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
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ABSTRACT
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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: 13081315, 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.
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MATERIALS AND METHODS
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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 300450 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-
-tubulin
NH2-terminal synthetic peptide (EEFATEGTDRKDVFFY, Sigma);
monoclonal antibody against
-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 2060% 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).
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RESULTS
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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.

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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.
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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
-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).

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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
(ej). 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
-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 -tubulin signal
separated from the apical domain IFs. Scale bars: 5
µm(ai) and 2 µm(j).
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In Fig. 2, examples of
fields with centrosomes within the same confocal optical section in nearby
cells are shown for control monolayers (ad) and
ATP-depleted cells (ej). As expected from previous
publications, the centrosomes in control cells were observed always embedded
in the apical IF submembrane cytoskeleton
(Fig. 2,
ad). 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
-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
-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
-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-
-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
-tubulin in fractions 1, 3, and 4 and of K19
and
-tubulin in fractions 15
(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
-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.

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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.
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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 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-
-tubulin antibody and FITC-phalloidin to
stain F-actin. In the control proximal tubules, very few 0.3- to 0.4-µm
spots of
-tubulin signal (centrosomes) were easily recognizable and
were mostly localized in the apical pole of the cells. However, a continuous
subapical layer of
-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
-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
-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
-tubulin was observed, even in tubules
(or cells) with a complete brush border
(Fig. 4, c vs.
d, arrows). Instead,
-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
-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.
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
-tubulin complexes (
-TurC; 42) remain in the supernatant and
-tubulin attached to the cytoskeleton in the pellet. In previous
studies, roughly 50% of the
-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
-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
-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
-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.
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Table 1. -Tubulin signal in the cortex of kidneys subjected to I/R as a
percentage of the band from the contralateral (sham surgery) kidney
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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-
-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.

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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- -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.
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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 S23 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).

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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).
eh: Confocal images of the same preparations at
higher magnification. Scale bars: 20 µm (ad) and
10 µm (eh).
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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
|
---|
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 (23 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 (13 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 (58%) 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 25 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.
 |
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