1 Department of Life Sciences, Indiana State University, Terre Haute, Indiana 47809; and 2 Department of Medicine, University of Colorado, Health Sciences Center, Denver, Colorado 80262
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ABSTRACT |
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The present studies investigated acute disruption of microvillar actin cytoskeleton and actin association with other cytoskeletal components in ATP-depleted rabbit proximal tubular cells. Video-enhanced differential-interference contrast microscopy and confocal microscopy were used to follow the fate of F-actin during the disruption of microvilli. Within individual cells, all microvilli collapsed simultaneously. Microvillar actin filaments underwent a parallel decrease in length. Using a sequential cytoskeletal extraction protocol and electron microscopy, we revealed in the present studies the coincident sequestration of a distinct, perinuclear pool of actin that was primarily absent in control cells. Actin sequestration progressed in a duration-dependent manner, occurring as early as 15 min of anoxia when cellular ATP dropped to <5% of control level. Phalloidin staining and depolymerization treatment showed the majority (>90%) of this sequestered actin to be F-actin. A microvillar actin bundling protein villin was also sequestered in the same perinuclear complex of anoxic proximal tubules. In conclusion, the present results demonstrate a coincident microvillar actin bundle disruption and the perinuclear sequestration of F-actin in ATP-depleted proximal tubular cells.
ischemia; ATP depletion; cytoskeleton; kidney; villin
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INTRODUCTION |
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AS A PRIMARY CONSEQUENCE of ischemia, anoxia-induced ATP depletion interrupts many energy-dependent cellular processes, including the maintenance of the structural integrity of the actin cytoskeleton. In renal proximal tubules (PT), actin filaments are concentrated in the microvilli of the apical membrane. These actin bundles in the microvilli play a critical role in supporting the microvillar plasma membrane (4), maintaining the maximized reabsorption surface, and anchoring membrane transporters. During renal ischemia, the disruption of the microvillar actin coincides with the early onset of microvillar blebbing, sloughing, and retraction (8-11, 14, 18, 21) and actin dissociation from basolateral membranes (18).
Alterations in the actin cytoskeleton during ischemia are not
likely due to actin degradation since total cellular actin content remains unchanged (3). Rather, these changes are likely caused by
altered interactions between actin and its binding proteins. A decrease
or increase in actin affinity to its binding proteins will destabilize
the whole structure of the actin cytoskeleton and affect actin
subcellular localization. Indeed, a redistribution of stainable F-actin
from the apical membrane to the perinuclear region following ATP
depletion has been demonstrated by previous immunofluorescence studies,
though mechanisms for the redistribution remain unclear (15).
Biochemical studies have revealed decreased association of actin with a
few actin-binding proteins, including spectrin (17), actin
depolymerizing factor (20), thymosin-B4 (16), and ezrin (3, 5). Actin
dissociation from some of these proteins may initiate redistribution of
actin from microvilli to other subcellular locations. Actin, after
being dissociated from microvilli, seems reassociated with the
cytoskeleton since actin remains in the Triton-insoluble cytoskeletal
fraction (3). The reassociation of actin with the perinuclear
cytoskeleton is likely mediated by an increase in the affinity of actin
with some proteins in that region. Except for an enhanced interaction
between actin and myosin 1 (22), no direct biochemical
evidence has been documented for the enhanced association of actin with
any other cytoskeletal component during ATP depletion.
On the basis of these findings, it was hypothesized that during ATP
depletion, depolymerized or fragmented actin filaments are reassociated
with the cytoskeleton in an abnormal way (Fig. 1). To test this hypothesis, attempts were
made to determine 1) whether actin bundles were disrupted
simultaneously during microvillar retraction and 2) whether any
difference in actin association with the cytoskeleton could be detected
between control (normoxic) and ATP-depleted (anoxic) PT cells. The
results indicate a simultaneous shortening of microvillar F-actin
bundles during microvillar retraction and an enhanced association
(sequestration) of a pool of actin with the perinuclear/nuclear
complex. The sequestration of actin in the perinuclear/nuclear complex
was also initially characterized.
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MATERIALS AND METHODS |
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Preparation of rabbit proximal tubular cells. PTs were isolated and purified as previously described (7). Briefly, female New Zealand White rabbits (1-2 kg; Harlan, Indianapolis, IN) were injected with heparin and euthanized with overdose of pentobarbital sodium (20 mg/100 g rabbit wt). The cortices were trimmed from the excised kidneys, minced, and digested for 60 min at 37°C in DMEM containing 150 U/ml collagenase (Worthington Biochemical, Freehold, NJ) and 1 U/ml DNase (Sigma, St. Louis, MO). The digested cortical tissue suspensions were washed free of collagenase, and PTs were isolated from other cell types by centrifugation on a self-generating 50% Percoll gradient for 30 min at 36,000 g. The collected PT bands were washed three times in DMEM and the final tubule pellets were resuspended in DMEM at a concentration of 2 mg protein/ml. The final PT suspension contains both intact tubules and clusters of PT cells from collagenase digestion.
ATP depletion of isolated PTs. ATP depletion was induced by anoxia, i.e., gassing PT suspensions in DMEM with 100% nitrogen for various duration, or chemical anoxia induced by antimycin A (10 µM), a site II mitochondrial inhibitor, or rotenone (10 µM), a site I mitochondrial inhibitor. Cellular ATP content was measured at 15, 30, 45, and 60 min of anoxia/chemical anoxia using luciferin-luciferase assay as previously described (6). After each experimental treatment, 300 µl of PT suspensions were mixed with an equal volume of 6% perchloric acid, followed by centrifugation at 15,000 g for 1 min. The supernatant is neutralized with 3.8 M KOH, and 10 µl of this sample was incubated for 15 min in 1 ml of luciferin-luciferase buffer containing 20 mg/ml luciferin-luciferase (Sigma), 25 mM glycylglycine, 10 mM MgSO4, and 0.5 mM EDTA, pH 7.5. Photon emission was measured in a Beckman model LS5000 scintillation counter and calibrated with ATP standards. ATP content is expressed as nanomoles per milligram protein and calculated as a percentage of control samples.
Online observation of microvillar retraction during ATP depletion. Freshly isolated rabbit PT suspension was placed in a microscopic perfusion chamber and maintained at 37°C using an air current incubator. Attachment of PT cells to the bottom of the chamber was strengthened with Cell-Tak (Becton-Dickinson Labware, Bedford, MA). Since the brush border lining the lumen of intact tubules was not visible from the outside, the clusters of tubular cells and the cells located at both ends of intact tubules were used to visualize the microvilli. Microvilli at both of the two places were monitored and videotaped via a video-enhanced differential-interference contrast (DIC) microscope (model IM-35 microscope; Zeiss, Oberkochen, Germany) throughout normoxic and anoxic incubation. Chemical anoxia was induced by adding 10 µM antimycin A into the chamber. Once microvillar retraction occurred, the PT cells were fixed immediately by adding 4% paraformaldehyde into the chamber. The fixed PT cells were stained with fluorescein phalloidin (Molecular Probes, Eugene, OR) to visualize F-actin by confocal microscopy (model LSM 410, Zeiss).
Triton solubilization of PTs. The association of actin and other proteins with the cytoskeleton was initially determined by their solubility in Triton X-100. After the experimental period, the PT were pelleted at 100 g for 2 min. The pellets were resuspended in 0.9 ml of ice-cold Triton extraction buffer [0.5% Triton X-100, 300 mM sucrose, 5 mM Tris · HCl, 2 mM EGTA, 200 µM phenylmethylsulfonyl fluoride (PMSF), 10 µM leupeptin, and 10 µM pepstatin, pH 7.4] for 10 min. The extracted suspension was then centrifuged at 4,000 g for 10 min. The Triton-soluble proteins (TSP) and proteins in extracellular medium (EP) were precipitated from the supernatant with 3.5× volume of 100% methanol and pelleted at 4,000 g for 10 min. The Triton-insoluble proteins (TIP), EP, and TSP were resuspended in equal volumes of PAGE buffer (5% SDS, 25% sucrose, 5 mM Tris · HCl, and 5 mM EDTA). The total protein in TIP and TSP fractions of PTs was measured using the bicinchoninic assay (Pierce Chemical). For quantitation of actin and ezrin in the EP, TIP, and TSP fractions, equal volume of each fraction was run on an 8% polyacrylamide gel, transferred to nitrocellulose, and analyzed by Western blotting.
Sequential extraction of PT cells. Sequential extraction of PT cells with a series of buffers was used to dissect the cytoskeleton and reveal any difference in actin association with the cytoskeleton between normal and ATP-depleted PT cells. After having been exposed to normoxic or anoxic incubation, PTs were extracted by digitonin, calcium, Triton X-100, and ATP, sequentially. First, equal amounts of anoxic or normoxic PTs (40 mg protein) were permeabilized with 1 ml of digitonin extraction buffer (0.012% digitonin, 280 mM sucrose, 200 mM KCl, 20 mM HEPES, and 2 mM EGTA, pH 7.4) on ice with agitation for 10 min to extract cytosolic proteins. The digitonin extraction protocol was modified from published methods (19). After digitonin permeabilization, the whole suspension was centrifuged at 4,000 g for 3 min to separate cytosolic proteins (supernatant) from the digitonin-insoluble fraction (pellet). The efficiency of digitonin permeabilization was examined by measuring the representative cytosolic protein lactate dehydrogenase (LDH) in digitonin-soluble and -insoluble fractions. Second, the digitonin-insoluble fraction of normal and ATP-depleted PT was further extracted on ice with agitation for 10 min in 1 ml of calcium extraction buffer (6 mM CaCl2, 10% Triton, 280 mM sucrose, 200 mM KCl, 20 mM HEPES, 2 mM EGTA, 200 µM PMSF, 10 µM leupeptin, and 10 µM pepstatin A, pH 7.4). This calcium extraction was intended to solubilize membrane lipids as well as break the microvillar cytoskeleton, since our previous observation revealed this effect of calcium (unpublished observation). Then, the whole suspension was centrifuged (4,000 g, 3 min) to separate the calcium-soluble from -insoluble fractions. Third, the calcium-insoluble pellet was further extracted in 1 ml of 10% Triton extraction buffer (10% Triton, 280 mM sucrose, 200 mM KCl, 20 mM HEPES, 2 mM EGTA, 200 µM PMSF, 10 µM leupeptin, and 10 µM pepstatin A, pH 7.4). The Triton extraction was designed to remove any proteins that were trapped in the remnant cytoskeleton after calcium extraction. Finally, after separation of the Triton-soluble fraction from the insoluble one, the Triton-insoluble pellet was extracted again in 1 ml of ATP extraction buffer (3 mM ATP, 280 mM sucrose, 200 mM KCl, 20 mM HEPES, and 2 mM EGTA, pH 7.4). The ATP-soluble and -insoluble fractions were separated by centrifugation at 4,000 g for 3 min. After the sequential extraction, five fractions were obtained (Digitonin-S, Calcium-S, Triton-S, ATP-S, and pellet). Each soluble fraction was mixed with equal volume (1 ml) of 2× PAGE buffer, and the final ATP-insoluble pellet was resuspended in 2 ml of 1× PAGE buffer.
Actin content in equal volumes of each of the five fractions was measured by Western blotting. The final pellet fraction was also fixed either in 4% paraformaldehyde for F-actin staining or 2% glutaraldehyde for electron microscopy.Western blotting. Samples were electrophoresed in 8% polyacrylamide gel and transferred to a NitroPure membrane (MSI, Westboro, MA). The blotted proteins were then probed with primary antibodies (monoclonal anti-actin and anti-villin from Chemicon, Temecula, CA; monoclonal anti-ezrin from Immusine, Hayward, CA) and peroxidase-conjugated goat anti-mouse IgG (Jackson ImmunoResearch Laboratories, West Grove, PA). The presence of the antigens was indirectly detected using enhanced chemiluminescence (ECL; Amersham, Arlington Heights, IL) and quantitated by scanning densitometry within linear range by series dilution.
Electron microscopy. In preparation for electron microscopy, samples were fixed with 2% glutaraldehyde in 100 mM NaH2PO4 overnight at 4°C, then dehydrated in ethanol and embedded in Epon 812. Sections were double stained with uranyl acetate and lead citrate and viewed with a Philips model 312 electron microscope.
LDH release. LDH release, an index of plasma membrane integrity, was measured as previously described (3). Extracellular LDH was measured by layering the tubule suspension on 2:1 n-butyl:dioctyl phthalate followed by centrifugation at 14,000 g for 30 s. This procedure pelleted the cellular contents and left the extracellular contents in suspension above the phthalate. Samples of 300 µl of extracellular medium as well as 300 µl of the total suspension were mixed with 60 µl of 2% Triton X-100 before analysis. The LDH activities were converted to percent release (supernatant activity/total activity × 100).
Detection of F-actin in the pellet after sequential extraction. The presence of F-actin in the final pellet of anoxic PT after the sequential extraction was determined with two approaches. First, the pellet was fixed in 4% paraformaldehyde, and stained with fluorescein phalloidin (Molecular Probes) to visualize F-actin. Second, a previously published method (12) was modified to determine whether treating the pellet with an actin depolymerization buffer could solubilize the sequestered actin. Anoxic pellets were incubated in actin depolymerization buffer (2 mM HEPES, 3 mM ATP, 0.2 mM CaCl2, and 0.5 mM dithiothreitol, pH 8.0) at 4°C for 1.5 h. Suspensions were centrifuged at 17,000 g for 35 min to separate soluble actin from cytoskeleton-associated actin. The insoluble pellet and soluble fractions were mixed with 2× SDS-PAGE buffer to an equal volume. Twenty microliters of each fraction were resolved in an 8% SDS polyacrylamide gel, transferred to nitrocellulose, and analyzed for the quantitation of actin by Western blotting.
Data analysis. All data are from at least three independent experiments and are expressed as means ± SE, unless otherwise noted in text. Comparisons between individual groups were made using Student's t-test, with P < 0.05 considered to be statistically significant.
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RESULTS |
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Microvillar retraction and simultaneous shortening of microvillar
actin filaments during chemical anoxia.
Online monitoring of microvilli using video-enhanced DIC microscopy
recorded the temporal sequence of microvillar retraction during
chemical anoxia induced by antimycin A. Microvilli retraction appeared
to be an independent event between individual PT cells. As shown in
Fig. 2, the microvilli on one PT cell
(cell 1) just finished retraction while the microvilli on the
adjacent cell (cell 2) remained unchanged. Most PT cells
underwent microvillar retraction within 20-30 min of chemical
anoxia. However, a few PT cells were able to maintain normal
microvillar morphology during 60 min of anoxia. Within each individual
PT cell, all microvilli retracted simultaneously. The duration from the
beginning to the completion of the microvillar retraction varied from
30 s to 2 min. The tips of microvilli in some cells enlarged right
before microvillar retraction. Because of both the limited focusing
depth of DIC microscopy and the motion of microvilli and cell bodies during microvillar retraction, it is difficult to obtain
high-resolution DIC images during retraction. Alternatively, the
enlarged microvillar tips of ATP-depleted cells were documented in the
electron micrograph in Fig. 3.
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No increase in Triton-soluble actin during ATP depletion.
Triton X-100 solubility has been widely used as a measure of
cytoskeleton association (3, 16, 18). PT cells were exposed to normoxia
or anoxia, then separated into TIP and TSP fractions. In addition, the
extracellular medium was collected to quantify the release of EP by PT
cells during anoxia. Despite the shortening of the microvillar actin
filaments during anoxia, no significant changes (P < 0.05) in
actin Triton solubility were observed (Fig. 4). The percentage of actin in TIP, TSP,
and EP was 79 ± 4%, 19 ± 3%, and 1 ± 1% under normoxia and 73 ± 5%, 26 ± 6%, and 4 ± 2% under anoxia, respectively. In
contrast to actin, another microvillar cytoskeletal protein, ezrin,
exhibited a significant increase in Triton solubility during anoxia
(Fig. 4). Ezrin dissociated from the cytoskeleton as indicated by the
decrease in TIP ezrin from 90 ± 2% to 40 ± 4% (P < 0.01, n = 5) and the increase in TSP and EP ezrin from 10 ± 2% to 26 ± 3% and 1 ± 1% to 34 ± 5% (P < 0.01, n = 5), respectively. The above morphological
evidence, immunocytochemical studies of actin filaments, and the
dissociation of ezrin clearly indicate the breakdown of microvillar
cytoskeleton and actin bundles. The unchanged Triton solubility of
actin suggests that fragments of actin filaments or depolymerized actin
monomers are reassociated with the insoluble cellular components.
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Differential association of actin with insoluble cellular components
in control and anoxic PT cells.
PT cells, after being exposed to normoxia or anoxia, were sequentially
extracted by digitonin, calcium, Triton, and ATP extraction buffers to
experimentally manipulate the interactions between actin and the
insoluble cellular components. Anoxic and normoxic PT membranes were
first permeabilized with digitonin buffer to solubilize free cytosolic
proteins. LDH release by control cells after digitonin treatment
demonstrated efficient permeabilization, with >90% of total LDH
activity in the supernatant. The total actin content before and after
the sequential extraction did not change significantly (data not
shown), indicating no degradation of actin during the extraction. After
sequential extractions, almost all actin in normal PT cells was
solubilized, with little or no actin left in the final pellet (Fig.
5). However, a significant amount of actin
in anoxic PT cells remained in the insoluble cellular components (Fig.
5). The sequestration of actin in the insoluble cellular components was
also observed during ATP depletion induced by inhibiting the
mitochondrial respiratory chain with chemical uncouplers (data not
shown). These results indicate an abnormally enhanced interaction
(sequestration) between actin and the insoluble cellular components
during ATP depletion. The sequestration of actin occurred as early as
15 min of anoxia and reached a plateau at 45 min of anoxia (Fig.
6A). The parallel measurement of
cellular ATP content during anoxia indicated that cellular ATP dropped to <5% of control level (Fig. 6B) when actin sequestration
occurred at 15 min of anoxia.
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Electron micrograph of the insoluble cellular components that
sequester actin.
Electron microscopy studies revealed that the final insoluble cellular
fractions obtained after sequential extraction of normoxic and anoxic
PT cells were a perinuclear remnant cytoskeleton/nucleus complex (Fig.
7). No apparent microvilli or terminal web
structure were found in the insoluble mass, and no morphological
difference was visible between normoxic and anoxic pellets (Fig. 7).
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F-actin sequestered in the insoluble cellular components.
Immunocytochemistry studies using fluorescent phalloidin revealed
F-actin in the insoluble cellular components (Fig.
8A). F-actin was found in the
perinuclear region but not inside the nuclei (Fig. 8A).
Furthermore, actin depolymerization treatment (see MATERIALS AND
METHODS) solubilized 91 ± 2% (mean ± SD, n = 5) of
the sequestered actin in the final insoluble cellular components in
anoxic PT cells, indicating the presence of F-actin (Fig. 8B).
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Copresence of villin in the insoluble cellular components.
Western blotting with antibody against villin, a microvillar marker
protein, revealed the presence of villin in the insoluble cellular
components that sequestered actin in anoxic PT cells (Fig.
9). Since villin is mainly localized to the
microvilli of PT cells, appearance of villin in the insoluble cellular
components suggests that microvillar cytoskeletal components are
sequestered in the perinuclear remnant cytoskeleton/nucleus complex
during anoxia.
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DISCUSSION |
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The results from the present studies provide novel information on microvillar retraction and an enhanced interaction of actin with the perinuclear complex during ATP depletion.
Observation of the dynamic process of microvillar retraction. Direct online observation of microvilli during ATP depletion using video-enhanced DIC microscopy, for the first time, revealed the entire process of microvillar retraction as a swift all-or-none event in each individual PT cell (Fig. 2). The simultaneous retraction of all microvilli in a PT cell during ATP depletion suggests that some critical threshold of cellular ATP concentrations initiates microvilli retraction. The retraction seems initiated at the tips of the microvilli, because enlargement of the tips occurred often right before microvillar retraction (Fig. 3). Studies on F-actin immunofluorescence revealed that microvillar actin filaments were shortened at the end of microvillar retraction (Fig. 2C). Thus microvillar retraction is more likely caused by disruption of actin bundles in microvilli than by intrusion of microvillar actin bundles into terminal web or the shrinkage of terminal web.
An abnormal actin interaction with the perinuclear cytoskeleton. The loss of microvillar F-actin did not result in a proportional increase in free cytosolic Triton-soluble actin (Fig. 4). These results, consistent with previous findings (3, 10, 11, 14, 21), suggest that fragments of the actin cytoskeleton in the microvilli are reassociated with the cytoskeleton. Indeed, an abnormally enhanced association of an actin pool with the perinuclear cytoskeleton/nucleus complex was identified after further dissecting the cytoskeleton in anoxic PT cells.
The sequential extraction employed in the present studies solubilized nearly all actin in normal PT cells but not in anoxic cells (Fig. 5). The pool of sequestered actin in anoxic PT cells must be interacting with the insoluble cellular components in a way different from those in normal cells. Isolation of this pool of actin and the associated insoluble cellular components makes it possible to further explore molecular details of the sequestration of actin, as well as provide information on the mechanisms of actin cytoskeleton breakdown in ATP-depleted PT. Electron microscopy studies identified the pellet that sequestered actin as remnant perinuclear cytoskeleton and nuclei (Fig. 7). No visible microvilli or terminal web was found in the pellet. Therefore, the present biochemical and morphological evidence suggests that fragments of actin filaments are sequestered at the perinuclear/nuclear region. These findings are consistent with the aggregation of actin in perinuclear region revealed by immunostaining of actin in cultured cells during ATP depletion (1, 11, 15).F-actin sequestered in the perinuclear cytoskeleton. Phalloidin staining revealed a high abundance of F-actin in the perinuclear site of the insoluble pellet in anoxic PT, which was absent in normoxic controls (Fig. 8A). Furthermore, 91% of the sequestered actin in the perinuclear/nuclear complex of anoxic PT cells was solubilized by actin depolymerization treatment (12) (Fig. 8B). The finding of F-actin sequestration may provide clues about the disruption of microvillar actin cytoskeleton. Although depolymerization of F-actin bundles cannot be excluded, the evidence of sequestered F-actin would implicate that the disruption of the microvillar actin cytoskeleton is caused, at least in part, by severing of actin bundles in ATP-depleted PT. Indeed, distinct pools of subcellular actin may be affected differently during ischemic PT injury, but further investigation is required to completely elucidate the mechanism of actin cytoskeleton breakdown.
The origin of the sequestered actin was initially sought considering that the sequestered fragments of microvillar cytoskeleton may contain not only actin but also other microvillar proteins such as villin. Indeed, the microvillar cytoskeletal marker protein villin (2) was also found in the perinuclear/nuclear complex (Fig. 9). It is possible that the actin bundling protein villin was associated with fragmented microvillar actin bundles and sequestered together with actin. It is also likely that villin is involved in the severing of actin bundles during anoxia, because the actin-severing capability of villin has been shown by in vitro experiments (2, 13). The relationship between villin and actin in the perinuclear complex is currently under investigation to determine whether villin is directly bound to actin. In summary, these findings demonstrate that actin is sequestered in a perinuclear complex during ATP depletion in a time-dependent manner after microvillar retraction and microvillar F-actin shortening. Initial characterization of the sequestered actin presented clues that may lead to the identification of the actin-sequestering protein(s) and the origin of the sequestered actin. The pathological significance of sequestering actin during ATP depletion is not yet clear. The sequestration of critical cytoskeletal proteins such as actin may be a mechanism to protect these proteins from being destroyed by proteolysis so that they can be recruited back during recovery. The method of sequential extraction for isolation of an actin-sequestering perinuclear complex will allow for more detailed investigations of the molecular mechanism involved in actin cytoskeleton alterations during ischemia. ![]() |
ACKNOWLEDGEMENTS |
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This work was partially supported by American Liver Foundation Grant ALF PN-9801-0141 (to R. B. Doctor).
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FOOTNOTES |
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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. §1734 solely to indicate this fact.
Address for reprint requests and other correspondence: J. Chen, Dept. of Life Sciences, Indiana State Univ., Terre Haute, IN 47809 (E-mail: lschen{at}scifac.indstate.edu).
Received 13 July 1999; accepted in final form 12 January 2000.
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