Membrane repolarization is delayed in proximal tubules after ischemia-reperfusio - 《American Journal of Physiology》美国生理学杂志干细胞之家



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发表于 2009-4-21 13:42 |只看该作者 |倒序浏览 |打印

作者：Flavia A. Wald, Yolanda Figueroa, Andrea S. Oriolo, and Pedro J. I. Salas作者单位：Department of Cell Biology and Anatomy, University of Miami School ofMedicine, Miami, Florida 33136
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      【摘要】
   We have previously shown that microtubule-organizing centers (MTOCs) attachto the apical network of intermediate filaments (IFs) in epithelial cells inculture and in epithelia in vivo. Because that attachment is important for thearchitecture of microtubules (MTs) in epithelia, we analyzed whether chemical anoxia in LLC-PK 1 and CACO-2 cells or unilateralischemia-reperfusion in rat kidney (performed under fluorane anesthesia) hadan effect on the binding and distribution of MTOCs. In culture, we found thatchemical anoxia induces MTOC detachment from IFs by morphological andbiochemical criteria. In reperfused rat proximal tubules, noncentrosomal MTOCswere fully detached from the cytoskeleton and scattered throughout thecytoplasm 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 markersexpressed in S2 and S3 segments were depolarized at the same stage. At 8 daysafter reperfusion, membrane polarity, MTOCs, and MTs were back to normal.Na -K -ATPase was also found redistributed, not to theapical domain but rather to an intracellular compartment, as described byothers (Alejandro VS, Nelson W, Huie P, Sibley RK, Dafoe D, Kuo P, ScandlingJD Jr., and Myers BD. Kidney Int 48: 1308-1315, 1995). Theprolonged depolarization of the apical membrane may have implications in thepathophysiology of acute renal failure.
      【关键词】 ischemic acute renal failure apical polarity intermediate filaments
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SEVERAL PROCESSES INVOLVED in cellular injury during ischemia and after ischemia-reperfusion (I/R) have been reported in recent years in aneffort 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 aresignificant pathophysiological mechanisms of ischemic injury in proximaltubule cells ( 44 ). Mostattention has been focused on the disarrangement of the actin cytoskeleton( 20, 26 ), although it is known thatcortical fodrin, villin, and tight junctions are also affected( 9, 16, 27 ). Interestingly, however,it has been recently reported that disorganization of the actin cytoskeletonalone is not sufficient to explain the anoxic disruption of the plasmamembrane ( 13 ). In addition, Brown and co-workers ( 1 ) alsoshowed depolymerization of microtubules (MTs) in proximal tubules in vivoduring 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 MTsin a polarized fashion with the minus ends toward the apical membrane( 5 ). Although this apicobasalpolarization of MTs is not absolutely essential for polarization( 19, 38 ), it is thought toparticipate in the vesicular traffic bound to the apical membrane( 24 ), especially for vesiclesinvolved in transcytosis from basolateral to apical membrane( 8 ). In this regard, thenucleating activity of MTOCs must be necessary to repolymerize MTs after I/R.Because MTOCs cap the minus ends of MTs( 45 ), it is expected thatMTOCs positioned in their normal (apical) localization will reorganize MTswith the correct orientation (minus end apical). Conversely, if MTOCs becomedelocalized during ischemia, the MTs formed during the recovery period willhave abnormal orientations. In that scenario, aberrant MTs may transport thecarrier vesicles that originate in the trans -Golgi network andcontain apical cargo ( 28 ) toincorrect regions of the cytoplasm. Therefore, newly formed MTs with anincorrect orientation may contribute to the mispolarization of the plasmamembrane more seriously than a simple depolymerization of MT.8 e+ a7 \* D4 W! i
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Because our previous work suggests that the apical localization of MTOCs insimple epithelia depends on binding to apical intermediate filament (IF)( 37 ), we decided to analyzewhether the IFs are also disrupted by anoxia and whether the MTOCs are still bound to IF and apically localized in proximal tubule kidney cells afterischemia. 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 isknown to disrupt the MTs ( 1 )but causes little or no necrosis and only modest amounts of apoptosis onlywithin the first 24 h after ischemia( 31 ). The results indicatethat MTOCs become delocalized after chemical anoxia in tissue culture or as aresult of I/R in vivo. Furthermore, the anomalous localization of MTOCsresults in a disorganized array of MTs and correlates with poor membranepolarization of proximal tubule cells, delayed by days respect to theformation of the F-actin brush border.: {5 @5 S9 D- X% k1 L* p

MATERIALS AND METHODS9 R5 }' F, W9 s1 l

Cell culture and ATP depletion. LLC-PK 1 (porcineproximal tubule) and CACO-2 (human colon carcinoma epithelial cells) wereobtained from ATCC and maintained by weekly passages in tissue culture plasticflasks in DMEM-F-12 nutrient mixture (DMEM/F-12, GIBCO) supplemented with 10⺶al bovine serum (Cellgro). For chemical anoxia experiments, the methoddescribed by Molitoris and co-workers ( 11 ) was used, with thefollowing modifications. Monolayers confluent for 4 (LLC-PK 1 ) or 9days (CACO-2) were incubated in Earle's balanced salt solution (GIBCOformulation) without glucose or other nutrients for 30 min and then changed tothe same solution supplemented with 0.5 µM antimycin A (Sigma), 1 mMadenosine, and 0.2 mM allopurinol( 15 ) for 1 h (ATP depletion). Recovery was initiated by four washes and incubation in the standard DMEM/F-12culture medium for various times. ATP determinations were performed using aluciferase-based kit (Calbiochem)." p" h; ?- i" L6 y6 I: q
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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 gwere anesthetized with 1.5% isoflurane and kept on a warm plate to maintaintemperature. The peritoneal cavity was opened, and the left renal vessels wereclamped for 30 min with two microvascular (no. 1) clamps. The success of theclamp, and, later, the reperfusion, was visualized by rapid changes in thecolor of the kidney. The animals were sutured and kept under an infrared lampuntil fully awake. After various periods of time, the animals were killed byan 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 postureof muscles and tail.
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Antibodies and fluorescent reagents. The antibodies used in thisstudy were as follows: polyclonal antibody (Ab) anti- -tubulin NH 2 -terminal synthetic peptide (EEFATEGTDRKDVFFY, Sigma);monoclonal antibody against -tubulin (DM 1A; Sigma); monoclonalantibody against keratin (K) 19 (in CACO-2 cells; RCK-108; Accurate Chemical); monoclonal antibody against K19 (in LLC-PK 1 cells; A53/BA2; Sigma);monoclonal antibody against K18, B23.1 (Biomeda); polyclonal antibodyanti-intestinal alkaline phosphatase (iAP; DAKO); polyclonal antibodyanti-carbonic anhydrase IV (CAIV), a generous gift from Dr. A. Waheed (St.Louis University School of Medicine); and polyclonal antibodyanti-Na -K -ATPase, a generous gift from Dr. W. J.Nelson (Stanford Univ.). Secondary fluorescent antibodies were affinitypurified (Jackson Laboratories). FITC-phalloidin was purchased from MolecularProbes.( ?  m- J  h& q1 Z( t

Immunofluorescence and confocal microscopy. Immunofluorescence andconfocal microscopy of tissue culture cells were performed as described before( 37 ). Rat kidneys were fixedby intracardiac perfusion of the whole animal with warm 3% formaldehyde, 0.1% glutaraldehyde. The kidneys were rapidly extracted from the animal, sectionedwith a razor blade into 1-mm-thick sections, and further fixed in the samefixative for 2 h. Then, the sections were infused overnight in PBSsupplemented with 27% (wt/vol) sucrose, 0.3% formaldehyde, and 0.01%glutaraldehyde. The kidneys were frozen in the same solution in isopentane atmelting point and stored at -70°C. Sectioning was performed in a cryostat at -25°C. The sections, attached to a glass slide, werethawed by immersion in the same fixative described above and processed forimmunofluorescence following the standard protocol( 37 ). For quantification ofsignal in confocal sections, highly confocal images, collected at 0.7 Airyunits, were acquired, with care taken that no pixels in the positive signalwere saturated. Then, the images were analyzed by selecting separately in eachcell section the basolateral domain and the corresponding apical domain. Ineach selected area, we calculated the product of number of pixels timesaverage pixel values (luminosity channel) and the basolateral-to-apical ratiofor each cell. The data are presented as means ± SD of those ratios. Apoptosis in kidney sections was evaluated using a DNA fragmentation detectionkit (Oncogene Research Products), based on the end labeling of fragmented DNA,with biotin-labeled deoxynucleotides by Klenow DNA polymerase I.8 H. v7 k' @) R2 K4 `: t" |* D
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Cell extraction, fractionation, and immunoprecipitation of sonication fragments. Cell extraction, fractionation, and immunoprecipitation ofsonication fragments were done as previously described ( 34, 37 ). Briefly, confluentmonolayers of LLC-PK 1 cells were grown in one roller bottle. Thecells were extracted in 1% Triton X-100, in PBS supplemented with 2 mM EDTA,and a complete cocktail of antiproteases (Sigma), and the pellet wasextensively sonicated (3 min total, with 10-s periods of sonication and 15-slapses to dissipate heat). The sonication fragments of the Triton-insolublecytoskeleton were separated in 10-ml 20-60% sucrose continuous gradientsin 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 wasdone following standard procedures. The protein A-agarose beads werecentrifuged through a 0.5-ml 30% sucrose cushion. All these centrifugationswere done at 14,000 rpm for 2 s to minimize nonspecific copelleting of unboundcytoskeletal fragments. Immunoblotting was performed as described before( 37 ).
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RESULTS
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Relocalization of centrosomes during chemically induced ATP depletionand ischemia. To study the effects of chemical anoxia in tissue culturecells, we used a well-established protocol that is known to cause a rapid andreproducible depletion of ATP( 11 ). The efficiency ofchemically induced ATP depletion in our cells and the kinetics of recoveryafter 1-h depletion were assessed in LLC-PK 1 (porcine proximaltubule) and CACO-2 (human colon carcinoma) cells using a standard luciferaseassay. 90% within the first 10 minafter treatment with antimycin A, and significant recoveries of ATP levelswere started between 2 and 4 h after the cells were replaced in DMEM( Fig. 1 ). Thus the kinetics ofATP depletion/recovery for LLC-PK 1 were similar to those inprevious reports.  k- D! S9 r1 H: k1 m! V

Fig. 1. ATP levels in CACO-2 ( A ) and LLC-PK 1 ( B )epithelial cells in tissue culture during and after chemically induced ATPdepletion. The ATP from quadruplicate confluent monolayers ( 3 x 10 5 cells each) was extracted at various times of incubation innutrient-free medium supplemented with antimycin A, adenosine, and allopurinol(depletion) and after a 1-h depletion at various times of recovery in normalculture medium. ATP was measured by luciferase luminescence by integratinglight emission for 1 min.% V- }, |. x9 S, H, d  l

To analyze the localization of centrosomes and to extend the observationsto a different cell line, LLC-PK 1 and CACO-2 monolayers weresubjected to ATP depletion for 1 h and 1-h recovery. Double-immunofluorescence experiments were performed colocalizing the -tubulin signal ( Fig. 2, green) with thecortical 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 Ikeratin, distributed under the apical and lateral domains inLLC-PK 1 cells, but the IF cortical signal was thicker in the kidneycells than in the intestinal cells ( Fig. 2, a, c, e, g, and i ). More importantly, ATPdepletion did not affect the IF cytoskeleton in either LLC-PK 1 orCACO-2 cells ( Fig. 2 ). Asimilar result was observed, reciprocally, using anti-K18 or anti-K19antibodies in CACO-2 or LLC-PK 1 cells, respectively (notshown).
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Fig. 2. Centrosomes detach from their normal apical location at the apicalintermediate filament (IF) network after 1-h ATP depletion inLLC-PK 1 and CACO-2 cells. Monolayers of LLC-PK 1 ( a, c, e, g, i ) and CACO-2 cells( b, d, f, h, j ) were grown onTranswell filters. Some cells were subjected to ATP depletion for 1 h( e - j ). The cells were fixed after 1-h recovery andprocessed for double immunofluorescence with monoclonal Abs against K18(LLC-PK 1 cells) or K19 (CACO-2 cells) (red channel) and apolyclonal antibody against a conserved NH 2 -terminal polypeptide of -tubulin (green channel). Two examples of each control and 3 examplesof ATP-depleted cells were analyzed by confocal microscopy, deconvolution, and3D reconstruction and are shown in the XZ plane (perpendicular to themonolayer), with the apical side up. Arrows point at -tubulin signalseparated from the apical domain IFs. Scale bars: 5µm( a - i ) and 2 µm( j ).
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In Fig. 2, examples offields with centrosomes within the same confocal optical section in nearbycells are shown for control monolayers ( a - d ) andATP-depleted cells ( e - j ). As expected from previouspublications, the centrosomes in control cells were observed always embeddedin the apical IF submembrane cytoskeleton( Fig. 2, a - d ). In ATP-depleted cells, a proportionof the centrosomes was found separated from the apical IF cytoskeleton( Fig. 2, arrows). Theproportion was smaller in LLC-PK 1 cells ( 28%) than in CACO-2cells ( 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

Because colocalization does not necessarily imply attachment, the physicalconnection of insoluble -tubulin-containing structures and IF wasanalyzed as before ( 37 ) byfragmenting the Triton-insoluble cytoskeleton of LLC-PK 1 cells withextensive sonication and separating the smallest fragments by size in sucrosegradients by rate centrifugation. This procedure was originally devised toperform immunoprecipitation in highly insoluble multiprotein complexes( 34 ). We collected the topfive fractions of the gradient that contain fragments small enough to beimmunoprecipitated (larger fragments tend to nonspecifically copellet withagarose beads). Each aliquot was divided into two equal parts, one of themimmunoprecipitated with an irrelevant rabbit IgG as a negative control( Fig. 3, - lanes), andthe other with anti- -tubulin Ab( Fig. 3, lanes). Theimmunoprecipitates were analyzed by immunoblot using anti-K18 or anti-K19monoclonal antibody. In general, the previous findings in CACO-2( 37 ) were extended toLLC-PK 1 cells. In control cells, there was coimmunoprecipitation ofK18 and -tubulin in fractions 1, 3, and 4 and of K19and -tubulin in fractions 1-5 ( Fig. 3 ). Thiscoimmunoprecipitation was mostly abolished in LLC-PK 1 cellssubjected to 1-h ATP depletion. K18 blots showed only background levels, andK19 showed only some ATP depletion-insensitive coimmunoprecipitation with -tubulin in fraction 4 ( Fig. 3 ). Controls showing thatthis coimmunoprecipitation cannot be explained by simple physical trapping orartifactual protein binding as a result of detergent extraction have beenperformed elsewhere ( 37, see Fig. 8 ). These results indicatethat the attachment between MTOC and IFs is largely broken after 1-h ATPdepletion in LLC-PK 1 and CACO-2 cells in tissue culture./ _4 t9 S5 Z# P0 O! Y3 H
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Fig. 3. ATP depletion inhibits coimmunoprecipitation of insoluble -tubulinwith K18 and 19 in LLC-PK 1 cells. LLC-PK 1 cellmonolayers were grown in 2 roller bottles ( 1.5 x 10 8 cells each). One of the cultures was ATP depleted for 1 h. The cells wereextracted in Triton X-100, and the insoluble cytoskeletal preparations wereextensively sonicated. The fragments were separated by size by ratecentrifugation in sucrose gradients and fractions 1-5 ( top half of the gradient) divided in 2 equal aliquots each. Thealiquots were immunoprecipitated with anti- -tubulin antibody (  lanes)or an irrelevant rabbit IgG (- lanes). The immunoprecipitates wereanalyzed by immunoblot using monoclonal Abs against K18 and K19 sequentiallyon the same blot. Left : molecular mass standards (in kDa).0 E. s+ b: x+ ~! `" \

Fig. 8. Intracellular localization of "depolarized"Na -K -ATPase after I/R in proximal tubules. Sections ofthe right (sham; 3 days after operation) and left (I/R) kidneys fixed 3 and 5days after renal artery clamp operation were processed for indirectimmunofluorescence with an anti-Na -K -ATPase antibody(red channel) and FITC-phalloidin (green). The arrows point to cells withintracellular distribution of Na -K -ATPase. Bars: 10µm./ k. A$ C8 b" L" p- T
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Ischemic injury in vivo causes detachment of MTOCs in proximal tubules. To test whether MTOCs also detach from the apical domain in vivoafter ischemia, rat kidneys were subjected to a 30-min clamp of the renalvessels. The contralateral kidneys were used as a sham operation control. Weanalyzed the kidneys at 24 h after I/R, a time that has been widely studiedbut that yields results difficult to analyze because of the complexity of theeffects. We also analyzed the kidneys at 3, 5, 7, and at 8 days after I/R. Thelatter was chosen because our preliminary experiments showed that the changesdescribed in this section last up to 7 days after I/R, and kidney functionalparameters with a 30-min bilateral clamp protocol have been reported to benearly normal as early as 5 days after I/R( 23 ). In addition, Hropot andco-workers ( 21 ) found plasmacreatinine and sodium still abnormal after 7 days for a 40-min bilateral clampmodel. Even in a different system, the human renal allograft, patients recovering well achieve acceptable glomerular filtration rates on day7 ( 14 ).' o7 ~, m5 {3 y" o6 M/ o9 W7 r

Frozen sections from reperfused or sham-operated kidneys were analyzed byimmunofluorescence with anti- -tubulin antibody and FITC-phalloidin tostain F-actin. In the control proximal tubules, very few 0.3- to 0.4-µmspots of -tubulin signal (centrosomes) were easily recognizable andwere mostly localized in the apical pole of the cells. However, a continuoussubapical layer of -tubulin signal was observed in most tubules both in kidneys subjected to a sham operation ( Fig.4 a ) and in kidneys from animals not subjected to surgery(not shown). When analyzed in sections perfectly perpendicular to the axis ofthe tubule, the -tubulin signal( Fig. 4 a, inset, green) could be localized immediately below the F-actin signalin the brush border ( Fig. 4 a, inset, red; arrows point at basal actinsignal). This image is consistent with findings in our laboratory and byothers as well that a layer of noncentrosomal MTOCs lies under the apicaldomain ( 3, 25 ). During the first 24 hafter I/R, the apical -tubulin layer became very discontinuous ordisappeared altogether (not shown). However, because the entire apical domain is degraded ( 44 ), it isdifficult to draw any conclusions about the fate of MTOCs at that time. Moreinteresting were the results from kidneys at 3 days following I/R. At thisstage, the F-actin component of the brush border was mostly repaired in many(but not all) cells, as judged by a phalloidin label( Fig. 4 d, arrows). However, no subapical layer of -tubulin was observed, even in tubules(or cells) with a complete brush border( Fig. 4, c vs. d, arrows). Instead, -tubulin signal was eitherdiffuse in the cytoplasm or concentrated under the lateral domain in somecells ( Fig. 4 c,arrows). At 8 days after I/R, tubules subjected to a 30-min ischemic injuryhad recovered the normal distribution of -tubulin and the images becameindistinguishable from controls in all cells (like in Fig. 4 a ). This result suggests that the repolarization of MTOCs is delayed with respect to thereassembly of the F-actin brush border.! i) V7 s$ a  ?) b9 B3 x/ G% E
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Fig. 4. Polarization of microtubule-organizing centers (MTOCs) is delayed respectto the formation of the brush border in proximal tubules. Frozen sections fromthe kidneys of an animal that was subjected to a 30-min clamp of the leftrenal vessels followed by a 3-day recovery ( c and d ) andsham-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- -tubulinantibody ( a and c ). Inset : higher magnificationview of a segment of a proximal tubule overlapping pseudocolor images ofphalloidin (red) and anti- -tubulin (green) signals. L, lumen;arrowheads, basal actin signal. Arrows point out depolarized -tubulinsignal ( c ) and fully reassembled F-actin brush border ( d ).Scale bars: 20 µm ( a - d ) and 5 µm( inset ).
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The same experiment was repeated in several animals, i.e., the harvestingof kidneys at 1, 3, and 8 days after I/R. To assess the attachment of MTOCs tothe cytoskeleton, pieces of the cortex, freshly obtained from the animal, werehomogenized and centrifuged at low speed, so that the soluble 27S -tubulin complexes ( -TurC; 42) remain in the supernatant and -tubulin attached to the cytoskeleton in the pellet. In previousstudies, roughly 50% of the -tubulin was found soluble( 37, 42 ), and that was the case forkidney cortexes subjected to a sham operation (S in Fig. 5, pellets vs. soluble).Despite the disruption in the architecture of the tissue, and with somevariability, 37% of the -tubulin (with respect to control) was stillassociated with the cytoskeletal pellet after 1 day of recovery (I in Fig. 5 and Table 1 ). This result can bedue to minimal amounts of MTOCs still attached to the apical cytoskeleton,which may go undetected by regular morphology. Interestingly, however, no -tubulin was found associated with the pellet fraction at 3 days afterI/R ( Fig. 5, Table 1 ). At 8 days after I/R, the fraction of -tubulin in the pellet resembled again that of thesham-operated kidneys ( Fig. 5, Table 1 ). Together with thedata in Fig. 4, this resultsuggests that the bonding of MTOCs to the cytoskeleton breaks down after I/Rand that it takes days to be reestablished. This is a much longer time periodthan it takes to reassemble apical F-actin at the brush border.7 d  T1 Y9 [7 q7 m. {8 T

Fig. 5. MTOCs are detached from the cytoskeleton in the renal cortex 3 days afteran ischemic injury. The left kidneys of rats were subjected to a 30-min clampfollowed by 1, 3, or 8 days of reperfusion (I). The corresponding rightkidneys were used as sham controls (S). Each S and I pair corresponds to adifferent animal. Pieces of cortex were homogenized in nonionic detergent, andthe cytoskeletal (detergent insoluble) fraction was pelleted at low-speedcentrifugation, which was devised to maintain -tubulin (tub.) complexes( -TurC, 42) in the supernatant. The pellets were rehomogenized bysonication. Then, 30 µg total protein of each sample were separated bySDS-PAGE and analyzed by immunoblot with an anti- -tubulin antibody. Thesame membranes were reprobed with an anti-keratin antibody (pellets) or ananti- -tubulin antibody (supernatants) to independently assess proteinload in the same lanes.
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Table 1. -Tubulin signal in the cortex of kidneys subjected to I/R as apercentage of the band from the contralateral (sham surgery) kidney/ ^" Y' S& m3 T: c0 @/ t

Disorganization of MTs at 3 days after I/R in proximal tubules. Itis known that MTs disassemble within the first 24 h after I/R( 1 ). We confirmed thatobservation in kidney sections stained with an anti- -tubulin antibody.The few MTs remaining at 1 day after I/R, however, were still oriented in theapicobasal 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 ofthe delocalization of MTOCs described above at 3 days after I/R, we wereprompted to analyze their organization at that particular time. Observation of the sections with regular epifluorescence microscopy revealed that the densityof MTs per cell was similar to that in sham-operated kidneys (shown only inconfocal microscopy, Fig. 6, a vs. b ), suggesting that an active process of repolymerizationof tubulin occurs between 1 and 3 days after I/R. The thickness of thesections ( 5 µm), compared with the diameter of single MTs ( 20nm), however, made it necessary to visualize MT bundles under confocalmicroscopy using a high degree of confocality (0.7 Airy units), which giveshigh resolution in the z -axis. Proximal tubules from kidneyssubjected to a sham operation observed under those conditions showed MTbundles mostly oriented in the apicobasal axis( Fig. 6 a ), asdescribed extensively before( 3, 5, 37 ). In the proximal tubulesat 3 days after I/R, on the other hand, different arrangements were observed.In some cases, MTs were visualized as a disorganized network( Fig. 6 b, red channel,arrows), mostly separated from the apical F-actin layer (green channel). Inother sections, MTs adopted a basolateral distribution, and many bundles were roughly perpendicular to the apicobasal axis( Fig. 6 c, arrows), anorientation seldom seen in control cells.

Fig. 6. Microtubules remain disorganized 3 days after ischemia-reperfusion (I/R) intubules with normal F-actin distribution. Sections of the right (sham; a ) and left (I/R; b and c ) kidneys fixed 3 daysafter renal artery clamp operation were stained with an anti- -tubulinmonoclonal Ab (red channel) and subsaturation concentrations ofFITC-phalloidin to minimally counterstain the brush-border F-actin (greenchannel). The sections were analyzed by confocal microscopy at 0.7 Airy units.Arrows, abnormal microtubule distribution. Scale bars: 10 µm.3 ~' v$ }+ s1 i9 C# g4 x
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Lack of polarity of apical plasma membrane markers at 3 days after I/Rin proximal tubules showing an apparently normal F-actin brush border. Because MTOCs cap the minus end of MTs ( 45 ) and because MTOCs seem tobe scattered in the cytoplasm at 3 days after I/R( Fig. 4 ) free from their normalcytoskeletal attachment ( Fig.5 ), we reasoned that the disorganized MTs at 3 days after I/R( Fig. 6 ) must have their minusends randomly distributed in the cells. Therefore, MTs would not only not contribute to the normal polarization of the plasma membrane but wouldactually tend to randomize apical membrane proteins. To test this hypothesis,we analyzed the polarization of two apical membrane markers normally expressedin 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 byphalloidin distribution ( Fig. 7, d vs. b ). CAIV and iAP were observed only in the brushborder in all tubules where they are expressed in sham-operated kidneys( Fig. 7 a, Table 2 ). At 1 day after I/R,the iAP signal could not be observed and the CAIV signal was weak anddepolarized in at least one cell in 72% of the S2-3 sections( Table 2 ). This result isconsistent 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. 7 c, arrows) wereobserved in one or more cells in 55% of the S3 sections( Table 2 ). Similarly, 64% ofthe sections positive for iAP showed at least one cell depolarized( Table 2 ). The number ofdepolarized cells decreased significantly on day 5, although isolateddepolarized cells were observed up to 7 days after I/R for CAIV( Table 2 ).' {6 x# {5 K; l6 d- H& X
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Fig. 7. Substantial amounts of basolateral carbonic anhydrase IV (CAIV) signalremain 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 renalartery clamp operation were processed for indirect immunofluorescence with ananti-CAIV antibody ( a, c, e, g ) followedby a specific secondary antibody coupled to CY3 and FITC-phalloidin to showF-actin ( b, d, f, h ). e - h : Confocal images of the same preparations athigher magnification. Scale bars: 20 µm ( a - d ) and10 µm ( e - h ).7 s: g, n5 s7 c/ z6 a  ~
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Table 2. Delayed recovery of epithelial polarization in S3
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To obtain a more precise assessment of the magnitude of the depolarizationof CAIV and iAP, we measured the total signal in the apical and basolateraldomain 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 imagesin which high-intensity pixels are saturated, this method can provide a fairestimation of the relative distribution of a membrane marker in a way that accounts for the differences in membrane folding( 36 ). In proximal tubulessubjected to sham surgery, only 3.6 ± 4.2% of total CAIV signal waslocalized to the basolateral domain ( Fig. 7, a and e; only detectable with digitalmicroscopy). In kidneys at 3 days after I/R, on the other hand, proximaltubule cells showed 48.2 ± 13.3% of total cellular CAIV signallocalized to the basolateral domain ( Fig.7, c, arrows, and g ). In other words, thesecells "look" partially polarized because the apical signal islocalized in a smaller area of the section with a high level of membranefolding (brush-border microvilli), whereas the basolateral signal is in a muchmore stretched membrane array. However, nearly equal amounts of the apicalmembrane marker are localized in each domain, and, thus these cells are almosttotally depolarized./ z& ?' a, C9 {$ T+ s- e

Intracellular localization ofNa -K -ATPase after I/R. To analyze the polarity of basolateral membrane proteins during the same periodafter I/R, we conducted experiments similar to those depicted in Fig. 7 but using ananti-Na -K -ATPase antibody instead. The antibody showeda 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 tubulecells showed intracellular images, even in cells with an already fullyreassembled actin brush border ( Fig.8, arrows). At this stage, however, little or no apicaldistribution of Na -K -ATPase was observed. At 5 daysafter I/R, only scattered cells (21% of the sections, an average of 1.2cells/section) were still showing the intracellular distribution ofNa -K -ATPase, and with few exceptions (one shown in Fig. 8 C ), noNa -K -ATPase signal was found in the apical domaineither ( Fig. 8 ). These resultssuggest that Na -K -ATPase also redistributes at the3-day stage after I/R but not in the apicobasal axis but rather into anintracellular compartment and fully confirm the observations of Alejandro andco-workers ( 2 ) in humanallografts. In addition, the results also indicate that different mechanismsgovern the polarization of apical and basolateral markers, as described beforein Madin-Darby canine kidney cells( 43 ) and that reassembly ofthe brush-border actin is not an indicator of full polarization.+ ~: Q$ R9 V8 V' F, D, f
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DISCUSSION
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The results in this work point at three general conclusions. First, IFs arestable after ATP depletion in culture or I/R in vivo (Figs. 2 and 5 ). Second, I/R leads to aseparation of MTOCs from the cytoskeleton, in culture( Fig. 2 ) and in vivo (Figs. 4 and 5 ). This detachment results ina 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 whenthe microfilaments are fully reorganized (2-3 days after I/R)( Fig. 6 ). Third, at least someplasma membrane proteins remain depolarized after the reorganization of the brush border, as defined by F-actin, and repolarize roughly with the samekinetics as MTOCs and MTs reacquire their normal subcellular organization( Fig. 7 ).

The initial experiments in this work were performed in two epithelial celllines in tissue culture: LLC-PK 1, derived from the proximal tubule,and CACO-2, derived from the intestine. The results from both cell linesindicate that ATP depletion has a significant impact on the apicallocalization of MTOCs. In proximal tubules, centrosomes were difficult toidentify, and perhaps they are absent altogether in most cells. Therefore, werather focused on the layer of noncentrosomal MTOCs that has been visualized in vivo in other epithelia as well( 3 ). The dramatic fate of theapical pole in vivo as a result of the I/R injury, extensively described byMolitoris and co-workers ( 11 ),made any comparison of the redistribution of MTOCs in culture vs. in vivonearly impossible. In addition, the analysis of the initial 24 h after I/R iscomplicated by the presence of apoptotic cells( 31 ). Biochemical andmorphological data in this work (Figs. 4 and 5 ) indicate that between 24 and72 h after I/R, nearly all MTOCs become detached from the cytoskeleton and arescattered 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 casesremained unaffected by ATP depletion or I/R (Figs. 2 and 5, respectively). The immediateconsequence of the detachment of MTOCs was that MTs, although fullyrepolymerized during the same period of time (1-3 days after I/R),became a disorganized network, instead of the normal array of bundles highlyoriented in the apicobasal axis ( Fig.6 ). Although we cannot assert a cause-and-effect relationship, itis safe to conclude that the disorganization of MTs correlates with poorlevels of plasma membrane polarity, which only revert over a period of days tonearly normal at 5 days after I/R and, in some isolated depolarized cells, upto 7 days after I/R ( Fig. 7, Table 2 ).$ [! K, ^& ]2 F3 o) N, `; [% S$ t' e, e

There are a number of possible explanations for the delayed recovery ofdistribution 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 asimple action of caspases. However, proximal tubule cells at 3 days after I/Rdisplayed a nearly normal F-actin cytoskeleton and full-length MTs (althoughin abnormal orientations) (Figs. 6 b and 7 d ). Furthermore, andin agreement with previous publications using the same I/R protocol( 31 ), we found only modestproportions 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 inthe first place, as opposed to more drastic ischemic injuries (e.g., 1 h) thatresult in higher percentages of apoptotic cells, and, eventually, a latesecondary peak of apoptosis( 41 ). In addition, we havedemonstrated that, at least in intestinal epithelia, apoptosis does not disrupt epithelial polarity as long as the cell maintains the integrity of itsplasma membrane ( 3 ). On theother 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 areeither upregulated or downregulated using the same I/R protocol at 1 or 3 daysafter I/R ( 48 ). More specifically, the expression of vimentin has been implicated as a reporter ofdedifferentiation in proximal tubule cells after ischemia, and, along with theexpression of proliferating cell nuclear antigen (PCNA), it was found in asubstantial number of S3 cells at 2-5 days postischemia using a harsher ischemic injury ( 46 ). In oursystem, vimentin was expressed in
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One lesson that we can draw from various studies of the acquisition ofmembrane polarity in tissue culture cells is that the time course ofpolarization of various components may be different and sometimescounter-intuitive. For example, we have demonstrated that apical polarity canbe established before basolateral polarity and in the absence oftight-junctions ( 43 ).Grinsdstaff et al. ( 19 )demonstrated that membrane polarity can precede the polarization of MTs. Thehierarchy of events during the establishment of polarity is, therefore, stillpoorly understood. Our data here suggest a sequence of two groups of events: 1 ) an early reestablishment of the F-actin-based brush border, alongwith an incipient, partial, establishment of membrane polarity completed at 3days 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-actinreorganization, a much delayed reestablishment of the polarized arrangement ofMTs, along with the full polarization of the plasma membrane, bothaccomplished 1 wk after I/R. In other words, the reestablishment of thesubmembrane F-actin does not seem to be sufficient for a full polarization ofthe cells.7 m6 ?$ ?: ^9 L/ |  n: P* h: k
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An obvious possibility is that depolarization of apical membrane proteinsmay be the consequence of open tight junctions and be unrelated to thedisorganization of MTs. In this regard, Kwon and co-workers( 22 ) have reported up to 57篶kleak of the glomerular filtrate in transplanted patients with sustained ARF, as demonstrated by fractional clearance of dextrans. However, it is ofnote that the same group in the above-mentioned and previous publications( 2 ) found redistribution ofNa -K -ATPase to a cytoplasmic pool but not to theapical membrane. The same result was confirmed here for the 30-min unilateralischemia 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 disruptedwith much shorter times of ATP depletion than the fence function( 6 ).

Although we have not analyzed the polarity of apical ion transporters, thebasolateral mislocalization of such will only potentiate the effects ofmislocalized Na -K -ATPase by shortcircuiting theremaining pumps at the basolateral membrane and contributing to the failure inproximal sodium reabsorption. Recently, it has been speculated that it mayeven contribute to elevated tubule pressure( 32 ), the predominant cause ofhypofiltration in ARF. While it is clear that the modest proportion ofdepolarized 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 thepossibility that slow apical polarization may play a role in delaying recoveryafter ARF under other ischemic conditions, such as allografts, as well. Otherconsequences of a sustained depolarization of apical proteins may go beyond ionic transport. Meprin, a brush-border protease, for example, has been foundrelocalized to the basolateral domain, where it caused fragmentation of theextracellular matrix after I/R in rats( 12 ). These potentialconnections with the pathogenesis of ARF warrant the need for futureinvestigations into the molecular mechanisms involved in theattachment-detachment of MTOCs in response to I/R injury in kidney.
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DISCLOSURES! C& s4 k! ~: \  w: f* p
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This work was supported by National Institute of Diabetes and Digestive andKidney Diseases Grant RO1-DK-57805 (to P. J. I. Salas).+ p5 X0 {/ f. h' N! W( b

ACKNOWLEDGMENTS
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The authors are indebted to Dr. N. A. Ameen for help with animal handlingand inspiring discussions and Dr. A. Waheed (St. Louis University School ofMedicine) for kindly providing anti-CAIV polyclonal antibody.* b1 q3 T# {* n- A
      【参考文献】
Abbate M,Bonventre JV, and Brown D. The microtubule network of renal epithelialcells is disrupted by ischemia and reperfusion. Am J Physiol RenalFluid Electrolyte Physiol 267:F971-F978, 1994.# k6 V4 u3 c. g: H9 p! m9 e
& b: T1 [9 g  c2 A$ Y

Alejandro VS,Nelson W, Huie P, Sibley RK, Dafoe D, Kuo P, Scandling JD Jr, and MyersBD. Postischemic injury, delayed function andNa /K -ATPase distribution in the transplantedkidney. Kidney Int 48:1308-1315, 1995./ ^4 @2 @% I  D& E, Y! [, V: O2 G

Ameen NA,Figueroa Y, and Salas PJI. Anomalous apical plasma membrane phenotype inCK8-deficient mouse enterocytes indicates a novel role of intermediatefilaments in the polarization of simple epithelia. J CellScience 114:563-575, 2001.; I; A. L& f3 U- K

Apodaca G, KatzLA, and Mostov KE. Receptor-mediated transcytosis of IgA in MDCK cells isvia apical recycling endosomes. J Cell Biol 125: 67-78,1994.: K4 f' Z% ^) }5 e9 D

5 k: V  T& Y1 w% q9 _1 Z. S3 U

Bacallao R,Antony C, Dotti C, Karsenti E, Stelzer EHK, and Simons K. The subcellularorganization of Madin-Darby canine kidney cells during the formation of apolarized epithelium. J Cell Biol 109: 2817-2832,1989.

2 S. j, [6 X' A0 U* @# o
% q" M5 p* q: `0 m5 D
Bacallao R,Garfinkel A, Monke S, Zampighi G, and Mandel L. ATP depletion: a novelmethod to study junction properties in epithelial tissues. I. Rearrangement ofthe actin cytoskeleton. J Cell Sci 107: 3301-3313,1994.2 T9 b% O/ q- Z, T. d! q9 t; S

$ F+ }" k- I% m4 s
Bonventre JV. Mechanisms of ischemic acute renal failure. Kidney Int 43:1160-1178, 1993.

Breitfeld PP,McKinnon WC, and Mostov KE. Effect of nocodazole on vesicular traffic tothe apical and basolateral surfaces of polarized MDCK cells. J CellBiol 111:2365-2373, 1990.- K* ?) y# n0 A+ {7 {
8 U+ \: {1 ?5 f( Y" Z# G* }

6 _. |8 q0 H" U, M) b; M
Brown D, Lee R,and Bonventre JV. Redistribution of villin to proximal tubule basolateralmembranes after ischemia and reperfusion. Am J Physiol RenalPhysiol 273:F1003-F1012, 1997.* G! y% V# B$ a  S! x; E
0 {7 Q8 z' @" w! N

& W3 N; w( C; a5 S/ R
Buendia B,Bré MH, Griffiths G, and Karsenti E. Cytoskeletal control ofcentrioles movement during the establishment of polarity in MDCK cells. J Cell Biol 110:1123-1135, 1990.
/ Z6 q4 ^# }% |" P; h; o& Z
  I' `2 {$ n5 l8 b1 b6 T
; q/ g1 K: ~( w: y4 u
Canfield PE,Geerdes AM, and Molitoris BA. Effect of reversible ATP depletion on tightjunction integrity in LLC-PK 1 cells. Am J Physiol RenalFluid Electrolyte Physiol 261:F1038-F1045, 1991.
: |6 q) `$ B% B7 S! M; l8 r
+ \2 H: N& b0 X, _9 X  h% J

Carmago S, ShahSV, and Walker PD. Meprin, a brush-border enzyme, plays an important rolein hypoxic/ischemic acute renal tubular injury in rats. KidneyInt 61:959-966, 2002.

- n6 X( F% B5 w5 G. Q' T. |

Chen J, Dai J,Grant RL, Doctor RB, Sheetz MP, and Mandel LJ. Loss of cytoskeletalsupport is not sufficient for anoxic plasma membrane disruption in renalcells. Am J Physiol Cell Physiol 272: C1319-C1328,1997./ ]) t) b" P9 r4 ^% ~$ [/ V/ h9 k

* v* i" O* M, P: b

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 humanpostischemic acute renal failure. Am J Physiol RenalPhysiol 277:F312-F318, 1999.# s  h- w! Y7 v# R- g; K& W

/ I% ~1 w9 U; U' F, P4 Z

Doctor RB,Bacallao R, and Mandel LJ. Method for recovering ATP content andmitochondrial function after chemical anoxia in renal cell cultures. Am J Physiol Cell Physiol 266:C1803-C1811, 1994.6 q, v. W* c8 v' @

4 ]# O2 A) ]5 X# ]
* c! a' D( Y8 \6 x+ `/ t; n( i# j
Doctor RB,Bennett V, and Mandel LJ. Degradation of spectrin and ankyrin in theischemic rat kidney. Am J Physiol Cell Physiol 264: C1003-C1013,1993.; P9 e& h0 A0 R' G
7 }3 A! ]$ k$ E' @
4 M6 X. B1 t7 b

Donnahoo KK,Shames BD, Harken AH, and Meldrum DR. The role of tumor necrosis factor inrenal ischemia-reperfusion injury. J Urol 162: 196-203,1999.6 n! A: t$ N0 `; C  Y, q

' Y5 q# r8 ?" R
! b$ _0 n' H0 j
Figueroa Y,Wald FA, and Salas PJI. p34 cdc2 Mediatedphosphorylation mobilizes microtubule organizing centers from the apicalintermediate filament scaffold in Caco-2 epithelial cells. J BiolChem 277:37848-37854, 2002.

0 V9 h# ]7 A3 X  O2 H3 Z/ L; o
2 A% c' W6 o0 R6 h# n
Grindstaff KK,Bacallao RL, and Nelson WJ. Apiconuclear organization of microtubules doesnot specify protein delivery from the trans-Golgi network to differentmembrane domains in polarized epithelial cells. Mol BiolCell 9:685-699, 1998.6 i( x- l% u# |+ v/ e2 @
1 P  E# `6 W# @! T" f: |
4 H9 Y, ]( w) @: {* O  y# X$ \: o

Golenhofen N,Doctor RB, Bacallao R, and Mandel LJ. Actin and villin compartmentationduring ATP depletion and recovery in renal cultured cells. KidneyInt 48:1813-1845, 1995.
; B  L0 @7 W2 y3 t- q- f. T, q
3 I. g0 i9 q. z( H
2 B& A' f/ v: C  M' m$ g
Hropot M,Juretschke HP, Langer KH, and Schwark JR. S3226, a novel NHE3 inhibitor,attenuates ischemia-induced acute renal failure in rats. KidneyInt 60:2283-2289, 2001., u( i4 L6 P( X

' Z# p# H5 \- y5 F" R3 Z

Kwon O,Corrigan G, Myers BD, Sibley R, Scandling JD, Dafoe D, Alfrey E, and NelsonWJ. Sodium reabsorption and distribution ofNa /K -ATPase during postischemic injury in the renalallograft. Kidney Int 55:963-975, 1999.4 {: U! T9 v$ N9 E' t, g7 X8 S6 ~

% L$ q* H$ g6 n4 b) x

Kwon TH,Frøkiær J, Han JS, Knepper MA, and Nielsen S. Decreasedabundance of major Na transporters in kidneys of rats withischemia-induced acute renal failure. Am J Physiol RenalPhysiol 278:F925-F939, 2000.) V' P$ n  l- j( `; Y: ~

( I: B! b& d% j! S( N
7 h: C6 ]: X& v% u
Lafont F,Burkhardt JK, and Simons K. Involvement of microtubule motors inbasolateral and apical transport in kidney cells. Nature 372:801-803, 1994.
1 I$ g6 s. z6 P8 ]
' B8 r3 _$ l( d: L) n1 _

Meads T andSchroer TA. Polarity and nucleation of microtubules in polarizedepithelial cells. Cell Motil Cytoskeleton 32: 273-288,1995.4 Z2 t$ U4 W* ^3 M  I

) |! Z8 Y- H! a; m
) c8 p* T. _' n1 y$ y9 r7 ]  e
Molitoris BA. Ischemia-induced loss of epithelial polarity:potential role of the actin cytoskeleton. Am J Physiol Renal FluidElectrolyte Physiol 260:F769-F778, 1991.
9 w1 U9 T9 n) ~$ L

Molitoris BA,Falk SA, and Dahl RH. Ischemia-induced loss of epithelial polarity. Roleof the tight junction. J Clin Invest 84: 1334-1339,1989.8 J- d8 L  p% Z

% a+ P& h, u2 ]% U) i* C

Nelson WJ andYeaman C. Protein trafficking in the exocytic pathway of polarizedepithelial cells. Trends Cell Biol 11: 483-486,2001.7 b% G8 Q; p$ ?" C

, r8 D& W. ~% l4 i
Nakagawa T,Sasahara M, Haneda M, Kataoka H, Nakagawa H, Yagi M, Kikkawa R, and HazamaF. Role of PDGF B-chain and PDGF receptors in rat tubular regenerationafter acute injury. Am J Pathol 155: 1689-1699,1999.

0 ^/ {8 H1 w8 \& d, Z( `
Nouwen EJ andDe Broe ME. Human intestinal versus tissue-nonspecific alkalinephosphatase as complementary urinary markers for the proximal tubule. Kidney Int Suppl 47:S43-S51, 1994.

1 O  A, k& f6 n; P9 f& K) _
Oberbauer R,Schwarz C, Regele HM, Hansmann C, Meyer TW, and Mayer G. Regulation ofrenal tubular cell apoptosis and proliferation after ischemic injury to asolitary kidney. J Lab Clin Med 138: 343-351,2001.

$ @. x- s6 ^( \, M" H
# e8 Y6 v! T' J- r. M/ r( K6 z
Ramaswamy D,Corrigan G, Polhemus C, Boothroyd D, Scandling J, Sommer FG, Alfrey E, HigginsJ, Deen WM, Olshen R, and Myers BD. Maintenance and recovery stages ofpostischemic acute renal failure in humans. Am J Physiol RenalPhysiol 282:F271-F280, 2002.) e. t: @* G9 p1 J9 k1 \0 `# g- v  s

- M/ m) c' m) g+ n; _  p
4 r! e: |0 g, e1 R
Reinsch S andKarsenti E. Orientation of spindle axis and distribution of plasmamembrane proteins during cell division in polarized MDCKII cells. JCell Biol 126:1509-1526, 1994.4 L0 k) J  W0 m( _$ v7 w
% @! y9 u) p8 q
  N6 Z4 s% S8 W8 j
. F5 I% t: U: Y5 ~( A( ], h
Rizzolo LJ andJoshi HC. Apical orientation of the microtubule organizing center andassociated gamma-tubulin during the polarization of the retinal pigmentepithelium in vivo. Dev Biol 157: 147-156,1993.

( d8 a# r9 O) ~

Rodriguez ML,Brignoni M, and Salas PJI. A specifically apical sub-membrane intermediatefilament cytoskeleton in non-brush-border epithelial cells. J CellSci 107:3145-3151, 1994.
% P6 _2 Q% Z" y2 `
+ K/ ?7 v* ]' T- @

Rodriguez-Boulan E, Salas PJ, Sargiacomo M, Lisanti M, Lebivic A,Sambuy Y, Vega-Salas D, and Graeve L. Methods to estimate the polarizeddistribution of surface antigens in cultured epithelial cells. Methods Cell Biol 32:37-56, 1989.

) A  V; O+ B) u6 a

Salas PJI. Insoluble -tubulin containing structures are anchored to the apicalnetwork of intermediate filaments in polarized CACO-2 epithelial cells. J Cell Biol 146:645-658, 1999.

. D! U2 |0 x" V5 F  a/ Z: c& ]
9 H$ I) P0 g1 T% f" K. C
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 biogenesisof epithelial cell surface polarity. J Cell Biol 102: 1853-1867,1986.; U5 N! X3 J4 T: [- c3 K

/ J! A. \, o- ]5 O/ T: [

Safirstein R,Di Mari J, Megyesi J, and Price P. Mechanisms of renal repair and survivalfollowing acute injury. Semin Nephrol 18: 519-522,1998.
2 C' t2 |$ X3 s3 k# F
6 F. a3 N' L2 b% ]

Schwartz GJ,Olson J, Kittelberger AM, Matsumoto T, Waheed A, and Sly WS. Postnataldevelopment of carbonic anhydrase IV expression in rabbit kidney. Am J Physiol Renal Physiol 276:F510-F520, 1999.
6 J6 U- F4 z: P* ^
1 q* p4 G5 \, @. P" N$ p7 Z( i7 s2 F

Shimizu A andYamanaka N. Apoptosis and cell desquamation in repair process of ischemictubular necrosis. Virchows Arch B Cell Pathol 65: 171-180,1993.7 `! [2 Z- x! }8 i. `# u: j
. V, g" R6 q& N/ F' n) D) U

Stearns T andKirschner M. In vitro reconstitution of centrosome assembly and function:the central role of -tubulin. Cell 76: 623-637,1994.+ u( q6 c; F  t$ p

Vega-Salas DE,Salas PJI, Gundersen D, and Rodriguez-Boulan E. Formation of the apicalpole of epithelial (Madin-Darby canine kidney) cells: polarity of an apicalprotein is independent of tight junctions while segregation of a basolateralmarker requires cell-cell interactions. J Cell Biol 104: 905-916,1987./ R( N. k4 @% u  B; }1 V
( |* r8 ?/ }' ?$ k! `7 ^) R

Wagner MC andMolitoris BA. Renal epithelial polarity in health and disease. Pediatr Nephrol 13:163-170, 1999.- y) K; x# O3 _- w$ U4 N
$ p# o+ q( v7 P- R

Wiese C andZheng Y. A new function for the -tubulin ring complex as amicrotubule minus-end cap. Nat Cell Biol 2: 358-364,2000.8 N4 s) j" _5 @

. z8 K9 _2 }2 H! N/ c' t/ Q+ a

Witzgall R,Brown D, Schwarz C, and Bonventre JV. Localization of proliferating cellnuclear antigen, vimentin, c-Fos, and clusterin in the postischemic kidney.Evidence for a heterogenous genetic response among nephron segments, and alarge pool of mitotically active and dedifferentiated cells. J ClinInvest 93:2175-2188, 1994.

* L3 D* q. |6 C1 m" A2 i

Yin T, SandhyG, Wolfgang CD, Burrier A, Webb RL, Rigel DF, Hai T, and Whelan J. Tissue-specific pattern of stress kinase activation in ischemic/reperfusedheart and kidney. J Biol Chem 272: 19943-19950,1997.

Yoshida T, TangSS, Hsiao LL, Jensen RV, Ingelfinger JR, and Gullans SR. Global analysisof gene expression in renal ischemia-reperfusion in the mouse. Biochem Biophys Res Commun 291:787-794, 2002.
# n$ ?, ~1 o1 K2 r$ u/ i# ?
$ S) j2 o/ p3 U+ K. w2 ]
, E- S% T' [& Y* a) Q3 a0 @( p
Zuk A,Bonventre JV, Brown D, and Matlin KS. Polarity, integrin and extracellularmatrix dynamics in the postischemic rat kidney. Am J Physiol CellPhysiol 275:C711-C731, 1998.

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