AJP - Renal Ad Instruments
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     

Am J Physiol Renal Physiol 285: F230-F240, 2003. First published April 22, 2003; doi:10.1152/ajprenal.00024.2003
0363-6127/03 $5.00
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
285/2/F230    most recent
00024.2003v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (5)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wald, F. A.
Right arrow Articles by Salas, P. J. I.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wald, F. A.
Right arrow Articles by Salas, P. J. I.

Membrane repolarization is delayed in proximal tubules after ischemia-reperfusion: possible role of microtubule-organizing centers

Flavia A. Wald, Yolanda Figueroa, Andrea S. Oriolo, and Pedro J. I. Salas

Department of Cell Biology and Anatomy, University of Miami School of Medicine, Miami, Florida 33136

Submitted 17 January 2003 ; accepted in final form 17 April 2003


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
We have previously shown that microtubule-organizing centers (MTOCs) attach to the apical network of intermediate filaments (IFs) in epithelial cells in culture and in epithelia in vivo. Because that attachment is important for the architecture of microtubules (MTs) in epithelia, we analyzed whether chemical anoxia in LLC-PK1 and CACO-2 cells or unilateral ischemia-reperfusion in rat kidney (performed under fluorane anesthesia) had an effect on the binding and distribution of MTOCs. In culture, we found that chemical anoxia induces MTOC detachment from IFs by morphological and biochemical criteria. In reperfused rat proximal tubules, noncentrosomal MTOCs were fully detached from the cytoskeleton and scattered throughout the cytoplasm at 3 days after reperfusion, when brush borders were mostly reassembled. At that time, MTs were also fully reassembled but, as expected, lacked their normal apicobasal orientation. Two apical membrane markers expressed in S2 and S3 segments were depolarized at the same stage. At 8 days after reperfusion, membrane polarity, MTOCs, and MTs were back to normal. Na+-K+-ATPase was also found redistributed, not to the apical domain but rather to an intracellular compartment, as described by others (Alejandro VS, Nelson W, Huie P, Sibley RK, Dafoe D, Kuo P, Scandling JD Jr., and Myers BD. Kidney Int 48: 1308–1315, 1995). The prolonged depolarization of the apical membrane may have implications in the pathophysiology of acute renal failure.

ischemic acute renal failure; apical polarity; intermediate filaments

SEVERAL PROCESSES INVOLVED in cellular injury during ischemia and after ischemia-reperfusion (I/R) have been reported in recent years in an effort to understand the pathophysiology of ischemic acute renal failure (ARF) (7, 17). It is generally accepted that disorganization of the cytoskeleton and loss of membrane polarity are significant pathophysiological mechanisms of ischemic injury in proximal tubule cells (44). Most attention has been focused on the disarrangement of the actin cytoskeleton (20, 26), although it is known that cortical fodrin, villin, and tight junctions are also affected (9, 16, 27). Interestingly, however, it has been recently reported that disorganization of the actin cytoskeleton alone is not sufficient to explain the anoxic disruption of the plasma membrane (13). In addition, Brown and co-workers (1) also showed depolymerization of microtubules (MTs) in proximal tubules in vivo during the first 24 h after I/R.

Our laboratory and others have reported that microtubule organizing centers (MTOCs) are apical in simple epithelia (3, 4, 10, 25, 34, 37), thus organizing the MTs in a polarized fashion with the minus ends toward the apical membrane (5). Although this apicobasal polarization of MTs is not absolutely essential for polarization (19, 38), it is thought to participate in the vesicular traffic bound to the apical membrane (24), especially for vesicles involved in transcytosis from basolateral to apical membrane (8). In this regard, the nucleating activity of MTOCs must be necessary to repolymerize MTs after I/R. Because MTOCs cap the minus ends of MTs (45), it is expected that MTOCs positioned in their normal (apical) localization will reorganize MTs with the correct orientation (minus end apical). Conversely, if MTOCs become delocalized during ischemia, the MTs formed during the recovery period will have abnormal orientations. In that scenario, aberrant MTs may transport the carrier vesicles that originate in the trans-Golgi network and contain apical cargo (28) to incorrect regions of the cytoplasm. Therefore, newly formed MTs with an incorrect orientation may contribute to the mispolarization of the plasma membrane more seriously than a simple depolymerization of MT.

Because our previous work suggests that the apical localization of MTOCs in simple epithelia depends on binding to apical intermediate filament (IF) (37), we decided to analyze whether the IFs are also disrupted by anoxia and whether the MTOCs are still bound to IF and apically localized in proximal tubule kidney cells after ischemia. We performed the analysis in two epithelial cell lines, and in vivo, using a mild I/R protocol of unilateral clamp of the renal vessels, which is known to disrupt the MTs (1) but causes little or no necrosis and only modest amounts of apoptosis only within the first 24 h after ischemia (31). The results indicate that MTOCs become delocalized after chemical anoxia in tissue culture or as a result of I/R in vivo. Furthermore, the anomalous localization of MTOCs results in a disorganized array of MTs and correlates with poor membrane polarization of proximal tubule cells, delayed by days respect to the formation of the F-actin brush border.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Cell culture and ATP depletion. LLC-PK1 (porcine proximal tubule) and CACO-2 (human colon carcinoma epithelial cells) were obtained from ATCC and maintained by weekly passages in tissue culture plastic flasks in DMEM-F-12 nutrient mixture (DMEM/F-12, GIBCO) supplemented with 10% fetal bovine serum (Cellgro). For chemical anoxia experiments, the method described by Molitoris and co-workers (11) was used, with the following modifications. Monolayers confluent for 4 (LLC-PK1) or 9 days (CACO-2) were incubated in Earle's balanced salt solution (GIBCO formulation) without glucose or other nutrients for 30 min and then changed to the same solution supplemented with 0.5 µM antimycin A (Sigma), 1 mM adenosine, and 0.2 mM allopurinol (15) for 1 h (ATP depletion). Recovery was initiated by four washes and incubation in the standard DMEM/F-12 culture medium for various times. ATP determinations were performed using a luciferase-based kit (Calbiochem).

Kidney ischemia in vivo. Animal handling was in compliance with Public Health Service Policy on Humane Care and Use of Laboratory Animals. Male Sprague-Dawley rats (Charles River Laboratories) weighing 300–450 g were anesthetized with 1.5% isoflurane and kept on a warm plate to maintain temperature. The peritoneal cavity was opened, and the left renal vessels were clamped for 30 min with two microvascular (no. 1) clamps. The success of the clamp, and, later, the reperfusion, was visualized by rapid changes in the color of the kidney. The animals were sutured and kept under an infrared lamp until fully awake. After various periods of time, the animals were killed by an overdose of pentobarbital sodium (1 mg/10 g body wt) and intracardiac perfusion of formaldehyde fixative (in the case of immunofluorescence experiments). The success of the perfusion was assessed by the rigid posture of muscles and tail.

Antibodies and fluorescent reagents. The antibodies used in this study were as follows: polyclonal antibody (Ab) anti-{gamma}-tubulin NH2-terminal synthetic peptide (EEFATEGTDRKDVFFY, Sigma); monoclonal antibody against {alpha}-tubulin (DM 1A; Sigma); monoclonal antibody against keratin (K) 19 (in CACO-2 cells; RCK-108; Accurate Chemical); monoclonal antibody against K19 (in LLC-PK1 cells; A53/BA2; Sigma); monoclonal antibody against K18, B23.1 (Biomeda); polyclonal antibody anti-intestinal alkaline phosphatase (iAP; DAKO); polyclonal antibody anti-carbonic anhydrase IV (CAIV), a generous gift from Dr. A. Waheed (St. Louis University School of Medicine); and polyclonal antibody anti-Na+-K+-ATPase, a generous gift from Dr. W. J. Nelson (Stanford Univ.). Secondary fluorescent antibodies were affinity purified (Jackson Laboratories). FITC-phalloidin was purchased from Molecular Probes.

Immunofluorescence and confocal microscopy. Immunofluorescence and confocal microscopy of tissue culture cells were performed as described before (37). Rat kidneys were fixed by intracardiac perfusion of the whole animal with warm 3% formaldehyde, 0.1% glutaraldehyde. The kidneys were rapidly extracted from the animal, sectioned with a razor blade into 1-mm-thick sections, and further fixed in the same fixative for 2 h. Then, the sections were infused overnight in PBS supplemented with 27% (wt/vol) sucrose, 0.3% formaldehyde, and 0.01% glutaraldehyde. The kidneys were frozen in the same solution in isopentane at melting point and stored at –70°C. Sectioning was performed in a cryostat at –25°C. The sections, attached to a glass slide, were thawed by immersion in the same fixative described above and processed for immunofluorescence following the standard protocol (37). For quantification of signal in confocal sections, highly confocal images, collected at 0.7 Airy units, were acquired, with care taken that no pixels in the positive signal were saturated. Then, the images were analyzed by selecting separately in each cell section the basolateral domain and the corresponding apical domain. In each selected area, we calculated the product of number of pixels times average pixel values (luminosity channel) and the basolateral-to-apical ratio for each cell. The data are presented as means ± SD of those ratios. Apoptosis in kidney sections was evaluated using a DNA fragmentation detection kit (Oncogene Research Products), based on the end labeling of fragmented DNA, with biotin-labeled deoxynucleotides by Klenow DNA polymerase I.

Cell extraction, fractionation, and immunoprecipitation of sonication fragments. Cell extraction, fractionation, and immunoprecipitation of sonication fragments were done as previously described (34, 37). Briefly, confluent monolayers of LLC-PK1 cells were grown in one roller bottle. The cells were extracted in 1% Triton X-100, in PBS supplemented with 2 mM EDTA, and a complete cocktail of antiproteases (Sigma), and the pellet was extensively sonicated (3 min total, with 10-s periods of sonication and 15-s lapses to dissipate heat). The sonication fragments of the Triton-insoluble cytoskeleton were separated in 10-ml 20–60% sucrose continuous gradients in a SW41 swinging bucket rotor at 15,000 rpm for 50 min at 4°C. Usually, the top five 1-ml fractions of the gradient were used. Immunoprecipitation was done following standard procedures. The protein A-agarose beads were centrifuged through a 0.5-ml 30% sucrose cushion. All these centrifugations were done at 14,000 rpm for 2 s to minimize nonspecific copelleting of unbound cytoskeletal fragments. Immunoblotting was performed as described before (37).


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Relocalization of centrosomes during chemically induced ATP depletion and ischemia. To study the effects of chemical anoxia in tissue culture cells, we used a well-established protocol that is known to cause a rapid and reproducible depletion of ATP (11). The efficiency of chemically induced ATP depletion in our cells and the kinetics of recovery after 1-h depletion were assessed in LLC-PK1 (porcine proximal tubule) and CACO-2 (human colon carcinoma) cells using a standard luciferase assay. In both cell lines, ATP levels fell >90% within the first 10 min after treatment with antimycin A, and significant recoveries of ATP levels were started between 2 and 4 h after the cells were replaced in DMEM (Fig. 1). Thus the kinetics of ATP depletion/recovery for LLC-PK1 were similar to those in previous reports.



View larger version (23K):
[in this window]
[in a new window]
  		Fig. 1. ATP levels in CACO-2 (A) and LLC-PK1 (B) epithelial cells in tissue culture during and after chemically induced ATP depletion. The ATP from quadruplicate confluent monolayers (~3 x 105 cells each) was extracted at various times of incubation in nutrient-free medium supplemented with antimycin A, adenosine, and allopurinol (depletion) and after a 1-h depletion at various times of recovery in normal culture medium. ATP was measured by luciferase luminescence by integrating light emission for 1 min.

 

To analyze the localization of centrosomes and to extend the observations to a different cell line, LLC-PK1 and CACO-2 monolayers were subjected to ATP depletion for 1 h and 1-h recovery. Double-immunofluorescence experiments were performed colocalizing the {gamma}-tubulin signal (Fig. 2, green) with the cortical IF cytoskeleton (Fig. 2, red). To show the apical domain alone, K19 was used as a marker of IF in the CACO-2 cells. As in CACO-2 cells (37), K18, the other type I keratin, distributed under the apical and lateral domains in LLC-PK1 cells, but the IF cortical signal was thicker in the kidney cells than in the intestinal cells (Fig. 2, a, c, e, g, and i). More importantly, ATP depletion did not affect the IF cytoskeleton in either LLC-PK1 or CACO-2 cells (Fig. 2). A similar result was observed, reciprocally, using anti-K18 or anti-K19 antibodies in CACO-2 or LLC-PK1 cells, respectively (not shown).



View larger version (37K):
[in this window]
[in a new window]
  		Fig. 2. Centrosomes detach from their normal apical location at the apical intermediate filament (IF) network after 1-h ATP depletion in LLC-PK1 and CACO-2 cells. Monolayers of LLC-PK1 (a, c, e, g, i) and CACO-2 cells (b, d, f, h, j) were grown on Transwell filters. Some cells were subjected to ATP depletion for 1 h (e–j). The cells were fixed after 1-h recovery and processed for double immunofluorescence with monoclonal Abs against K18 (LLC-PK1 cells) or K19 (CACO-2 cells) (red channel) and a polyclonal antibody against a conserved NH2-terminal polypeptide of {gamma}-tubulin (green channel). Two examples of each control and 3 examples of ATP-depleted cells were analyzed by confocal microscopy, deconvolution, and 3D reconstruction and are shown in the XZ plane (perpendicular to the monolayer), with the apical side up. Arrows point at {gamma}-tubulin signal separated from the apical domain IFs. Scale bars: 5 µm(a–i) and 2 µm(j).

 

In Fig. 2, examples of fields with centrosomes within the same confocal optical section in nearby cells are shown for control monolayers (a–d) and ATP-depleted cells (e–j). As expected from previous publications, the centrosomes in control cells were observed always embedded in the apical IF submembrane cytoskeleton (Fig. 2, a–d). In ATP-depleted cells, a proportion of the centrosomes was found separated from the apical IF cytoskeleton (Fig. 2, arrows). The proportion was smaller in LLC-PK1 cells (~28%) than in CACO-2 cells (~49%) but in both cases significantly larger than the proportion of centrosomes normally found separated from the IF in control interphasic cells (0% in all our samples). In control cells, a modest proportion of centrosomes (usually <10%) does appear attached to the lateral domain IF (37), which was not the case in the ATP-depleted cells either. In some cases, the {gamma}-tubulin signal usually interpreted as centrosomes (~0.4-µm spheres, often observed in pairs) appeared fragmented in ATP-depleted cells (Fig. 2, h and i). The noncentrosomal {gamma}-tubulin signal (smaller dots or diffuse signal), normally at or within 1 µm of the terminal web (25, 37), also appeared separated from the apical submembrane cytoskeleton, especially in LLC-PK1 cells (Fig. 2e).

Because colocalization does not necessarily imply attachment, the physical connection of insoluble {gamma}-tubulin-containing structures and IF was analyzed as before (37) by fragmenting the Triton-insoluble cytoskeleton of LLC-PK1 cells with extensive sonication and separating the smallest fragments by size in sucrose gradients by rate centrifugation. This procedure was originally devised to perform immunoprecipitation in highly insoluble multiprotein complexes (34). We collected the top five fractions of the gradient that contain fragments small enough to be immunoprecipitated (larger fragments tend to nonspecifically copellet with agarose beads). Each aliquot was divided into two equal parts, one of them immunoprecipitated with an irrelevant rabbit IgG as a negative control (Fig. 3, – lanes), and the other with anti-{gamma}-tubulin Ab (Fig. 3, + lanes). The immunoprecipitates were analyzed by immunoblot using anti-K18 or anti-K19 monoclonal antibody. In general, the previous findings in CACO-2 (37) were extended to LLC-PK1 cells. In control cells, there was coimmunoprecipitation of K18 and {gamma}-tubulin in fractions 1, 3, and 4 and of K19 and {gamma}-tubulin in fractions 1–5 (Fig. 3). This coimmunoprecipitation was mostly abolished in LLC-PK1 cells subjected to 1-h ATP depletion. K18 blots showed only background levels, and K19 showed only some ATP depletion-insensitive coimmunoprecipitation with {gamma}-tubulin in fraction 4 (Fig. 3). Controls showing that this coimmunoprecipitation cannot be explained by simple physical trapping or artifactual protein binding as a result of detergent extraction have been performed elsewhere (37, see Fig. 8). These results indicate that the attachment between MTOC and IFs is largely broken after 1-h ATP depletion in LLC-PK1 and CACO-2 cells in tissue culture.



View larger version (40K):
[in this window]
[in a new window]
  		Fig. 3. ATP depletion inhibits coimmunoprecipitation of insoluble {gamma}-tubulin with K18 and 19 in LLC-PK1 cells. LLC-PK1 cell monolayers were grown in 2 roller bottles (~1.5 x 108 cells each). One of the cultures was ATP depleted for 1 h. The cells were extracted in Triton X-100, and the insoluble cytoskeletal preparations were extensively sonicated. The fragments were separated by size by rate centrifugation in sucrose gradients and fractions 1–5 (top half of the gradient) divided in 2 equal aliquots each. The aliquots were immunoprecipitated with anti-{gamma}-tubulin antibody (+ lanes) or an irrelevant rabbit IgG (– lanes). The immunoprecipitates were analyzed by immunoblot using monoclonal Abs against K18 and K19 sequentially on the same blot. Left: molecular mass standards (in kDa).

 


View larger version (49K):
[in this window]
[in a new window]
  		Fig. 8. Intracellular localization of "depolarized" Na+-K+-ATPase after I/R in proximal tubules. Sections of the right (sham; 3 days after operation) and left (I/R) kidneys fixed 3 and 5 days after renal artery clamp operation were processed for indirect immunofluorescence with an anti-Na+-K+-ATPase antibody (red channel) and FITC-phalloidin (green). The arrows point to cells with intracellular distribution of Na+-K+-ATPase. Bars: 10 µm.

 

Ischemic injury in vivo causes detachment of MTOCs in proximal tubules. To test whether MTOCs also detach from the apical domain in vivo after ischemia, rat kidneys were subjected to a 30-min clamp of the renal vessels. The contralateral kidneys were used as a sham operation control. We analyzed the kidneys at 24 h after I/R, a time that has been widely studied but that yields results difficult to analyze because of the complexity of the effects. We also analyzed the kidneys at 3, 5, 7, and at 8 days after I/R. The latter was chosen because our preliminary experiments showed that the changes described in this section last up to 7 days after I/R, and kidney functional parameters with a 30-min bilateral clamp protocol have been reported to be nearly normal as early as 5 days after I/R (23). In addition, Hropot and co-workers (21) found plasma creatinine and sodium still abnormal after 7 days for a 40-min bilateral clamp model. Even in a different system, the human renal allograft, patients recovering well achieve acceptable glomerular filtration rates on day 7 (14).

Frozen sections from reperfused or sham-operated kidneys were analyzed by immunofluorescence with anti-{gamma}-tubulin antibody and FITC-phalloidin to stain F-actin. In the control proximal tubules, very few 0.3- to 0.4-µm spots of {gamma}-tubulin signal (centrosomes) were easily recognizable and were mostly localized in the apical pole of the cells. However, a continuous subapical layer of {gamma}-tubulin signal was observed in most tubules both in kidneys subjected to a sham operation (Fig. 4a) and in kidneys from animals not subjected to surgery (not shown). When analyzed in sections perfectly perpendicular to the axis of the tubule, the {gamma}-tubulin signal (Fig. 4a, inset, green) could be localized immediately below the F-actin signal in the brush border (Fig. 4a, inset, red; arrows point at basal actin signal). This image is consistent with findings in our laboratory and by others as well that a layer of noncentrosomal MTOCs lies under the apical domain (3, 25). During the first 24 h after I/R, the apical {gamma}-tubulin layer became very discontinuous or disappeared altogether (not shown). However, because the entire apical domain is degraded (44), it is difficult to draw any conclusions about the fate of MTOCs at that time. More interesting were the results from kidneys at 3 days following I/R. At this stage, the F-actin component of the brush border was mostly repaired in many (but not all) cells, as judged by a phalloidin label (Fig. 4d, arrows). However, no subapical layer of {gamma}-tubulin was observed, even in tubules (or cells) with a complete brush border (Fig. 4, c vs. d, arrows). Instead, {gamma}-tubulin signal was either diffuse in the cytoplasm or concentrated under the lateral domain in some cells (Fig. 4c, arrows). At 8 days after I/R, tubules subjected to a 30-min ischemic injury had recovered the normal distribution of {gamma}-tubulin and the images became indistinguishable from controls in all cells (like in Fig. 4a). This result suggests that the repolarization of MTOCs is delayed with respect to the reassembly of the F-actin brush border.



View larger version (53K):
[in this window]
[in a new window]
  		Fig. 4. Polarization of microtubule-organizing centers (MTOCs) is delayed respect to the formation of the brush border in proximal tubules. Frozen sections from the kidneys of an animal that was subjected to a 30-min clamp of the left renal vessels followed by a 3-day recovery (c and d) and sham-operated kidney (sections from the right kidney; a and b) were processed with fluorescent phalloidin (F-actin; b and d) or for indirect immunofluorescence with anti-{gamma}-tubulin antibody (a and c). Inset: higher magnification view of a segment of a proximal tubule overlapping pseudocolor images of phalloidin (red) and anti-{gamma}-tubulin (green) signals. L, lumen; arrowheads, basal actin signal. Arrows point out depolarized {gamma}-tubulin signal (c) and fully reassembled F-actin brush border (d). Scale bars: 20 µm (a–d) and 5 µm (inset).

 

The same experiment was repeated in several animals, i.e., the harvesting of kidneys at 1, 3, and 8 days after I/R. To assess the attachment of MTOCs to the cytoskeleton, pieces of the cortex, freshly obtained from the animal, were homogenized and centrifuged at low speed, so that the soluble 27S {gamma}-tubulin complexes ({gamma}-TurC; 42) remain in the supernatant and {gamma}-tubulin attached to the cytoskeleton in the pellet. In previous studies, roughly 50% of the {gamma}-tubulin was found soluble (37, 42), and that was the case for kidney cortexes subjected to a sham operation (S in Fig. 5, pellets vs. soluble). Despite the disruption in the architecture of the tissue, and with some variability, 37% of the {gamma}-tubulin (with respect to control) was still associated with the cytoskeletal pellet after 1 day of recovery (I in Fig. 5 and Table 1). This result can be due to minimal amounts of MTOCs still attached to the apical cytoskeleton, which may go undetected by regular morphology. Interestingly, however, no {gamma}-tubulin was found associated with the pellet fraction at 3 days after I/R (Fig. 5, Table 1). At 8 days after I/R, the fraction of {gamma}-tubulin in the pellet resembled again that of the sham-operated kidneys (Fig. 5, Table 1). Together with the data in Fig. 4, this result suggests that the bonding of MTOCs to the cytoskeleton breaks down after I/R and that it takes days to be reestablished. This is a much longer time period than it takes to reassemble apical F-actin at the brush border.



View larger version (40K):
[in this window]
[in a new window]
  		Fig. 5. MTOCs are detached from the cytoskeleton in the renal cortex 3 days after an ischemic injury. The left kidneys of rats were subjected to a 30-min clamp followed by 1, 3, or 8 days of reperfusion (I). The corresponding right kidneys were used as sham controls (S). Each S and I pair corresponds to a different animal. Pieces of cortex were homogenized in nonionic detergent, and the cytoskeletal (detergent insoluble) fraction was pelleted at low-speed centrifugation, which was devised to maintain {gamma}-tubulin (tub.) complexes ({gamma}-TurC, 42) in the supernatant. The pellets were rehomogenized by sonication. Then, 30 µg total protein of each sample were separated by SDS-PAGE and analyzed by immunoblot with an anti-{gamma}-tubulin antibody. The same membranes were reprobed with an anti-keratin antibody (pellets) or an anti-{alpha}-tubulin antibody (supernatants) to independently assess protein load in the same lanes.

 

View this table:
[in this window]
[in a new window]
  		Table 1. {gamma}-Tubulin signal in the cortex of kidneys subjected to I/R as a percentage of the band from the contralateral (sham surgery) kidney

 

Disorganization of MTs at 3 days after I/R in proximal tubules. It is known that MTs disassemble within the first 24 h after I/R (1). We confirmed that observation in kidney sections stained with an anti-{alpha}-tubulin antibody. The few MTs remaining at 1 day after I/R, however, were still oriented in the apicobasal axis (not shown), a result consistent with the persistence of 37% of the MTOCs still attached to the cytoskeleton (Fig. 5, Table 1). However, because of the delocalization of MTOCs described above at 3 days after I/R, we were prompted to analyze their organization at that particular time. Observation of the sections with regular epifluorescence microscopy revealed that the density of MTs per cell was similar to that in sham-operated kidneys (shown only in confocal microscopy, Fig. 6, a vs. b), suggesting that an active process of repolymerization of tubulin occurs between 1 and 3 days after I/R. The thickness of the sections (~5 µm), compared with the diameter of single MTs (~20 nm), however, made it necessary to visualize MT bundles under confocal microscopy using a high degree of confocality (0.7 Airy units), which gives high resolution in the z-axis. Proximal tubules from kidneys subjected to a sham operation observed under those conditions showed MT bundles mostly oriented in the apicobasal axis (Fig. 6a), as described extensively before (3, 5, 37). In the proximal tubules at 3 days after I/R, on the other hand, different arrangements were observed. In some cases, MTs were visualized as a disorganized network (Fig. 6b, red channel, arrows), mostly separated from the apical F-actin layer (green channel). In other sections, MTs adopted a basolateral distribution, and many bundles were roughly perpendicular to the apicobasal axis (Fig. 6c, arrows), an orientation seldom seen in control cells.



View larger version (47K):
[in this window]
[in a new window]
  		Fig. 6. Microtubules remain disorganized 3 days after ischemia-reperfusion (I/R) in tubules with normal F-actin distribution. Sections of the right (sham; a) and left (I/R; b and c) kidneys fixed 3 days after renal artery clamp operation were stained with an anti-{alpha}-tubulin monoclonal Ab (red channel) and subsaturation concentrations of FITC-phalloidin to minimally counterstain the brush-border F-actin (green channel). The sections were analyzed by confocal microscopy at 0.7 Airy units. Arrows, abnormal microtubule distribution. Scale bars: 10 µm.

 

Lack of polarity of apical plasma membrane markers at 3 days after I/R in proximal tubules showing an apparently normal F-actin brush border. Because MTOCs cap the minus end of MTs (45) and because MTOCs seem to be scattered in the cytoplasm at 3 days after I/R (Fig. 4) free from their normal cytoskeletal attachment (Fig. 5), we reasoned that the disorganized MTs at 3 days after I/R (Fig. 6) must have their minus ends randomly distributed in the cells. Therefore, MTs would not only not contribute to the normal polarization of the plasma membrane but would actually tend to randomize apical membrane proteins. To test this hypothesis, we analyzed the polarization of two apical membrane markers normally expressed in S2 and S3 segments of the proximal tubule, CAIV (S2 and S3 marker) (40) and iAP (S3 marker) (30). At 3 days after I/R, F-actin in the brush borders was similar to control tubules, as judged by phalloidin distribution (Fig. 7, d vs. b). CAIV and iAP were observed only in the brush border in all tubules where they are expressed in sham-operated kidneys (Fig. 7a, Table 2). At 1 day after I/R, the iAP signal could not be observed and the CAIV signal was weak and depolarized in at least one cell in 72% of the S2–3 sections (Table 2). This result is consistent with previous publications describing the loss of apical membrane and depolarization of the cells immediately after ischemia (2, 44). Strikingly, however, on day 3 significant levels of basolateral CAIV (Fig. 7c, arrows) were observed in one or more cells in 55% of the S3 sections (Table 2). Similarly, 64% of the sections positive for iAP showed at least one cell depolarized (Table 2). The number of depolarized cells decreased significantly on day 5, although isolated depolarized cells were observed up to 7 days after I/R for CAIV (Table 2).



View larger version (101K):
[in this window]
[in a new window]
  		Fig. 7. Substantial amounts of basolateral carbonic anhydrase IV (CAIV) signal remain 3 days after I/R in tubules with a reestablished F-actin brush border. Sections of the right (sham) and left (I/R) kidneys fixed 3 days after renal artery clamp operation were processed for indirect immunofluorescence with an anti-CAIV antibody (a, c, e, g) followed by a specific secondary antibody coupled to CY3 and FITC-phalloidin to show F-actin (b, d, f, h). e–h: Confocal images of the same preparations at higher magnification. Scale bars: 20 µm (a–d) and 10 µm (e–h).

 

View this table:
[in this window]
[in a new window]
  		Table 2. Delayed recovery of epithelial polarization in S3

 

To obtain a more precise assessment of the magnitude of the depolarization of CAIV and iAP, we measured the total signal in the apical and basolateral domain in high z-axis resolution confocal optical sections (Fig. 7, e and g). For each cell, the signal was weighed as total pixels x average pixel intensity in each area. With care taken to avoid images in which high-intensity pixels are saturated, this method can provide a fair estimation of the relative distribution of a membrane marker in a way that accounts for the differences in membrane folding (36). In proximal tubules subjected to sham surgery, only 3.6 ± 4.2% of total CAIV signal was localized to the basolateral domain (Fig. 7, a and e; only detectable with digital microscopy). In kidneys at 3 days after I/R, on the other hand, proximal tubule cells showed 48.2 ± 13.3% of total cellular CAIV signal localized to the basolateral domain (Fig. 7, c, arrows, and g). In other words, these cells "look" partially polarized because the apical signal is localized in a smaller area of the section with a high level of membrane folding (brush-border microvilli), whereas the basolateral signal is in a much more stretched membrane array. However, nearly equal amounts of the apical membrane marker are localized in each domain, and, thus these cells are almost totally depolarized.

Intracellular localization of Na+-K+-ATPase after I/R. To analyze the polarity of basolateral membrane proteins during the same period after I/R, we conducted experiments similar to those depicted in Fig. 7 but using an anti-Na+-K+-ATPase antibody instead. The antibody showed a basolateral image in sham kidneys. In 3-day I/R kidneys, a large proportion (72% of the proximal tubules, an average of 8 cells/section, Table 2) of the proximal tubule cells showed intracellular images, even in cells with an already fully reassembled actin brush border (Fig. 8, arrows). At this stage, however, little or no apical distribution of Na+-K+-ATPase was observed. At 5 days after I/R, only scattered cells (21% of the sections, an average of 1.2 cells/section) were still showing the intracellular distribution of Na+-K+-ATPase, and with few exceptions (one shown in Fig. 8C), no Na+-K+-ATPase signal was found in the apical domain either (Fig. 8). These results suggest that Na+-K+-ATPase also redistributes at the 3-day stage after I/R but not in the apicobasal axis but rather into an intracellular compartment and fully confirm the observations of Alejandro and co-workers (2) in human allografts. In addition, the results also indicate that different mechanisms govern the polarization of apical and basolateral markers, as described before in Madin-Darby canine kidney cells (43) and that reassembly of the brush-border actin is not an indicator of full polarization.


   DISCUSSION
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
The results in this work point at three general conclusions. First, IFs are stable after ATP depletion in culture or I/R in vivo (Figs. 2 and 5). Second, I/R leads to a separation of MTOCs from the cytoskeleton, in culture (Fig. 2) and in vivo (Figs. 4 and 5). This detachment results in a nonpolarized distribution of MTOCs and a disorganized architecture of MTs that last up to 1 wk after a mild ischemic injury, well beyond the time when the microfilaments are fully reorganized (2–3 days after I/R) (Fig. 6). Third, at least some plasma membrane proteins remain depolarized after the reorganization of the brush border, as defined by F-actin, and repolarize roughly with the same kinetics as MTOCs and MTs reacquire their normal subcellular organization (Fig. 7).

The initial experiments in this work were performed in two epithelial cell lines in tissue culture: LLC-PK1, derived from the proximal tubule, and CACO-2, derived from the intestine. The results from both cell lines indicate that ATP depletion has a significant impact on the apical localization of MTOCs. In proximal tubules, centrosomes were difficult to identify, and perhaps they are absent altogether in most cells. Therefore, we rather focused on the layer of noncentrosomal MTOCs that has been visualized in vivo in other epithelia as well (3). The dramatic fate of the apical pole in vivo as a result of the I/R injury, extensively described by Molitoris and co-workers (11), made any comparison of the redistribution of MTOCs in culture vs. in vivo nearly impossible. In addition, the analysis of the initial 24 h after I/R is complicated by the presence of apoptotic cells (31). Biochemical and morphological data in this work (Figs. 4 and 5) indicate that between 24 and 72 h after I/R, nearly all MTOCs become detached from the cytoskeleton and are scattered throughout the cytoplasm. Importantly, the data in tissue culture cells (Fig. 2) and in vivo (Fig. 5) consistently indicate that the detachment of MTOCs is not a consequence of depolymerization of IFs, the normal anchor of MTOCs (18, 37), that in all cases remained unaffected by ATP depletion or I/R (Figs. 2 and 5, respectively). The immediate consequence of the detachment of MTOCs was that MTs, although fully repolymerized during the same period of time (1–3 days after I/R), became a disorganized network, instead of the normal array of bundles highly oriented in the apicobasal axis (Fig. 6). Although we cannot assert a cause-and-effect relationship, it is safe to conclude that the disorganization of MTs correlates with poor levels of plasma membrane polarity, which only revert over a period of days to nearly normal at 5 days after I/R and, in some isolated depolarized cells, up to 7 days after I/R (Fig. 7, Table 2).

There are a number of possible explanations for the delayed recovery of distribution of MTOCs at 3 days after I/R. The first is that apoptosis, could, in principle, account for the separation of MTOCs from the cytoskeleton by a simple action of caspases. However, proximal tubule cells at 3 days after I/R displayed a nearly normal F-actin cytoskeleton and full-length MTs (although in abnormal orientations) (Figs. 6b and 7d). Furthermore, and in agreement with previous publications using the same I/R protocol (31), we found only modest proportions of apoptotic cells (5–8%) during the first 24 h after I/R, and none at 3 days (not shown). This was the reason the protocol was chosen in the first place, as opposed to more drastic ischemic injuries (e.g., 1 h) that result in higher percentages of apoptotic cells, and, eventually, a late secondary peak of apoptosis (41). In addition, we have demonstrated that, at least in intestinal epithelia, apoptosis does not disrupt epithelial polarity as long as the cell maintains the integrity of its plasma membrane (3). On the other hand, we cannot rule out that the early detachment of MTOCs from IF found in tissue culture cells may be due to caspase activation. Second, dedifferentiation, in a general sense, may occur because tens of genes are either upregulated or downregulated using the same I/R protocol at 1 or 3 days after I/R (48). More specifically, the expression of vimentin has been implicated as a reporter of dedifferentiation in proximal tubule cells after ischemia, and, along with the expression of proliferating cell nuclear antigen (PCNA), it was found in a substantial number of S3 cells at 2–5 days postischemia using a harsher ischemic injury (46). In our system, vimentin was expressed in <2% of the cells at 3 days after I/R. Interestingly, some of the vimentin-expressing cells actually had a polarized distribution of CAIV (not shown). Therefore, while it is quite likely that changes in the pattern of gene expression may be responsible for the detachment of MTOCs from the cytoskeleton, the phenomenon cannot be assigned to a simple epithelial-to-mesenchymal transition. Third, changes in the intracellular signaling/regulatory pathways, on the other hand, are more likely to be responsible for a phenomenon that occurs >24 h after I/R. Although the molecular identity of the "glue" attaching MTOCs to IFs has not yet been established, we have recently identified an ~190-kDa protein that, on phosphorylation with p34cdc2, seems to mediate the detachment of MTOCs from IF (18). Although we have no evidence that the same protein is responsible for the detachment of MTOCs from IF at day 3 after I/R, our working hypothesis is that a posttranslational modification of one of the components of the glue may be implicated in this phenomenon. For this reason, our laboratory is presently engaged in an attempt to identify the molecular components that mediate the binding of MTOCs to IFs. After I/R, specific signaling pathways become activated (39). The activation of "stress" kinases (p39 MAP, c-Jun, and ATF3), however, seems to be an early event after I/R (3 h) (47), and the exact nature of all the signaling/regulatory pathways active at 3 days after I/R is still unknown. Fourth, mitotic responses have been observed in S3 with harsher ischemic injuries, i.e., 40- (46) and 50-min ischemia (29). PDGF receptors have been implicated in this response (29) as well as stress kinases (39). In mitotically active tissue culture cells, MTOCs cycle from an IF-attached state in interphase to a free and mobile state during mitosis (18, 33). While there is no doubt that MTOCs are apical in quiescent cells, it is still conceivable that they may not reattach to the apical cytoskeleton in mitotically active S3 cells. We cannot rule out this scenario, either.

One lesson that we can draw from various studies of the acquisition of membrane polarity in tissue culture cells is that the time course of polarization of various components may be different and sometimes counter-intuitive. For example, we have demonstrated that apical polarity can be established before basolateral polarity and in the absence of tight-junctions (43). Grinsdstaff et al. (19) demonstrated that membrane polarity can precede the polarization of MTs. The hierarchy of events during the establishment of polarity is, therefore, still poorly understood. Our data here suggest a sequence of two groups of events: 1) an early reestablishment of the F-actin-based brush border, along with an incipient, partial, establishment of membrane polarity completed at 3 days after I/R, entirely consistent with the findings of Brown and co-workers (1); and 2) despite the repolymerization of MTs completed at the same time as the apical F-actin reorganization, a much delayed reestablishment of the polarized arrangement of MTs, along with the full polarization of the plasma membrane, both accomplished ~1 wk after I/R. In other words, the reestablishment of the submembrane F-actin does not seem to be sufficient for a full polarization of the cells.

An obvious possibility is that depolarization of apical membrane proteins may be the consequence of open tight junctions and be unrelated to the disorganization of MTs. In this regard, Kwon and co-workers (22) have reported up to 57% backleak of the glomerular filtrate in transplanted patients with sustained ARF, as demonstrated by fractional clearance of dextrans. However, it is of note that the same group in the above-mentioned and previous publications (2) found redistribution of Na+-K+-ATPase to a cytoplasmic pool but not to the apical membrane. The same result was confirmed here for the 30-min unilateral ischemia paradigm in the rat (Fig. 8). Those results would suggest a dissociation between the "fence" role of tight junctions, apparently conserved, and the "gate" role, clearly disrupted. In fact, in tissue culture cells, the gate function, measured by transepithelial resistance, can be disrupted with much shorter times of ATP depletion than the fence function (6).

Although we have not analyzed the polarity of apical ion transporters, the basolateral mislocalization of such will only potentiate the effects of mislocalized Na+-K+-ATPase by shortcircuiting the remaining pumps at the basolateral membrane and contributing to the failure in proximal sodium reabsorption. Recently, it has been speculated that it may even contribute to elevated tubule pressure (32), the predominant cause of hypofiltration in ARF. While it is clear that the modest proportion of depolarized cells in the 5- to 7-day period after I/R may not be significant for overall kidney function in the 30-min clamp paradigm, they highlight the possibility that slow apical polarization may play a role in delaying recovery after ARF under other ischemic conditions, such as allografts, as well. Other consequences of a sustained depolarization of apical proteins may go beyond ionic transport. Meprin, a brush-border protease, for example, has been found relocalized to the basolateral domain, where it caused fragmentation of the extracellular matrix after I/R in rats (12). These potential connections with the pathogenesis of ARF warrant the need for future investigations into the molecular mechanisms involved in the attachment-detachment of MTOCs in response to I/R injury in kidney.


   DISCLOSURES
 
This work was supported by National Institute of Diabetes and Digestive and Kidney Diseases Grant RO1-DK-57805 (to P. J. I. Salas).


   ACKNOWLEDGMENTS
 
The authors are indebted to Dr. N. A. Ameen for help with animal handling and inspiring discussions and Dr. A. Waheed (St. Louis University School of Medicine) for kindly providing anti-CAIV polyclonal antibody.


   FOOTNOTES
 

Address for reprint requests and other correspondence: P. J. I. Salas, Dept. of Cell Biology and Anatomy, R-124 -RMSB 4090, Univ. of Miami School of Medicine, 1600 NW 10th Ave., Miami, FL 33136 (E-mail: psalas{at}miami.edu).

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


   REFERENCES
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Abbate M, Bonventre JV, and Brown D. The microtubule network of renal epithelial cells is disrupted by ischemia and reperfusion. Am J Physiol Renal Fluid Electrolyte Physiol 267: F971–F978, 1994.[Abstract/Free Full Text]
  2. Alejandro VS, Nelson W, Huie P, Sibley RK, Dafoe D, Kuo P, Scandling JD Jr, and Myers BD. Postischemic injury, delayed function and Na+/K+-ATPase distribution in the transplanted kidney. Kidney Int 48: 1308–1315, 1995.[Web of Science][Medline]
  3. Ameen NA, Figueroa Y, and Salas PJI. Anomalous apical plasma membrane phenotype in CK8-deficient mouse enterocytes indicates a novel role of intermediate filaments in the polarization of simple epithelia. J Cell Science 114: 563–575, 2001.[Abstract]
  4. Apodaca G, Katz LA, and Mostov KE. Receptor-mediated transcytosis of IgA in MDCK cells is via apical recycling endosomes. J Cell Biol 125: 67–78, 1994.[Abstract/Free Full Text]
  5. Bacallao R, Antony C, Dotti C, Karsenti E, Stelzer EHK, and Simons K. The subcellular organization of Madin-Darby canine kidney cells during the formation of a polarized epithelium. J Cell Biol 109: 2817–2832, 1989.[Abstract/Free Full Text]
  6. Bacallao R, Garfinkel A, Monke S, Zampighi G, and Mandel L. ATP depletion: a novel method to study junction properties in epithelial tissues. I. Rearrangement of the actin cytoskeleton. J Cell Sci 107: 3301–3313, 1994.[Abstract]
  7. Bonventre JV. Mechanisms of ischemic acute renal failure. Kidney Int 43: 1160–1178, 1993.[Web of Science][Medline]
  8. Breitfeld PP, McKinnon WC, and Mostov KE. Effect of nocodazole on vesicular traffic to the apical and basolateral surfaces of polarized MDCK cells. J Cell Biol 111: 2365–2373, 1990.[Abstract/Free Full Text]
  9. Brown D, Lee R, and Bonventre JV. Redistribution of villin to proximal tubule basolateral membranes after ischemia and reperfusion. Am J Physiol Renal Physiol 273: F1003–F1012, 1997.[Abstract/Free Full Text]
  10. Buendia B, Bré MH, Griffiths G, and Karsenti E. Cytoskeletal control of centrioles movement during the establishment of polarity in MDCK cells. J Cell Biol 110: 1123–1135, 1990.[Abstract/Free Full Text]
  11. Canfield PE, Geerdes AM, and Molitoris BA. Effect of reversible ATP depletion on tight junction integrity in LLC-PK1 cells. Am J Physiol Renal Fluid Electrolyte Physiol 261: F1038–F1045, 1991.[Abstract/Free Full Text]
  12. Carmago S, Shah SV, and Walker PD. Meprin, a brush-border enzyme, plays an important role in hypoxic/ischemic acute renal tubular injury in rats. Kidney Int 61: 959–966, 2002.[Web of Science][Medline]
  13. Chen J, Dai J, Grant RL, Doctor RB, Sheetz MP, and Mandel LJ. Loss of cytoskeletal support is not sufficient for anoxic plasma membrane disruption in renal cells. Am J Physiol Cell Physiol 272: C1319–C1328, 1997.[Abstract/Free Full Text]
  14. Corrigan G, Ramaswamy D, Kwon O, Sommer FG, Alfrey EJ, Dafoe DC, Olshen RA, Scandling JD, and Myers BD. PAH extraction and estimation of plasma flow in human postischemic acute renal failure. Am J Physiol Renal Physiol 277: F312–F318, 1999.[Abstract/Free Full Text]
  15. Doctor RB, Bacallao R, and Mandel LJ. Method for recovering ATP content and mitochondrial function after chemical anoxia in renal cell cultures. Am J Physiol Cell Physiol 266: C1803–C1811, 1994.[Abstract/Free Full Text]
  16. Doctor RB, Bennett V, and Mandel LJ. Degradation of spectrin and ankyrin in the ischemic rat kidney. Am J Physiol Cell Physiol 264: C1003–C1013, 1993.[Abstract/Free Full Text]
  17. Donnahoo KK, Shames BD, Harken AH, and Meldrum DR. The role of tumor necrosis factor in renal ischemia-reperfusion injury. J Urol 162: 196–203, 1999.[Web of Science][Medline]
  18. Figueroa Y, Wald FA, and Salas PJI. p34cdc2 Mediated phosphorylation mobilizes microtubule organizing centers from the apical intermediate filament scaffold in Caco-2 epithelial cells. J Biol Chem 277: 37848–37854, 2002.[Abstract/Free Full Text]
  19. Grindstaff KK, Bacallao RL, and Nelson WJ. Apiconuclear organization of microtubules does not specify protein delivery from the trans-Golgi network to different membrane domains in polarized epithelial cells. Mol Biol Cell 9: 685–699, 1998.[Abstract/Free Full Text]
  20. Golenhofen N, Doctor RB, Bacallao R, and Mandel LJ. Actin and villin compartmentation during ATP depletion and recovery in renal cultured cells. Kidney Int 48: 1813–1845, 1995.
  21. Hropot M, Juretschke HP, Langer KH, and Schwark JR. S3226, a novel NHE3 inhibitor, attenuates ischemia-induced acute renal failure in rats. Kidney Int 60: 2283–2289, 2001.[Web of Science][Medline]
  22. Kwon O, Corrigan G, Myers BD, Sibley R, Scandling JD, Dafoe D, Alfrey E, and Nelson WJ. Sodium reabsorption and distribution of Na+/K+-ATPase during postischemic injury in the renal allograft. Kidney Int 55: 963–975, 1999.[Web of Science][Medline]
  23. Kwon TH, Frøkiær J, Han JS, Knepper MA, and Nielsen S. Decreased abundance of major Na+ transporters in kidneys of rats with ischemia-induced acute renal failure. Am J Physiol Renal Physiol 278: F925–F939, 2000.[Abstract/Free Full Text]
  24. Lafont F, Burkhardt JK, and Simons K. Involvement of microtubule motors in basolateral and apical transport in kidney cells. Nature 372: 801–803, 1994.[Medline]
  25. Meads T and Schroer TA. Polarity and nucleation of microtubules in polarized epithelial cells. Cell Motil Cytoskeleton 32: 273–288, 1995.[Web of Science][Medline]
  26. Molitoris BA. Ischemia-induced loss of epithelial polarity: potential role of the actin cytoskeleton. Am J Physiol Renal Fluid Electrolyte Physiol 260: F769–F778, 1991.[Abstract/Free Full Text]
  27. Molitoris BA, Falk SA, and Dahl RH. Ischemia-induced loss of epithelial polarity. Role of the tight junction. J Clin Invest 84: 1334–1339, 1989.[Web of Science][Medline]
  28. Nelson WJ and Yeaman C. Protein trafficking in the exocytic pathway of polarized epithelial cells. Trends Cell Biol 11: 483–486, 2001.[Web of Science][Medline]
  29. Nakagawa T, Sasahara M, Haneda M, Kataoka H, Nakagawa H, Yagi M, Kikkawa R, and Hazama F. Role of PDGF B-chain and PDGF receptors in rat tubular regeneration after acute injury. Am J Pathol 155: 1689–1699, 1999.[Abstract/Free Full Text]
  30. Nouwen EJ and De Broe ME. Human intestinal versus tissue-nonspecific alkaline phosphatase as complementary urinary markers for the proximal tubule. Kidney Int Suppl 47: S43–S51, 1994.[Medline]
  31. Oberbauer R, Schwarz C, Regele HM, Hansmann C, Meyer TW, and Mayer G. Regulation of renal tubular cell apoptosis and proliferation after ischemic injury to a solitary kidney. J Lab Clin Med 138: 343–351, 2001.[Web of Science][Medline]
  32. Ramaswamy D, Corrigan G, Polhemus C, Boothroyd D, Scandling J, Sommer FG, Alfrey E, Higgins J, Deen WM, Olshen R, and Myers BD. Maintenance and recovery stages of postischemic acute renal failure in humans. Am J Physiol Renal Physiol 282: F271–F280, 2002.[Abstract/Free Full Text]
  33. Reinsch S and Karsenti E. Orientation of spindle axis and distribution of plasma membrane proteins during cell division in polarized MDCKII cells. J Cell Biol 126: 1509–1526, 1994.[Abstract/Free Full Text]
  34. Rizzolo LJ and Joshi HC. Apical orientation of the microtubule organizing center and associated gamma-tubulin during the polarization of the retinal pigment epithelium in vivo. Dev Biol 157: 147–156, 1993.[Medline]
  35. Rodriguez ML, Brignoni M, and Salas PJI. A specifically apical sub-membrane intermediate filament cytoskeleton in non-brush-border epithelial cells. J Cell Sci 107: 3145–3151, 1994.[Abstract]
  36. Rodriguez-Boulan E, Salas PJ, Sargiacomo M, Lisanti M, Lebivic A, Sambuy Y, Vega-Salas D, and Graeve L. Methods to estimate the polarized distribution of surface antigens in cultured epithelial cells. Methods Cell Biol 32: 37–56, 1989.[Web of Science][Medline]
  37. Salas PJI. Insoluble {gamma}-tubulin containing structures are anchored to the apical network of intermediate filaments in polarized CACO-2 epithelial cells. J Cell Biol 146: 645–658, 1999.[Abstract/Free Full Text]
  38. Salas PJI, Misek D, Vega-Salas DE, Gundersen D, Cereijido M, and Rodriguez-Boulan E. Microtubules and actin filaments are not critically involved in the biogenesis of epithelial cell surface polarity. J Cell Biol 102: 1853–1867, 1986.[Abstract/Free Full Text]
  39. Safirstein R, Di Mari J, Megyesi J, and Price P. Mechanisms of renal repair and survival following acute injury. Semin Nephrol 18: 519–522, 1998.[Web of Science][Medline]
  40. Schwartz GJ, Olson J, Kittelberger AM, Matsumoto T, Waheed A, and Sly WS. Postnatal development of carbonic anhydrase IV expression in rabbit kidney. Am J Physiol Renal Physiol 276: F510–F520, 1999.[Abstract/Free Full Text]
  41. Shimizu A and Yamanaka N. Apoptosis and cell desquamation in repair process of ischemic tubular necrosis. Virchows Arch B Cell Pathol 65: 171–180, 1993.
  42. Stearns T and Kirschner M. In vitro reconstitution of centrosome assembly and function: the central role of {gamma}-tubulin. Cell 76: 623–637, 1994.[Web of Science][Medline]
  43. Vega-Salas DE, Salas PJI, Gundersen D, and Rodriguez-Boulan E. Formation of the apical pole of epithelial (Madin-Darby canine kidney) cells: polarity of an apical protein is independent of tight junctions while segregation of a basolateral marker requires cell-cell interactions. J Cell Biol 104: 905–916, 1987.[Abstract/Free Full Text]
  44. Wagner MC and Molitoris BA. Renal epithelial polarity in health and disease. Pediatr Nephrol 13: 163–170, 1999.[Web of Science][Medline]
  45. Wiese C and Zheng Y. A new function for the {gamma}-tubulin ring complex as a microtubule minus-end cap. Nat Cell Biol 2: 358–364, 2000.[Medline]
  46. Witzgall R, Brown D, Schwarz C, and Bonventre JV. Localization of proliferating cell nuclear antigen, vimentin, c-Fos, and clusterin in the postischemic kidney. Evidence for a heterogenous genetic response among nephron segments, and a large pool of mitotically active and dedifferentiated cells. J Clin Invest 93: 2175–2188, 1994.[Web of Science][Medline]
  47. Yin T, Sandhy G, Wolfgang CD, Burrier A, Webb RL, Rigel DF, Hai T, and Whelan J. Tissue-specific pattern of stress kinase activation in ischemic/reperfused heart and kidney. J Biol Chem 272: 19943–19950, 1997.[Abstract/Free Full Text]
  48. Yoshida T, Tang SS, Hsiao LL, Jensen RV, Ingelfinger JR, and Gullans SR. Global analysis of gene expression in renal ischemia-reperfusion in the mouse. Biochem Biophys Res Commun 291: 787–794, 2002.[Web of Science][Medline]
  49. Zuk A, Bonventre JV, Brown D, and Matlin KS. Polarity, integrin and extracellular matrix dynamics in the postischemic rat kidney. Am J Physiol Cell Physiol 275: C711–C731, 1998.[Abstract/Free Full Text]



This article has been cited by other articles:


Home page
 Am. J. Physiol. Renal Physiol.Home page
D. B. N. Lee, E. Huang, and H. J. Ward
Tight junction biology and kidney dysfunction
Am J Physiol Renal Physiol, January 1, 2006; 290(1): F20 - F34.
[Abstract] [Full Text] [PDF]

Home page
 Am. J. Physiol. Cell Physiol.Home page
K. Zahedi, Z. Wang, S. Barone, K. Tehrani, N. Yokota, S. Petrovic, H. Rabb, and M. Soleimani
Identification of stathmin as a novel marker of cell proliferation in the recovery phase of acute ischemic renal failure
Am J Physiol Cell Physiol, May 1, 2004; 286(5): C1203 - C1211.
[Abstract] [Full Text] [PDF]

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow All Versions of this Article:
285/2/F230    most recent
00024.2003v1
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Web of Science (5)
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Wald, F. A.
Right arrow Articles by Salas, P. J. I.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Wald, F. A.
Right arrow Articles by Salas, P. J. I.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
Visit Other APS Journals Online
Copyright © 2003 by the American Physiological Society.