), Na+/K+-ATPase (35), as well as adhering and catenins (37,38).
We demonstrate that that the relative amount of all three ZA proteins studied increased by about 45% in the Triton X-100–insoluble fraction of the cell during ATP depletion (Figure 4). There was a comparable fall of these proteins in the Triton X-100–soluble fraction, so that the total amount of all three proteins did not change (Figure 4). These data suggest that all three ZA proteins studied are partially redistributed from a cytosolic to a cytoskeletal compartment of the cell. We also demonstrate that ATP depletion increased the relative amount of actin present in the Triton X-100–insoluble fraction. Because polymerized actin, F-actin, is Triton X-100–insoluble and G-actin is Triton X-100–soluble this observation is consistent with the conclusion that ATP depletion induces polymerization of G-actin and is consistent with earlier work of Molitoris et al (5,39). This group first documented that ATP depletion induces polymerization of monomeric G-actin to F-actin.
Although we found no evidence of degradation of either E-cadherin or the catenins, Bush et al. (1) have demonstrated that ATP depletion of MDCK cells and ischemia of kidney in vivo result in selective degradation of E-cadherin and partial disruption of complexes. The discrepancy between these two studies is probably due to differences in the duration of ATP depletion. We examined the effects of less than 70 min of ATP depletion, a period that induces reversible epithelial cell injury. We have found that ATP depletion lasting more than 90 min causes irreversible cell injury (3). In contrast, Bush et al. (1) examined the effects on ZA proteins of far longer duration of ATP depletion/ischemia, ranging from 2 to 6 h.
We also determined whether complexes of E-cadherin–catenin remain intact or dissociate during ATP depletion. We show that the total amount of -catenin co-immunoprecipitated with E-cadherin is unchanged with ATP depletion although the amount of these complexes in the Triton X-100–insoluble pool increased (Figure 5, B and C). These data suggest that most of the ZA proteins are withdrawn as intact complexes from the basolateral membrane and remain associated with the cytoskeleton. We speculate that maintenance of ZA proteins as intact complexes attached to the cytoskeleton facilitates the rapid recovery of the ZA that occurs during ATP repletion (Figure 2).
Interestingly, these changes in distribution of ZA proteins are different from those induced by the "calcium switch" model, which has been used extensively to study the effects of reversible loss of ZA integrity on the redistribution of cadherin-catenin complexes. In both calcium switch (removal of extracellular calcium) and ATP depletion models, intact E-cadherin–catenin complexes are removed from the plasma membrane. Although these complexes are associated with the cytoskeleton during ATP depletion, the calcium switch model has the opposite effect; the complexes are redistributed to the Triton X-100–soluble (cytosolic) fraction.
This difference in the distribution of ZA protein complexes in these two models may be related to the disparity in the effects that these interventions have on the actin cytoskeleton. The calcium switch model leads to depolymerization of F-actin to G-actin (1,40). By contrast, our data and those of other investigators (5,39), demonstrate that ATP depletion results in the opposite effect, an increased polymerization of G-actin to F-actin (Figure 4). We speculate that the increased amount of polymerized actin during ATP depletion provides additional actin binding sites that recruit additional ZA complexes from the Triton X-100–soluble pool. A similar mechanism has been proposed for the incorporation of Na+-K+ ATPase into actin aggregates as this transporter is withdrawn from the basolateral membrane during ischemia (41).
In addition to studying effects of ATP depletion on the redistribution of ZA proteins, this study had a second goal: to determine whether -catenin plays a signaling role during or after ischemia. It is well known that -catenin, in addition to its structural role as a part of the ZA, is an important component of the Wnt/wingless signaling pathway (11–14). Cytosolic -catenin can bind to the TCF/LEF-1 family of transcription factors. This complex then translocates to the nucleus and transactivates a number of genes (11–14). The Wnt/wingless pathway was first described in Drosophila, in which it is important in development. However, more recent evidence has shown that homologues of -catenin and TCF/LEF-1 can modulate the expression of a number of genes including c-myc (16), cyclin D (18), and E-cadherin (17) in mammalian cells. Thus the Wnt/wingless pathway appears to be involved in regulating cell cycling, apoptosis, and cell differentiation in mammals (11–14).
We maintain that if ATP depletion activates the signaling function of -catenin, the total amount of -catenin in the nucleus should increase after ATP depletion and should be complexed with LEF-1. We examined this hypothesis in two ways, by immunohistochemistry of cell monolayers and by immunoblotting of isolated nuclei for -catenin and LEF-1. Both methods clearly demonstrate an increase in the amount -catenin in the nucleus of tubular cells after ATP depletion (Figures 2 and 6). The nuclear accumulation of -catenin during ATP depletion is associated with a concomitant fall in the Triton X-100–soluble (cytosolic) pool of -catenin (Figure 6) and with an increment in nuclear LEF-1 (Figure 7). The nuclear LEF-1 can be co-immunoprecipitated with antibody to -catenin but not -catenin, documenting that -catenin and LEF-1 form a complex. Furthermore, after ATP repletion, the nuclear content of -catenin remains elevated (Figures 2) while the Triton X-100–soluble pool fraction of -catenin remains reduced (Figure 4). The source of the -catenin that translocates into the nucleus during ATP depletion remains uncertain. However, our data are consistent with previous reports that -catenin involved in wnt/wingless signaling is derived predominantly from a transcriptionally competent cytosolic pool (19).
Our novel findings of nuclear accumulation of -catenin and LEF-1 are consistent with our hypothesis that the wnt/wingless signaling pathway is activated during ATP depletion (11–14). Although we have not directly assessed the transcriptional effect of nuclear -catenin/LEF-1 complexes, this pathway under certain circumstances in mammalian cells, is associated with expression of genes such as cyclin D, c-myc, and E-cadherin (14). We propose that activation of this pathway by ATP depletion may contribute to some of the alterations in gene expression that occur within tubular cell after ischemic injury and that are necessary for renal regeneration and repair (20,21). Our observation that the increase of nuclear -catenin appears to persist after ATP-repletion is consistent with this idea (Figure 2). The significance of these observations, as a general model for the responses of renal epithelial cells awaits confirmation in other cultured cells lines and in in vivo studies.
In summary, we have shown that ATP depletion leads to disassembly of the ZA during ATP depletion, an event associated with the reversible withdrawal of intact E cadherin–catenin complexes from the basolateral membrane and an increase in the amount of these ZA protein complexes bound to the cytoskeleton. We also demonstrate for the first time that ATP depletion is associated with translocation of -catenin and LEF-1 to the nucleus. We suggest that the preservation of intact ZA protein complexes and the recruitment of additional ZA complexes to the actin cytoskeleton during ATP depletion facilitates the reestablishment of an intact ZA and tight junction during recovery by allowing the rapid delivery of intact complexes back to the basolateral membrane. We also hypothesize that -catenin participates in signaling events that contribute to alterations in gene expression associated with renal ischemia and that mediate some of the events necessary for regeneration and repair of tubular cells after injury (20,21).
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Acknowledgments |
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-catenin an d plakoglobin in renal tubular cells. J Am Soc Nephrol 10: 2297–2305, 1999This article has been cited by other articles:
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  A. Havasi, Z. Li, Z. Wang, J. L. Martin, V. Botla, K. Ruchalski, J. H. Schwartz, and S. C. Borkan Hsp27 Inhibits Bax Activation and Apoptosis via a Phosphatidylinositol 3-Kinase-dependent Mechanism J. Biol. Chem., May 2, 2008; 283(18): 12305 - 12313. [Abstract] [Full Text] [PDF]
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 M. Endemann, H. Bergmeister, B. Bidmon, M. Boehm, D. Csaicsich, L. Malaga-Dieguez, K. Arbeiter, H. Regele, K. Herkner, and C. Aufricht Evidence for HSP-mediated cytoskeletal stabilization in mesothelial cells during acute experimental peritoneal dialysis Am J Physiol Renal Physiol, January 1, 2007; 292(1): F47 - F56. [Abstract] [Full Text] [PDF]
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K. Ruchalski, H. Mao, Z. Li, Z. Wang, S. Gillers, Y. Wang, D. D. Mosser, V. Gabai, J. H. Schwartz, and S. C. Borkan Distinct hsp70 Domains Mediate Apoptosis-inducing Factor Release and Nuclear Accumulation J. Biol. Chem., March 24, 2006; 281(12): 7873 - 7880. [Abstract] [Full Text] [PDF]
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M. D. Covington, K. J. Bayless, R. C. Burghardt, G. E. Davis, and A. R. Parrish Ischemia-induced cleavage of cadherins in NRK cells: evidence for a role of metalloproteinases Am J Physiol Renal Physiol, August 1, 2005; 289(2): F280 - F288. [Abstract] [Full Text] [PDF]
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S. Zhuang, Y. Yan, J. Han, and R. G. Schnellmann p38 Kinase-mediated Transactivation of the Epidermal Growth Factor Receptor Is Required for Dedifferentiation of Renal Epithelial Cells after Oxidant Injury J. Biol. Chem., June 3, 2005; 280(22): 21036 - 21042. [Abstract] [Full Text] [PDF]
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 Y. T. Becker, R. M. Hofmann, J. Yracheta, A. Friedl, and B. N. Becker Epithelial cell polarity and improved early outcomes in delayed graft function: a pilot study of polyclonal vs monoclonal antibodies Nephrol. Dial. Transplant., November 1, 2004; 19(11): 2886 - 2892. [Abstract] [Full Text] [PDF]
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D. Sinha, S. Bannergee, J. H. Schwartz, W. Lieberthal, and J. S. Levine Inhibition of Ligand-independent ERK1/2 Activity in Kidney Proximal Tubular Cells Deprived of Soluble Survival Factors Up-regulates Akt and Prevents Apoptosis J. Biol. Chem., March 19, 2004; 279(12): 10962 - 10972. [Abstract] [Full Text] [PDF]
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 K. Ruchalski, H. Mao, S. K. Singh, Y. Wang, D. D. Mosser, F. Li, J. H. Schwartz, and S. C. Borkan HSP72 inhibits apoptosis-inducing factor release in ATP-depleted renal epithelial cells Am J Physiol Cell Physiol, December 1, 2003; 285(6): C1483 - C1493. [Abstract] [Full Text] [PDF]
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 J. V. Bonventre and J. M. Weinberg Recent Advances in the Pathophysiology of Ischemic Acute Renal Failure J. Am. Soc. Nephrol., August 1, 2003; 14(8): 2199 - 2210. [Full Text] [PDF]
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D. Sinha, Z. Wang, V. R. Price, J. H. Schwartz, and W. Lieberthal Chemical anoxia of tubular cells induces activation of c-Src and its translocation to the zonula adherens Am J Physiol Renal Physiol, March 1, 2003; 284(3): F488 - F497. [Abstract] [Full Text] [PDF]
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P < 0.05 versus CN + glucose.
