Ischemia disrupts myosin I
in renal
tubules
Judy
Boyd-White1,
Anjaiah
Srirangam1,
Michael P.
Goheen2, and
Mark
C.
Wagner1
1 Renal Epithelial Biology Experimental Laboratories,
Division of Nephrology, Department of Medicine, and 2 Department
of Pathology and Laboratory Medicine, Indiana University School
of Medicine, Roudebush Veterans Affairs Medical Center,
Indianapolis, Indiana 46202
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ABSTRACT |
In these
studies we have examined rat kidneys biochemically and microscopically
to determine where myosin I
is located before, during, and after an
acute ischemic injury. Myosin I
is present in multiple
tubule segments including the brush border (BB) of the proximal tubule
cell (PTC). Its distribution is severely altered by a 15-min renal
artery clamp. Myosin I
is present in the urine during reflow and is
found in the numerous cellular blebs arising from the damaged PTC and
other tubules. Two hours of reflow result in a decrease in BB myosin
I
staining and an increase in its cytoplasmic staining.
Interestingly, the return of the F-actin in the BB precedes the return
of the myosin I
, suggesting that this myosin I isoform may not play
a role in rebuilding the microvilli after an ischemic injury. A
nonstructural role for this myosin, such as transport or channel
regulation, is supported by its presence in many tubule segments, all
of which have transport and channel requirements but do not all contain microvilli.
kidney; cytoskeleton; motor; disease; actin
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INTRODUCTION |
MYOSIN
I
is a member of the myosin superfamily that contains at least 15 distinct classes (33). All myosins contain an actin and
ATP binding site in their globular head or motor region, while the
myosin tail regions are quite diverse. Many myosin class members have
been identified by molecular techniques in both lower and higher
eukaryotes (23, 33, 40). In addition, multiple studies
have identified multiple myosins in the same tissue and/or cells,
suggesting that these molecular motors may coordinate their respective
or redundant roles in cells. The myosin I class members have three
functional domains (3, 12). At the amino terminus is a 70- to 80-kDa region that contains the ATP and actin binding site. The
-helical neck region contains one to six IQ (isoleucine-glutamine) motifs that interact with light chains that are calmodulin in vertebrates. The carboxy end domain, ~20 kDa, is enriched in basic amino acids that interact with acidic phospholipids. Numerous studies
have proposed a role for myosin I's in cell motility, endocytosis,
secretion, contractile function, and Golgi vesicle trafficking and in
linking the actin bundles of microvilli to the plasma membrane. All
these activities involve an interaction with membrane and actin
filaments, a dual property for which myosin I's are well equipped.
There are at least four distinct classes of myosin I that are
widespread in mammalian tissues, including kidney (12, 23, 33,
40). The four classes have been designated as I
, I
, IC,
and I
, whereas in rat they are termed myr1, myr2, myr3, and myr4
(3, 33). The human cDNA for myosin I
also has been characterized and is called Myo1c (33). Myosin I
was
the first mammalian myosin I purified (2). Western blots
and immunofluorescence have shown this isoform to be widely
distributed, being found in all rat tissues examined and present in
active motile regions of cells, i.e., lamellipodia and ruffles
(1, 12, 30, 36, 40). In addition, a punctate cytoplasmic
pattern is observed in both cell and tissue immunolabeling. Recent
studies suggest that myosin I may be involved in the regulation of
mechanochemical transduction channels in the apical region of hair cell
stereocilia in the inner ear (16, 18). We have previously
localized myosin I
to microvilli in a proximal tubule cell line,
LLCPK, and have shown that it was also present in the brush border (BB)
of proximal tubules (37).
Proximal tubule cells (PTC) have a specialized apical membrane that
mediates the selective and efficient reabsorption of ions, water, and
macromolecules from the glomerular filtrate (6, 24).
Reabsorption requires microvilli and delivery of ion channels, receptors, and regulatory molecules to their correct destination, thus
enabling the establishment and maintenance of surface membrane polarity. These processes depend, in part, on actin-surface membrane interactions and vesicular transport, both processes in which myosin I
has been implicated. The apical region of the PTC is a very dynamic
region composed of the terminal web and microvilli, each containing a
high concentration of actin and lacking microtubules. The pit area at
the base of the microvilli is a site of active endocytosis, and myosin
I
has been identified in an endosome fraction from this region
(17). Importantly, membrane channels and other proteins
and lipids are continuously delivered to the plasma membrane of all
kidney tubules to replenish and maintain a cellular environment capable
of executing urinary function. Ischemia in vivo or ATP
depletion in vitro severely alters PTC actin cytoskeleton and inhibits
ATP-dependent vectorial transport of organelles (4, 13, 24,
35). Data from numerous labs have documented that
ischemia-reperfusion results in cellular changes throughout the
kidney, including proximal and distal tubules (4, 13-15, 19,
21, 24, 25, 29). The alterations combined lead to cellular and
organ-level dysfunction. Return to the normal physiological state
requires cellular events involving both the actin cytoskeleton and
vectorial transport.
We previously documented the changes to myosin I
in an established,
reversible in vitro ischemic model, LLCPK ATP depleted/repleted (37). In that study, myosin I
was shown for the first
time to be present in microvilli, to undergo apparent cross-linking with actin after ATP depletion, and to return to a normal distribution after ATP repletion. Myosin I
also was observed in the cytoplasm and
at the lateral cell borders of cells. If this unconventional myosin
participates in the structural stability of the microvilli, its
presence in the BB should coincide with the return of the BB after
ischemia. However, a return after BB formation would suggest
that it is participating in some other cellular function in these
cells. Consequently, the objective of these studies was to analyze the
distribution of myosin I
in the rat kidney under normal
physiological conditions, during ischemia, and after
reperfusion following ischemia. Interestingly, we found myosin
I
in many tubule segments. We found that myosin I
is indeed
concentrated in the BB of PTCs but also is present in a punctate
pattern in the cytoplasm. Ischemia dramatically altered the BB
morphology and resulted in a loss of myosin I
from the BB and an
apparent dissociation from F-actin. However, Western blot analysis of
both kidney and urine samples from the injury and recovery process shows minimal loss of myosin I
. This finding suggests that the changes we observed by immunofluorescence are the result of dramatic redistribution and/or reorganization of myosin I
in the proximal tubule. Fifteen minutes of ischemia followed by 2 h of
recovery resulted in structurally normal microvilli. However, myosin
I
, detected by immunofluorescence, had not returned to its
predominant BB location in the proximal tubules but was found
predominantly in the cytoplasm. Even after 24 h of recovery we
observed more cytoplasmic and less BB staining of myosin I
. These
studies suggest that myosin I
does not play a structural role in the
early stages of microvilli rebuilding. In fact, its presence in all
kidney tubules suggests a more ubiquitous role for myosin I
,
possibly in vesicular transport or other cellular functions requiring
membrane-actin interactions.
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MATERIALS AND METHODS |
Antibodies.
The primary antibodies against myosin I
have been characterized and
used in multiple studies (1, 9, 36, 37). Briefly, highly
purified adrenal myosin I
was the antigen used, and the resulting
mouse monoclonal antibodies react with a specific band in the rat
kidney (Fig. 1). M2 and M3 have been
mapped to a tail region specific for this myosin I isoform, and M4 and
M5 have been mapped to a head domain region containing the actin and
ATP binding sites (36, 37). All antibodies give similar
immunolabeling results, while M2 and M3 have an apparent higher
affinity for SDS-denatured myosin I
and are routinely used for
Western blots. The actin antibody was from Chemicon (no.
MAB1501R; Temecula, CA).

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Fig. 1.
Myosin I is present throughout the kidney.
A: specificity of the antibodies used. Three samples were
blotted: rat kidney cortex (C), medulla (M), and partially purified
brush-border membranes (BBM, or B). Ten micrograms of SDS-solubilized
cortex (lanes 1 and 4), medulla (lanes
2 and 5), and BBM (lanes 3 and 6)
were loaded. Lanes 1-3 were stained with Gel Code
Silver Snap stain. Lanes 4-6 were Western blotted for
myosin I and actin. M2 and M5 monoclonal antibodies against myosin
I were used. Note the enrichment of the actin in the BBM sample.
B and C: BBM, cortex, or medulla samples were
extracted in <500 µl with 1% Triton X-100 (TX-100) plus (in mM) 250 sucrose, 6 PIPES, 2.5 HEPES, 0.2 EGTA, and 0.2 MgCl2, pH
6.9, on ice for 10 min. The solubilized proteins were separated from
the insoluble by centrifugation at 48,000 g for 30 min.
Equal amounts of cortex and medulla protein were extracted, and pellets
were resuspended to volume. Equal volumes were then loaded on the gel.
The BBM sample used for extraction contained approximately one-sixth
the amount of protein. S, supernatant; P, pellet.
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Western blot analysis.
SDS-PAGE was carried out using Bio-Rad Criterion Tris-HCl gels
(Hercules, CA) or Fisher PAGEr gold precast gels (Hanover Park, IL).
Prestained molecular mass standards from Sigma (St. Louis, MO) and
Bio-Rad were used. For immunoblotting, samples were resolved by
SDS-PAGE and transferred by electroblotting to Millipore polyvinylidene difluoride (Bedford, MA). The chemiluminescence detection method used
either the alkaline phosphatase substrate CSPD (Tropix) or Pierce's
ECL kit according to the manufacturer's instructions. Gels were
stained with either Gel-Code Blue (Pierce) or Gel-Code SilverSnap stain (Pierce).
Northern blot analysis.
At the specified time of ischemia and/or reperfusion, the
kidneys were briefly (<20 s) perfused with PBS, and the cortex, medulla, and papilla were separated and rapidly frozen in liquid nitrogen. Frozen tissue was homogenized with the Bio-Pulverizer (Biospec Products, Bartlesville, OK) in liquid nitrogen, and powdered tissue was then poured into a tube containing the TRI reagent (Molecular Research Center, Cincinnati, OH). Total RNA was prepared according to the manufacturer's protocol. For Northern blots, total
RNA was electrophoresed in 1% agarose gels containing 1.85% formaldehyde. Equal loading was assessed by optical density reading of
samples before loading and by analysis of 28S RNA bands after the gel
run. The RNA was transferred onto nylon membranes using the Schleicher
and Schuell Turbo blotter (Keene, NH). The membranes were prehybridized
for 15 min and hybridized for 2 h at 65°C using Rapid-hyb buffer
(Amersham). The probe was a 700-bp myosin I
cDNA tail fragment
labeled using the Rediprime DNA labeling system (Amersham). The filter
was washed twice with 2× sodium saline citrate (SSC)-0.1% SDS at room
temperature for 5 min each time, once for 15 min with 0.5× SSC-0.1%
SDS at 65°C, and once for 15 min with 0.2× SSC-0.1% SDS at 65°C.
Analysis and quantitation were conducted with a Molecular Dynamics
PhosphorImager. All analyses compared the clamped kidney with the
contralateral unclamped kidney. RNA levels of the paired kidneys were
normalized using a glyceraldehyde-3-phosphate dehydrogenase probe. Data
are presented as means ± SE. Statistical analysis was performed
with the ANOVA and Tukey's honestly significant difference analysis,
and a P value <0.05 was considered statistically significant.
Immunofluorescence of rat kidney sections.
Rat kidneys were perfusion-fixed according to the procedure of
Maunsbach and Afzelius (22). Briefly, the animals
were anesthetized with an intraperitoneal injection of
ketamine-xylazine and atropine. The abdominal aorta and viscera were
exposed via a midline incision, and sutures were placed superior and
inferior to the renal blood vessels to confine the perfusate flow
primarily to the kidneys. A catheter was secured in the abdominal
aorta in the retrograde direction for delivery of a prefixation rinse
of 20 ml of warmed and filtered 0.85% saline containing 3 U/ml of
heparin, followed by the fixation solution containing filtered 2%
paraformaldehyde in PBS, pH 7.4, warmed to 37°C. The kidneys were
perfusion-fixed for 5 min at a flow rate of 50 ml/min at an average
pressure of 140 mmHg. After perfusion fixation, the kidneys were
removed from the animal, the capsule and perirenal fat were dissected
away, and the tissue was trimmed for immersion fixation at 4°C.
Larger blocks were sectioned on a Vibratome for immunofluorescence, and smaller blocks ~1 mm3 were processed for electron
microscopy (EM). Sections 50-100 µm thick were obtained with the
use of the Vibratome, and the sections were placed in 0.20%
paraformaldehyde until labeled. For immunolabeling, the sections were
rinsed with PBS and placed into 1% SDS or 1% Triton X-100 for 5 min
(5). After detergent treatment, the tissue sections were
rinsed in PBS and placed in PBS-50 mM EGTA containing the primary
antibody (10 µg/ml). Incubation was at room temperature overnight,
with gentle agitation. The sections were extensively (minimum 6× 30 min) washed with PBS at room temperature, followed by an overnight
incubation with the secondary antibody (goat anti-mouse IgG conjugated
to Cy5, Jackson ImmunoResearch, or Alexa 568, Molecular Probes, Eugene, OR) 5 µg/ml in 3% BSA in PBS. F-actin was labeled by including Alexa
488 or Oregon Green phalloidin (Molecular Probes) diluted 1:200 with
the secondary antibody. FITC-labeled lectins were used at a
concentration of 10 µg/ml. The sections were again rinsed with PBS
and mounted on coverslips using ProLong (Molecular Probes). Immunofluorescence controls included omission of the primary antibody, use of multiple secondary antibodies that gave the same pattern for
myosin I
, and use of several additional irrelevant monoclonal antibodies at the same concentration as the myosin I
antibodies. Omission of both the primary and irrelevant antibodies resulted in
negligible staining.
A Bio-Rad (Hercules, CA) MRC 1024 laser scanning confocal microscope
was used to acquire images. Focal sections were obtained in 0.5- or
1-µm steps with Kalman averaging. Each black-and-white image
presented is a single focal plane unless stated otherwise. Three-dimensional (3-D) reconstruction of a series of images was performed with the use of MetaMorph's (Universal Imaging, West Chester, PA) stack 3-D reconstruction function. All digital images were
imported into MetaMorph, and image planes were selected and transferred
to PowerPoint for labeling and scaling. Color encoding was performed in
MetaMorph by assigning the red and blue channels to the myosin I
signal and the green channel to the F-actin signal. The resulting
images show myosin I
in purple and F-actin in green, and
colocalization is shown as white.
Induction of renal ischemia and urine collection.
In Sprague-Dawley rats utilized for microscopy and RNA isolation, renal
ischemia was induced by clamping of the renal pedicle of the
left kidney, with the contralateral kidney serving as the paired
control (26). The rats used for urine and total protein changes were subjected to a bilateral renal clamp. We found no difference in the morphological changes and distribution of myosin I
and F-actin between the 15-min unilateral and bilateral clamp models.
Rats were acclimated to metabolic cages for 2-3 days before the
bilateral clamp was performed. Urine was collected 24 h before surgery and after 2 and 24 h of reperfusion following the 15-min clamp. Urine volume was measured and monitored using Bayer Multistix for urinalysis. The urine was centrifuged for 15 min at 17,000 g, and the supernatant was assayed. Kidneys were removed at
2 or 24 h after reperfusion and separated into cortex and medulla (including papilla and pelvis). Tissue was solubilized with SDS. These
protein and urine samples were used for semiquantitative analysis of
myosin I
changes after the ischemic injury (see Fig. 4).
Other methods.
Tissue for EM analysis was obtained as described in
Immunofluorescence of rat kidney sections, with the tissue
blocks processed according to the procedure of Maunsbach and Afzelius
(22). Briefly, blocks were rinsed in PBS twice for 30 min
to remove fixative, postfixed in 1% osmium tetraoxide for 1 h at
4°C, rinsed in PBS twice for 30 min, dehydrated through an ethanol
series, and embedded in Spurrs resin. Thick sections (0.5 µm) were
cut on an RMC (Research Manufacturing Company, Tucson, AZ) microtome
and stained with 1% toluidine blue for light microscopy. Thin sections
(90-120 nm) were cut and stained with uranyl acetate followed by
lead citrate and were viewed with a Philips CM 10 electron microscope. Protein concentration was determined using Pierce's BCA Protein Assay
Reagent or Bio-Rad's Protein Assay.
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RESULTS |
Myosin I
is widely distributed in the rat kidney.
We previously showed that myosin I
was present in the microvilli of
LLCPK1 PTCs and that its distribution was altered by ATP depletion
(37). The primary objective in these studies was to
determine the distribution of myosin I
in the rat kidney under normal physiological conditions, after an ischemic insult, and during reperfusion. Figure 1A shows the specificity of the
antibodies used in these studies. Western blot analysis of rat kidney
cortex, medulla, and BB membranes (BBM) is shown.
Most myosin I
in rat kidneys is Triton X-100 insoluble in contrast
to that in LLCPK cells, which under control conditions is close to 90%
soluble (37). Figure 1, B and C,
shows the results from a Triton X-100 extraction of BBM, cortex, and
medulla. After extraction on ice, the sample was centrifuged,
and the supernatant and pellet were analyzed by silver stain (Fig.
1B) and Western blot (Fig. 1C) for myosin I
.
Note that the majority of myosin I
is present in the
F-actin-containing pellet in all samples. Over 90% of myosin I
was
found in the supernatant when ATP was added before centrifugation (data
not shown). This finding agrees with our data from the in vitro
ATP-depletion model. We observed no significant change in the Triton
X-100 solubility of myosin I
after ischemia or reperfusion
in rat kidney (data not shown). This is likely due to the highly
organized and developed cytoskeleton in vivo compared with the in vitro model.
Immunofluorescence images in Figs. 2 and
3 show that myosin I
is widely
distributed in the rat kidney. In Fig. 2, three rat kidney Vibratome
sections double-labeled for myosin I
(A, C, and E) and F-actin (B, D, and
F) are shown. Figure 2, A and B, represent single planes, while Fig. 2, C-F, are
reconstructions from multiple confocal planes. Note the prominent
myosin I
BB labeling in the proximal tubules. The intense labeling
of F-actin with phalloidin easily identifies proximal tubules. We often
observed a zone of reduced myosin I
labeling under the BB that may
correspond to the terminal web. There is also distinct cytoplasmic
myosin I
staining that appears rather punctate in many of the
tubules. In Fig. 3, sections were double-labeled for myosin I
(A and C) and peanut lectin (B and
D) to address the presence of myosin I
in non-PTCs.
Peanut lectin labels primarily the intercalated cells in the rat
kidney, though some staining has been observed in the inner medulla
(7, 20). Myosin I
is clearly present throughout the
cytoplasm of collecting duct cells and is often concentrated near the
apical surface of these cells. This distribution, along with its
presence in the BB of PTCs, is consistent with a role for myosin I
that involves regulating the dynamic actin network found near the
plasma membrane in these cells (6).

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Fig. 2.
Myosin I is concentrated in the brush border (BB) of
proximal tubule cells (PTC). Vibratome sections of rat kidney were
double-labeled for myosin I and F-actin. A, C,
and E show low- and high-power views of myosin I labeling
in primarily proximal tubules (PT), whereas B, D,
and F show F-actin labeling in the same tubule field.
C and D are reconstructions from 24 0.5-µm
planes, and E and F are reconstructions from 19 0.5-µm planes. Note the prominent staining of myosin I in the BB,
the frequent weak staining zone under the BB, and consistent but weaker
cytoplasmic staining. Bars = 10 µm.
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Fig. 3.
Presence of myosin I in nonproximal tubules. Vibratome sections
of rat kidney were immunolabeled for myosin I and peanut lectin.
A and C show the myosin I signal, whereas
B and D show the peanut lectin labeling in the
same tubule field. Note the presence of myosin I in the non-PTC,
i.e., peanut lectin-positive tubules in addition to PTC. Bars = 10 µm.
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Changes induced by ischemia.
Ischemia has been shown to alter many cellular proteins and
functions and has been most extensively studied in the PTCs (4, 24, 35). Our objective in these studies was to assess changes to
myosin I
caused by ischemia and to determine when myosin
I
returns to normal. We chose to use a 15-min ischemic clamp
because structural alterations to the proximal tubules have been
documented to follow this period of ischemia (26, 38,
39). Furthermore, this brief ischemic injury will result
in less cell death and enable a quicker return to normal cell structure
and function.
The first analysis addressed whether ischemia-reperfusion
altered levels of myosin I
mRNA. Quantitation of Northern blots, shown in Fig.
4A, showed
changes to myosin I
expression in both cortex and medulla after a
15-min clamp and different times of recovery. Note that after 2 and
4 h of reperfusion, there was an increase in expression of myosin
I
mRNA in both the cortex and medulla. After 24 h there was a
significant decrease in myosin I
expression in the medulla that did
not return to normal levels till after 3 days of reperfusion. Myosin
I
mRNA in the cortex returned to normal levels within 3 days of
reperfusion. These data suggest that myosin I
mRNA expression is
regulated differently in cortex and medullary cells and that its
recovery from an ischemic injury occurs more slowly in the
medullary cells. An increase in mRNA expression could be caused by the
need to replace myosin I
that has been lost. Western blot analysis
of kidney proteins (Fig. 4B) was conducted to determine
whether myosin I
protein levels change after an ischemic
injury. Quantitation using the UN-SCAN-IT program (Silk Scientific,
Orem, Utah) on scanned blots showed that changes between control and
recovery time points varied <10%. In addition, urine was collected
and analyzed for its protein content to further evaluate myosin
I
release. Figure 4C shows the changes in urine protein
before and after the ischemic clamp. Note that the urine output
(ml/h) is similar under all conditions. To evaluate the urine protein
content of the control and 15-min ischemia-24-h reperfusion
collections, they were concentrated fivefold. The 15-min
ischemia-2-h reperfusion sample was not concentrated. The most
notable difference besides the increase in albumin after ischemia was the minor increase in the release of myosin I
and actin at the 15-min ischemia-2-h reperfusion collection.

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Fig. 4.
Ischemia causes minor alterations to kidney
myosin I mRNA and protein levels. A: Northern analysis of
myosin I mRNA levels in both cortex and medulla is presented at
selected time points of ischemia-reperfusion. All analyses
compared the clamped kidney with the contralateral unclamped kidney.
For each time point, a minimum of 3 samples was analyzed. Data are
presented as means ± SE. Cortex and medulla values were combined
to perform statistical analysis using ANOVA and the Tukey's honestly
significant difference test. The variable tested was recovery time.
*Significant differences (P < 0.05) were found between
2 and 4 h of recovery compared with the 24-h recovery time point.
B: a Gel-Code Blue-stained gel and Western blot of 2 identical gels that contained kidney cortex and medulla samples under
control conditions, after 15-min ischemia and 2-h recovery
(15-2 h), or after 15-min ischemia and 24-h recovery
(15-24 h). Total protein (50 µg) was loaded in each lane. The
gel for Western blotting was cut and probed for myosin I or actin
(A), and their respective bands are shown below the Gel-Code
Blue-stained gel. C: a Gel-Code Blue-stained gel and Western
blot of rat urine samples (lanes 1-8) under control
(before clamping), after 15-min ischemia and 2-h recovery
(15-2 h), or after 15-min ischemia and 24-h recovery
(15-24 h). Results from identical gels blotted for myosin I and
actin are presented below the stained gel. The control and 15-24 h
urine samples were concentrated 5-fold because of their low protein
levels. Concentrated urine (5 µl) was loaded for the control and
15-24 h samples, and 5 µl of collected urine was loaded for the
15-2 h collection.
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Figure 5 shows two rat kidney sections
double-labeled for myosin I
(A and C) and
F-actin (B and D) after 15 min of
ischemia. Note the presence of myosin I
closer to the apical
surface and its presence in cellular blebs (Fig. 5C,
arrows). These alterations were observed in tubules containing distinct
F-actin labeling, i.e., proximal tubules, and also in those without
distinct F-actin labeling, i.e., nonproximal tubules (Fig.
5A, star). The changes in location of myosin I
are not
restricted to proximal tubules and suggest that recovery will require
reorganization and retargeting of this protein.

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Fig. 5.
Cellular redistribution of myosin I following ischemia.
Vibratome sections from ischemic kidneys were double-labeled
for myosin I (A and C) and F-actin
(B and D). Note the presence of myosin I in
cellular blebs from both PTC (C, arrows) and non-PTC
(A, star). Myosin I was also found in the tubule lumens.
C and D are single confocal planes, and
A and B are reconstructions from 14 1-µm
confocal planes. Bars = 10 µm.
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Recovery from ischemia: Does myosin I
participate in rebuilding the microvilli?
Having established that myosin I
undergoes dramatic changes with
ischemia, our next objective was to establish the structural relationship between the reappearance of microvilli and the return of
myosin I
to the BB. Figure 6 shows the
overall cell structure of primarily proximal tubules at both the light
and EM level. While there are clearly some areas lacking complete
microvilli that can be found after only 2 h of reperfusion (Fig.
6F, arrow), the majority of the proximal tubules appear to
have reformed normal BB with microvilli.

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Fig. 6.
Effect of ischemia-reperfusion on PT morphology. The
overall cell structure of PTs is shown at both the light level, stained
with toluidine blue (Tol. Blue; A-C), and at the
electron microscopy (EM) level (D-F) under control
conditions (A and D), after 15 min of
ischemia (B and E), and after 15 min of
ischemia followed by 2 h of reperfusion (C and
F). Asterisks identify PT lumens. Note the prominent BB and
microvilli in A and D, their dramatic change in
B and E, and their structural return by 2 h
of reperfusion in C and F. The arrow in
F identifies a PT with incomplete recovery of its
microvilli. Bars = 10 µm.
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Figure 7 highlights the major
observations reported for myosin I
under control (A) and
ischemic (B) conditions and shows the location of
myosin I
after 15 min of ischemia and 2 (C-F) or 24 h (H and I) of
reperfusion. Color images present myosin I
in purple and F-actin in
green. Colocalization between myosin I
and F-actin is indicated as
white. Note the prominent BB labeling for both F-actin and myosin I
in the control state (Fig. 7A), where significant white is
indicated, and the dramatic disruption to both myosin I
and F-actin
after ischemia (Fig. 7B). With 2 h of
reperfusion, a modest increase in myosin I
mRNA was found (Fig.
4A) with a similar decrease in myosin I
protein occurring (Fig. 4B), and it also was found in the urine (Fig.
4C). Immunolocalization showed a more dramatic decrease in
myosin I
staining (Fig. 7, C, E, and
F) after 2 h of reperfusion. Although the magnitude of
the myosin I
decrease was variable, there was a significant reorganization of this protein after ischemia and reperfusion (compare the color image of the control in Fig. 7A with the
2-h recovery state in Fig. 7F). This was most evident by the
reduced staining of the BB for myosin I
. In addition, the intensity
of myosin I
labeling was often increased in nonproximal tubular cells at 2 h. These results suggest that myosin I
does not
participate in the early rebuilding of the microvilli that occurs after
reperfusion. By 24 h of reperfusion (Fig. 7, G-I)
there were clearly regions of proximal tubules that had myosin I
in
the BB (Fig. 7H, arrows), although there was still a higher
percentage of cytoplasmic labeling than observed in the normal
physiological state, and many proximal tubules had little myosin I
in the BB (Fig. H, triangles). In addition, we often
observed an increase in myosin I
staining in nonproximal tubules at
24 h of recovery. Interestingly, myosin I
was also found in the
urine from the 2- to 24-h reperfusion collection. The results combined
suggest that a dramatic cytoskeletal and membrane reorganization of
kidney cells has taken place, after 15-min clamp, that does not cause
any significant release of protein but requires the cells to retarget
and reorganize multiple cellular components, one of which is myosin
I
. Normal proximal tubule staining for myosin I
returned by 3 days of reperfusion (data not shown). To obtain a better understanding
of myosin I
changes in other tubule segments, changes to specific
tubular markers during ischemia-reperfusion must be followed.
These studies, along with studies aimed at better defining the recovery
of myosin I
in the PTC, are needed to provide a clearer picture of
the role of myosin I
in kidney tubule function and recovery from
ischemia.

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Fig. 7.
Cellular location of myosin I and F-actin after 15 min of
ischemia and 2 or 24 h of reperfusion. Control tissue and
ischemic tissue are presented in A and B,
respectively, as color composites of images in Fig. 2, E and
F, and Fig. 5, C and D. All color
overlays show myosin I in purple and F-actin in green, and
colocalization is indicated as white. D-F:
reconstructions of 15 0.5-µm planes that show myosin I , F-actin,
and the color composite from 15 min of ischemia and 2 h of
reperfusion. C: myosin I staining from a kidney treated
with 15 min of ischemia and 2 h of reperfusion;
indicates the PT. G-I: images of
myosin I (H and I)- and F-actin-labeled
(G) kidney sections after 15 min of ischemia and
24 h of reperfusion. Even after 24 h of recovery, we still
found most PTs lacking the normal staining pattern for myosin I .
Arrows identify PTs with some normal myosin I staining, and
triangles identify PTs lacking any significant myosin I staining in
the BB. Bars = 10 µm.
|
|
 |
DISCUSSION |
Ischemia triggers many cellular alterations along the
nephron (4, 24, 35, 39). Reduced blood flow results in low cellular ATP levels, effectively inhibiting many ATP-dependent processes such as phosphorylation, ion regulation, and vesicle trafficking. These alterations contribute to the cell injury observed and lead to tubule and kidney dysfunction. The best-studied
ischemic changes occur in proximal tubules, where the apical
cytoskeleton undergoes dramatic rearrangement after cessation of blood
flow in vivo. Actin, the major protein component of the microvilli, is
redistributed in the apical cytoplasm. Its regulation and associated proteins, including ezrin and actin-depolymerizing factor (ADF), are
also altered under ischemic conditions (10, 11,
31). It has become increasingly apparent that ischemia
disrupts the normal cellular morphology and physiology of the distal
segments of the nephron as well (14, 15, 19, 21, 29).
Return to the normal physiological state requires retargeting of
misdirected cellular components and rebuilding of the cytoskeleton.
Myosin I
is an actin-activated ATPase with both actin and membrane
binding domains (3). It has three calmodulin light chain
binding domains that bind calmodulin strongest in the absence of
calcium. Its widespread distribution argues for involvement in a
fundamental cell process shared by multiple cell types. In these
studies, we analyzed its tubular distribution and addressed its
potential structural role in PTC microvilli. It has previously been
suggested that myosin I tethers actin to the plasma membrane of the
microvillus (27). Under control conditions, myosin I
was found to be ~90% Triton insoluble, probably due to its
interaction with F-actin. Immunofluorescence labeling of myosin I
and F-actin in normal kidney tissue sections showed extensive
colocalization of these two proteins in the BBM. There is strikingly no
colocalization of myosin I
with actin in the stress fibers of these
cells. There appears to be a zone of minimal myosin I
labeling
beneath the BB, which may correspond to the proximal tubule terminal
web region. This zone is often followed by a punctate distribution of
myosin I
in the cytoplasm. The myosin just beneath the BB may be
associated with vesicles that are to be transported along actin
filaments to the microvillar membrane. Multiple studies have implicated myosin I's as playing a role in endocytosis and intracellular trafficking (23, 33, 40). Movement into and throughout the BB region, a region lacking microtubules, may utilize an actin-based motor such as myosin I
. Alternatively, myosin I
in the subapical region could be simply waiting for transport to the BB via another myosin motor. Distinguishing whether this myosin is actively involved in transport to the BB or a non-transport function would be aided by a
better understanding of what this myosin interacts with and how these
interactions are regulated. Interactions between multiple myosin I
proteins have been suggested as a prerequisite for this motor to
perform work, and the subapical punctate distribution may represent a
small cluster of myosin I
molecules (28). It is widely
accepted that the actin cytoskeleton plays a role in the regulation of
multiple channels; thus it is possible that myosin I actin-based motors
contribute to these regulatory processes (8, 32, 34,
38). Myosin I's may also help shuttle membrane vesicles
containing channels to and from the plasma membrane and/or may affect
the dynamics of the actin cytoskeleton, thus indirectly contributing to
channel regulation.
Under normal physiological conditions, myosin I
and actin are
concentrated in the BB of the proximal tubule. With ischemic injury, the actin cytoskeleton is disrupted, microvilli shorten and
fuse, ATP levels fall, and intracellular calcium levels rise. Low
levels of ATP favor formation of a rigor complex between myosin and
actin. Interestingly, after 15 min of ischemia, myosin I
is
observed in cellular blebs in the proximal tubule lumens without F-actin. This early separation of myosin I
and F-actin may result from the rapid degradation of microvillar F-actin by ADF, which is
present in these blebs, and the calcium activation of F-actin severing
proteins, such as gelsolin (6, 24, 31). What is clear is
that the microvillar F-actin undergoes dramatic alterations within 15 min of ischemia that result in apical enrichment of myosin I
and G-actin in this area (31, 35). A possible scenario to
explain what happens to myosin I
follows: increased intracellular calcium releases calmodulin from myosin I
, causing an increase in
ATPase activity and membrane binding. This enables the myosin to move
quickly up the collapsing microvillar F-actin, where it associates with
the membrane as F-actin breaks down. The polarity of F-actin (barbed
end apical) is such that myosin I
would move to the barbed end.
Myosin I
could then associate with a select group of acidic
phospholipids that collect in the blebs or bind to a receptor protein
that has accumulated in the blebs. We are currently examining some of
these possibilities by analyzing BBM vesicles from normal and
ischemic proximal tubules.
While the precise sequence of these cellular ischemic events is
uncertain, it is well established that return of physiological cell and
organ function requires reestablishment of the normal cell
architecture. This will involve, in part, rebuilding the apical
microvilli and retargeting membrane and proteins back to their correct
location. We showed in these studies that normal F-actin structure is
obtained after 2 h of reperfusion following a 15-min clamp.
Surprisingly, myosin I
normal staining pattern is not returned to
normal. In fact, myosin I
is present throughout the cytoplasm in
both proximal and distal tubules and often appears in larger punctate
structures than in the control tissue. This suggests that either the
cellular process(es) in which this motor protein participates has not
recovered or it may be actively assisting in the retargeting and/or
reorganization of actin cytoskeleton and membrane components. Even
after 24 h of reperfusion there are regions of PTC that do not
appear normal for myosin I
. However, at 2 and 24 h of recovery,
we have shown that there are only minor changes in both protein and
mRNA levels for myosin I
. This finding suggests that the changes we
observe by immunofluorescence are the result of dramatic cytoskeletal
and membrane reorganization that occurs after an ischemic
insult. Apparently, instead of simply making more of the myosin
I
, the cells have to retarget and reestablish this protein's normal
cellular interactions, which our data suggest is not complete even
after 24 h of recovery. We believe that interactions of myosin
I
interactions with membrane phospholipids, F-actin, and possibly
membrane proteins undergo significant reorganization during reperfusion
that ultimately contributes to the return of functioning kidneys.
During this reorganization, a reduced staining pattern for myosin I
is observed in the BB. Our results suggest that this is caused by a
redistribution of this myosin away from the BB. Analysis of distinct
subcellular fractions, i.e., BBM, will help to explain these changes.
We are presently examining the recovery time points following the
ischemic block to better define the physiological changes that
may coincide with the return of myosin I
. The major conclusion from
these studies is that myosin I
is present throughout the kidney
tubules, its distribution is severely altered by ischemia, and
its return follows the apparent structural return of the BB.
 |
ACKNOWLEDGEMENTS |
This research was supported by National Institute of Diabetes and
Digestive and Kidney Diseases Grant DK-54923.
 |
FOOTNOTES |
Address for reprint requests and other correspondence: M. C. Wagner, Indiana Univ. School of Medicine, 1120 South Dr., FH 115, Indianapolis, IN 46202-5116 (E-mail: wagnerm{at}iupui.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.
Received 17 October 2000; accepted in final form 25 May 2001.
 |
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