Department of Cellular Biology, University of Georgia, Athens, Georgia 30602, USA
* Author for correspondence (e-mail: fechheim{at}cb.uga.edu )
Accepted 26 February 2002
![]() |
Summary |
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Key words: Cytoskeleton, Actin-binding protein, Dictyostelium, Hirano body, Neurodegeneration
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Introduction |
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Cytoplasmic inclusions termed Hirano bodies have been described in a
variety of neurodegenerative diseases and other conditions that produce
persistent injury or stress (Hirano,
1994). The structures contain actin filaments and actin-associated
proteins (Galloway et al.,
1987
; Goldman,
1983
; Maciver and Harrington,
1995
). However, their mechanism of formation, composition and
relation to disease remains poorly understood.
In this study, we report the serendipitous development of a cultured cell
model for studies of Hirano bodies in the cellular slime mold
Dictyostelium discoideum, a lower eukaryote with a well-characterized
cytoskeleton, and facile methods for protein expression and creation of mutant
strains (Mann et al., 1998;
Noegel and Schleicher, 2000
).
The 34 kDa protein is one of 11 actin crosslinking proteins present in
Dictyostelium (Furukawa and
Fechheimer, 1997
). In vitro studies reveal that actin bundling by
the purified 34 kDa protein is calcium regulated
(Fechheimer, 1987
;
Fechheimer and Furukawa, 1993
;
Fechheimer and Taylor, 1984
;
Lim and Fechheimer, 1997
).
Experiments with defined segments of recombinant protein reveal three
actin-binding sites located at amino acids 1-123, 193-254 and 279-295
(Lim et al., 1999a
). The
strongest of these sites, located at amino acids 193-254, is necessary and
sufficient for co-sedimentation with F-actin in vitro
(Lim et al., 1999a
). The CT
fragment, comprising amino acids 124-295, lacks the inhibitory domain located
in the N-terminus that modulates the activity of the strong actin-binding site
through an intramolecular interaction (Lim
et al., 1999b
). Truncation of the inhibitory region from the CT
fragment results in enhanced binding and crosslinking of actin filaments that
is calcium-insensitive (Lim et al.,
1999b
).
In this paper, we report that expression of low levels of the CT protein in Dictyostelium induces formation of paracrystalline actin inclusions that resemble Hirano bodies in both ultrastructure and composition as assessed by immunocytochemistry. Similarly, expression of the CT fragment induces formation of Hirano bodies in murine L cells. These results show that formation of Hirano bodies is restricted neither to mammalian cells nor to nerve cells. Rather, the formation of Hirano bodies appears to be a general response to or consequence of aberrant function of the actin cytoskeleton. In addition, we report the first studies of the physiological effects of Hirano bodies on cell function.
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Materials and Methods |
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Growth and transformation of Dictyostelium
AX-2 and derivatives described below were routinely maintained in axenic
shaking cultures in HL-5 media (Loomis,
1971). Cell transformation was performed and growth rates were
measured as described previously (Rivero
et al., 1996
). Clones were screened by western blotting or
staining with rhodamine-labelled phalloidin to identify actin inclusions. Once
a clone was established, it was transferred to shaking culture in HL-5 medium,
and multiple aliquots were frozen at -80°C in HL-5 media supplemented with
10% DMSO. Since the incidence of actin inclusions decreased during routine
passage of cells, all assays were performed on cells that were maintained in
shaking culture for less than 30 days.
Development and germination
Cells were grown in HL-5 media in suspension to a concentration of
3x106 cells/ml, harvested by centrifugation, washed once in
17 mM Soerensen buffer, and 1x107 cells in 0.5 ml 17 mM
Soerensen phosphate buffer were plated on 100 mm non-nutrient agar plates.
Photographs were taken on a Wild 500 MacroScope.
To assess germination of spores, fruiting bodies were collected and the spores were placed in HL-5 media. The number of amoebae and ungerminated spores were counted after 4.5 hours to assess viability. The amoebae germinated from spores were stained with rhodamine-labeled phalloidin to identify actin inclusions.
Electron microscopy
Dictyostelium cells were fixed for transmission electron
microscopy as described previously (Novak
et al., 1995). Cells were embedded in EPON 812 and sectioned on an
RMC 5000 ultramicrotome (Tuscon, AZ) with a diamond knife. Micrographs were
taken on a Phillips 400 TEM.
Immunofluorescence
Dictyostelium cells were fixed for 20 minutes in 3.7% formaldehyde
in 17 mM phosphate buffer containing 1 mM CaCl2, pH 7.1, and
permeabilized using acetone at -20°C for 2 minutes as described previously
(Fechheimer, 1987). Cells were
stained with B2C monoclonal antibody to the 34 kDa protein
(Furukawa et al., 1992
) or
anti-myc monoclonal antibody 9E10 (ATCC CRL-1729). Tubulin and
Dictyostelium MAP were identified with rat monoclonal antibody YL 1/2
and antibody AX3, respectively (gift of M. Kimble, University of South
Florida). Myosin II was identified with mouse polyclonal antibody NU48 (gift
of R. Chisholm, Northwestern University Medical School). Myosin II heavy chain
was identified with polyclonal antibody 9555-3 (gift of A. De Lozanne,
University of Texas at Austin). Cofilin was identified with rabbit polyclonal
antibody (gift of H. Aizawa, Tokyo Metropolitan Institute of Medical Science,
and C. Chia, University of Nebraska). EF-1
, cofilin, ABP120 and
-actinin were identified with rabbit polyclonal antibodies (gift of J.
Condeelis, Albert Einstein College of Medicine). The rhodamine- and
fluorescein-conjugated secondary antibodies were obtained from Sigma Chemical
Co. (St Louis, MO). The rhodamine- and Oregon green 488-labelled phalloidins
were obtained from Molecular Probes (Eugene, OR).
Phagocytosis and pinocytosis
The phagocytosis assay was performed with fluorescently labeled yeast in
suspension essentially as described previously
(Maniak et al., 1995) and
modified by Meg Titus (University of Minnesota). The pinocytosis assay was
performed using lucifer yellow on cells attached to 24-well tissue culture
plates as described (Rivero et al.,
1996
).
Measurements of actin in Dictyostelium amoebae
The total actin content of the amoebae was assessed by western blotting.
Dictyostelium cells were counted, washed and lysed, and proteins were
resolved by SDS-PAGE. The samples were then blotted to nitrocellulose, labeled
with an anti-actin monoclonal primary antibody (10-B-3, gift of M. Kandasamy
and R. B. Meagher, University of Georgia) followed by an
alkaline-phosphatase-conjugated secondary antibody (Promega, Madison, WI), and
treated with BCIP/NBT to visualize the bands. The blot was photographed with
direct positive black and white film, and scanned using a Molecular Dynamics
Laser Scanning Densitometer.
F-actin levels for the whole population of cells were determined with
rhodamine-labelled phalloidin (Sigma, St. Louis, MO) using the method
described previously (Cano et al.,
1992). Data presented represent fluorescence units per mg total
protein from six independent samples.
Comparisons of the amount of F-actin in single cells were made by measurement of the fluorescence from single cells stained with rhodamine-labelled phalloidin. Images were recorded with a Mighty Max Cooled CCD camera (Princeton, Trenton, NJ). Care was taken not to saturate any pixels in the array. The mean pixel intensity of AX-2 and CT-myc cells were compared using IP lab image analysis software (Scanalytics, Fairfax, VA). The data reported represent mean pixel values from at least 24 individual cells.
Expression of CT fragment in murine L cells
The sequences encoding the CT fragment of the 34 kDa protein (aa 124-295)
were subcloned into the BamHI site of the vector pEGFP N1 (Clontech)
following amplification by PCR using custom oligonucleotides. The vector was
designed to express the CT fragment alone, and not a fusion protein of CT with
GFP. The coding sequence was confirmed by automated DNA sequencing.
L-cell fibroblasts were maintained in RPMI-1640 supplemented with 15% bovine serum, 100 U/ml penicillin/streptomycin, and 2 mM L-glutamine at 37°C in 5% CO2. The cells were plated at 1x105 cells/22 mm2 coverslip (50-60% confluence), and transfected using Lipofectamine Plus (Life Technologies) according to the manufacturer's protocol. Cells were transfected with no DNA (mock), or 1 µg of the pEGFP vector only or the pCTEGFP plasmid. The transfection efficiency was approximately 60% as assessed from the expression of GFP in the cells transfected with vector alone.
Cells were fixed 24 hours following replacement of complete media to the transfected cells. Coverslips were rinsed in PBS, transferred to 3.7% formaldehyde in PBS for 25 minutes, washed briefly three times in PBS, permeabilized in 0.1% Triton X-100 in PBS for 5 minutes, washed briefly three times in PBS, blocked in 1% BSA in PBS for 1 hour, and stained with Oregon Green 488-labelled phalloidin for 1 hour (Molecular Probes) before being mounted.
All slides were coded and scored by observers unbiased by knowledge of the identity of the treatment groups. Cells were counted and categorized according to the F-actin distribution as follows: (1) predominantly in cortical arrays, stress fibers, and filament bundles; (2) predominantly in punctate and stellar foci; and 3) contain a large aggregate/Hirano body. All experiments were repeated at least 3 times. Significance was judged using a Student's T test to compare the means ± standard deviations of the control and CT expressing cells.
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Results |
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CT-myc induces formation of actin-containing ellipsoids
In cells expressing full-length 34 kDa-myc protein, the localization of the
full-length 34 kDa protein with the myc epitope tag is identical to the
localization pattern for the native 34 kDa protein. Both native and tagged
proteins can be found in the leading edge and the cell cortex of vegetative
amoebae (Fig. 2). The actin
localization pattern in cells expressing 34 kDa-myc protein is the same as
AX-2 wild-type cells, and largely overlaps the localization of the 34 kDa
protein.
|
Surprisingly, cells expressing low levels of the CT-myc protein produce ellipsoidal inclusions that label strongly with rhodamine-labelled phalloidin, anti-34-kDa protein antibody (B2C), and anti-myc antibody (Figs 2, 3). The B2C monoclonal antibody to the 34 kDa protein recognizes an epitope within the CT fragment, and so is expected to label the expressed CT-myc (R. W. L. Lim, R.F. and M.F., unpublished). The ellipsoids do not incorporate all the endogenous full-length 34 kDa protein, as cells containing ellipsoids exhibit significant staining for the 34 kDa protein in the cortex. By contrast, staining of the cortex with the antibody to the myc epitope is less prominent, which indicates that the cortical staining emanates primarily from full-length endogenous 34 kDa protein. The difference in localization is apparent in a direct comparison of the anti-34-kDa protein and anti-myc antibodies (Fig. 3). The B2C antibody staining of both the native 34 kDa protein and the CT-myc protein is visible in both the cell cortex, the normal site of localization and the ellipsoid. By contrast, the actin-containing ellipsoid is stained predominantly when the cells are labeled with antibody to the myc epitope. Although the ellipsoid is roughly the same size as the nucleus, DAPI staining of DNA demonstrates that the ellipsoids are distinct from the nucleus (data not shown).
|
CT-myc cells were labeled with a variety of antibodies to assess the status
of selected known cytoskeletal proteins. The ellipsoids do not contain
-tubulin or a Dictyostelium MAP
(Kimble et al., 1992
), and no
differences in the pattern of interphase microtubules between wild-type cells
and cells expressing CT-myc are detected
(Fig. 4A). The ellipsoids are
also enriched in myosin II, cofilin and
-actinin
(Fig. 4B-D), but do not stain
with antibodies to EF-1
, ABP 120
(Fig. 4E,F), and myosin I (data
not shown).
|
Ultrastructure and actin filament organization in the ellipsoids
Ultrastructural observation reveals that the ellipsoids are present in the
cytoplasm of CT-myc cells, and that no lipid bilayer circumscribes the
ellipsoids (Fig. 5A). The
ellipsoids are most commonly located near the center of the cell, but they
also can be observed adjacent to the cell membrane.
|
The F-actin in the ellipsoids is highly ordered, as revealed by
transmission electron microscope images of thin sections. An oblique section
through the ellipsoids reveals a pattern that resembles a herringbone fabric
(Fig. 5B). Tilting the section
reveals that the herringbone appearance is a plane of section effect, as the
same field reveals parallel filaments when viewed at a 45° tilt
(Fig. 5C). This observation is
confirmed by cross-sections that reveal different degrees of order in the
ellipsoids. Some of the regions have an appearance similar to cytoplasmic
extensions that can be seen at the cell periphery, and presumably contain
loosely ordered actin that is not well preserved using methods of preparation
for electron microscopy that include treatment with OsO4
(Maupin-Szamier and Pollard,
1978) (Figs 5,
6). Many regions are highly
ordered, with the filaments arranged in a square packed arrangement
(Fig. 6A,D). In these square
packed regions the filaments are arranged in a double row, with a center to
center spacing of 20 nm, separated by a space of 25 nm between double rows.
The pattern of double rows is clearly visible in both longitudinal and
cross-sections (Fig. 6C,D).
Square packed actin bundles have not been observed previously in
Dictyostelium, but do form in vitro in mixtures of actin with
Dictyostelium elongation factor 1
(ABP 50)
(Owen et al., 1992
). However,
note that ABP-50 is not concentrated in these structures in vivo
(Fig. 4F). Hexagonally ordered
actin filaments represent the most compact state. In this arrangement each
actin filament is 15 nm from its six nearest neighbors
(Fig. 6B). In a fortuitous
section these different states of order can be seen in the same ellipsoid
(Fig. 6A), showing both the
square packed and hexagonally packed filament patterns.
|
Growth, development and endocytosis by CT-myc cells
Since the CT-myc cells reveal dramatic reorganization of F-actin, the
ability of the cells to perform the primary functions of growth and
development was assessed. CT-myc cells grow slowly in shaking axenic culture
at 20°C. The G418-resistant AX-2 cells expressing vector alone double 2.06
times per day as opposed to 2.04 doubles per day for 34 kDa-myc cells and only
1.46 doubles per day for CT-myc cells. To track the portion of the population
with ellipsoids, samples of CT-myc cells were taken daily and stained with
rhodamine phalloidin. The number of cells presenting ellipsoids remained
constant at about 40% until late log phase. During late log phase the number
of ellipsoids visible in the population was substantially smaller.
Furthermore, ellipsoids are rare in cells that have been maintained in culture
for more than 1 month. Each of these observations was confirmed in at least
two independent transformants, indicating that they are a consequence of
expression of CT-myc, and not from adventitious effects arising from
integration of the vector and selection of cell clones. Since loss of
expression is an inevitable consequence of the slow growth of the CT-myc
cells, new cultures were regularly established from frozen stock, and all data
presented are from cells maintained in axenic shaking culture for less than 30
days.
To assess the ability of CT-myc cells to undergo development, cells from shaking culture were collected and washed by centrifugation and plated on non-nutrient agar. The CT-myc cells are delayed in development by 3-6 hours (Fig. 7). When the wild-type and 34 kDa-myc expressing cells are making mounds, the CT-myc cells are only beginning to stream. When wild-type cells are beginning to culminate, the CT-myc cells have just started making mounds. After 30 hours, the CT-myc cells produce fruiting bodies with normal morphology, approximately 6 hours later than wild-type cells (Fig. 7).
|
The spores that are produced by CT-myc cells produce viable amoebae. After 4.5 hours in HL-5, 67% and 61% of the 34 kDa-myc and CT-myc spores germinated, respectively. The population of amoebae germinated from CT-myc spores contains about 40% individuals with large ellipsoids, a fraction similar to that of the population from which the spores were derived. Thus, the selection against CT-myc expression that was observed after long term culture was not evident after a single round of development and subsequent germination.
The presence of the 34 kDa protein and actin in the phagocytic cup
(Furukawa et al., 1992;
Furukawa and Fechheimer, 1994
;
Rezabek et al., 1997
) and the
participation of actin in macropinocytosis in Dictyostelium
(Hacker et al., 1997
)
suggested that alteration of the actin cytoskeleton by expression of CT-myc
might have a detrimental effect on endocytosis. Surprisingly, the rate of
phagocytosis of fluorescently labeled yeast in shaking culture is not
significantly different in assays using CT-myc and AX-2 cells (data not
shown). Further, the rate of pinocytotic uptake of lucifer yellow by adherent
cells in HL-5 axenic media reveals no significant difference between wild-type
AX-2 and CT-myc cells (data not shown).
Changes in actin in CT-myc cells
The presence of stable F-actin-containing ellipsoids in the cytoplasm of
Dictyostelium suggests that the actin levels and/or dynamics in the
cell may be altered. Actin comprises 8% of the protein in axenic
Dictyostelium amoebae (Brier et
al., 1983). CT-myc cells had a level of total actin that was
96±23% of wild-type as assessed by western blots of whole cell
lysates.
The total actin pool contains free G-actin, unpolymerized sequestered
monomeric actin, and F-actin in quantities that depend on changes in
actin-monomer-binding proteins, and the presence of free or capped barbed and
pointed ends on the actin filaments
(Fechheimer and Zigmond,
1993). The F-actin levels in the whole population were examined by
lysing cells, labeling the F-actin with rhodamine-labeled phalloidin, and
measuring bound phalloidin after high speed sedimentation of the actin
filaments. The F-actin levels in CT-myc cells are 120±13% of wild-type.
This difference is not statistically significant. However, since only 40% of
CT-myc cells contain ellipsoids, the difference in F-actin levels between
wild-type and CT-myc cells with ellipsoids may be more striking than is
revealed by measurements of the bulk population of CT-myc cells.
To compare F-actin levels between AX-2 and the subsets of CT-myc cells that do or do not contain ellipsoids, we examined the intensity of rhodamine-labelled, phalloidin-derived fluorescence from selected cells with images obtained and quantified using a cooled CCD camera. These comparisons were made by identifying rhodamine-labelled, phalloidinstained CT-myc cells that did or did not exhibit ellipsoids, and comparing their mean pixel intensity with AX-2 cells. Cells in the CT-myc population that did not display ellipsoids had 117±3.8% of the F-actin of wild-type. The level of F-actin in CT-myc cells presenting ellipsoids was 183±4.6% of wild-type. Thus, the F-actin levels in CT-myc cells containing ellipsoids is significantly higher than that in wild-type cells. Since CT-myc cells contain equivalent total actin and increased F-actin compared with wild-type cells, these cells must have a smaller pool of unpolymerized actin than wild-type cells.
Formation of Hirano bodies in L cells
The actin distribution in murine L cells expressing the CT fragment was
investigated to determine whether the results obtained in
Dictyostelium might be extended to a mammalian cell culture system.
Control (untransfected, mock transfected, or vector only transfected)
fibroblasts have a typical F-actin distribution characterized by cortical
filaments, filament bundles and few punctate foci
(Fig. 8A). Cells expressing the
CT fragment exhibited a loss of stress fibers and cortical F-actin, and
accumulated multiple punctate foci (Fig.
8B) and large aggregates resembling Hirano bodies
(Fig. 8C-E). These results were
quantified revealing that the decrease in stress fibers, and accumulation of
multiple foci and large aggregates are representative of the populations
(Fig. 8F). The differences in
actin distributions of control and CT-transfected cells were statistically
significant as assessed by a Student's t-test
(P<0.01).
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Discussion |
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Hirano bodies have been reported in normal and pathological specimens from
humans and animals providing our primary clues to their etiology. Hirano
bodies form in association with a broad array of conditions including
Alzheimer's disease (Gibson and Tomlinson,
1977; Mitake et al.,
1997
; Mori et al.,
1986
; Schmidt et al.,
1989
), Parkinson's disease
(Hirano et al., 1968
), Pick's
disease (Schochet et al.,
1968
), amyotrophic lateral sclerosis
(Hirano et al., 1968
), ataxic
Creutzfeldt-Jakob disease (Cartier et al.,
1985
), kuru (Field et al.,
1969
), scrapie (Field and
Narang, 1972
), Papovavirus
(Hadfield et al., 1974
),
chronic alcoholism (Lass and Hagel,
1994
), diabetes (Sima and
Hinton, 1983
), cancer (Fu et
al., 1975
; Gessaga and Anzil,
1975
), muscle degeneration
(Fisher et al., 1972
), and
neuronal degeneration associated with abnormal copper homeostasis
(Anzil et al., 1974
;
Nagara et al., 1980
;
Peterson et al., 1986
;
Waggoner et al., 1999
).
Hirano bodies have been observed most frequently in the hippocampus
(Cartier et al., 1985;
Gibson and Tomlinson, 1977
;
Hirano et al., 1968
;
Mitake et al., 1997
;
Mori et al., 1986
;
Ogata et al., 1972
;
Schmidt et al., 1989
;
Schochet et al., 1968
), but
have also been observed in other regions of the brain including Purkinje cells
(Yamamoto and Hirano, 1985
),
cerebellum (Nagara et al.,
1980
; Peterson et al.,
1986
), cerebral cortex (Anzil
et al., 1974
), and peripheral neurons
(Atsumi et al., 1980
;
Doering and Aguayo, 1987
;
Sima and Hinton, 1983
;
Yagishita et al., 1979
).
Hirano bodies are not restricted to neurons; they have been reported in
oligodendroglial cells (Gibson,
1978
; Okamoto et al.,
1982
; Sima and Hinton,
1983
), muscle fibers
(Fernandez et al., 1999
;
Fisher et al., 1972
;
Tomonaga, 1983
), and testis
(Setoguti et al., 1974
).
These findings have supported a variety of suggestions regarding the
physiological or pathological significance of Hirano bodies. Because Hirano
bodies were reported in brain tissue from individuals with a wide variety of
conditions, they have been characterized as a nonspecific manifestation of
neuronal degeneration (Schochet and
McCormick, 1972), nonspecific changes with no relation to
pathology (Ogata et al.,
1972
), and nonspecific arrangements of filament units largely
devoid of cytopathological significance
(Gessaga and Anzil, 1975
).
However, both the data in the literature and our results presented above
support a broader interpretation. Since the filament arrangement is
paracrystalline and highly ordered and contains actin but neither microtubules
nor intermediate filaments, the claim that Hirano bodies are a nonspecific
arrangement of filaments is clearly not well supported. Moreover, the finding
of Hirano bodies in a variety of tissues and in association with a broad array
of conditions does not prove that they are nonspecific and lacking in
cytopathological significance. Rather, we propose that a range of conditions
may generate a signal(s) that causes rearrangement of the actin cytoskeleton
and induces the formation of Hirano bodies. Our finding that Hirano bodies can
be induced to form by expression of an activated and unregulated fragment of
an actin crosslinking protein strongly supports this interpretation.
It is interesting to ponder the mechanism of formation of the Hirano
bodies. First, the CT fragment binds with high affinity to actin filaments in
vitro, and would disassociate slowly from actin filaments strongly delaying
their disassembly (Zigmond et al.,
1992). The CT fragment induces formation of large extended tangles
of actin filaments in vitro (Lim et al.,
1999a
; Lim et al.,
1999b
). This could result in accumulation of ordered structures,
promoted by entropic forces that would induce alignment into a highly ordered
stable array (Furukawa and Fechheimer,
1997
). Second, the presence of myosin II in the Hirano bodies
(Fig. 4B) raises the
possibility that active generation of contractile force is involved in the
formation of these structures. Third, it is possible that following initial
formation of a number of small arrays, formation of a single large Hirano body
occurs by transport directed towards the minus ends of microtubules as has
been shown for formation of aggresomes
(Johnston et al., 1998
). In
addition, it is possible that aberrant function of the actin cytoskeleton
generates a signal for formation of Hirano bodies. Studies of such potential
mechanisms will be an important goal of future research efforts.
Our studies of live Dictyostelium cells containing Hirano bodies are the first studies of the effects of these inclusions on cell physiology, since prior studies have employed fixed material. Dictyostelium cells with Hirano bodies grow slowly in suspension, and develop more slowly (Fig. 7) than wild-type cells, but perform phagocytosis and pinocytosis normally. These cells have normal total actin levels, but an increased amount of F-actin, and consequently, a decreased amount of unpolymerized actin compared with wild-type cells. Thus, Hirano bodies may act as a sink for F-actin, sequestering actin and reducing the concentration of free actin available for cellular processes. Thus, the slow growth and slow development phenotypes we observe in CT-myc cells may be due to a reduction in the unpolymerized actin pool. Alternatively, these phenotypes may arise from mechanical hindrance of the ellipsoid and/or disruption of the viscoelastic properties of cytoplasm resulting from CT-myc expression. It is also possible that the presence of a large highly crosslinked ellipsoid may impede processes that require global cytoskeletal changes.
Our findings show clearly that Hirano bodies are not necessarily linked to
a stage in cell death. They are toxic neither in Dictyostelium, nor
in mouse fibroblasts. What then, if any, is the possible significance of
Hirano bodies? Actin/cofilin rods are bundles of actin-containing ADF/cofilin
that are induced in the cytoplasm following either ATP depletion, oxidative
stress (Minamide et al., 2000)
or NaCl treatment (Nishida et al.,
1987
), and in the nucleus following DMSO treatment
(Ono et al., 1993
) or heat
shock (Nishida et al., 1987
).
These ADF/cofilin actin rods do not reveal the paracrystalline filament
organization characteristic of Hirano bodies. Further, ADF/cofilin rods fail
to stain with phalloidin, in marked contrast to Hirano bodies, which are
strongly labeled both with fluorescent phalloidin and with antibody to
cofilin. The change in the twist of the actin filament that results from
cofilin-binding explains the reported absence of phalloidin binding to
ADF/cofilin rods (McGough et al.,
1997
). These ADF/cofilin actin rods are associated with >97% of
amyloid deposits in brain from Alzheimer's patients, and can form in neurites
in cultures resulting in loss of distal microtubules and absence of growth
cones (Minamide et al., 2000
).
These exciting recent results focus attention on aberrant behavior of the
actin cytoskeleton as a potentially important feature of loss of cell function
in neurodegenerative diseases. Our results show that it cannot be assumed that
formation of Hirano bodies is deleterious to cell function. The formation of
Hirano bodies may be an adaptive change that promotes cell function. It is
possible that the sequestration of actin, cofilin, and other components in
Hirano bodies may promote cytoskeletal function by removing them from other
regions of the cell. Future studies of actin/cofilin rods and Hirano bodies
both in Dictyostelium and in mammalian cells may help to elucidate
the role(s) of the actin cytoskeleton in the progression of neurodegenerative
diseases.
In closing, we propose that formation of Hirano bodies is a general cellular response to or a consequence of aberrant function of the actin cytoskeleton. The unregulated fragment of the 34 kDa protein is the first well-characterized signal shown to induce formation of these assemblies. These findings provide insight into the delicate balance between formation and disassembly of crosslinked actin structures that is necessary for the proper function of the actin cytoskeleton. Actin-binding proteins must be regulated or bind weakly to actin to prevent the accumulation of crosslinked structures. The ability to induce Hirano bodies in cultured cell lines will allow us to explore the mechanism of their formation, and potential adaptive or pathological roles in the various conditions with which they are associated.
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Acknowledgments |
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![]() |
Footnotes |
---|
C-terminal portion of the Dictyostelium 34 kDa actin-bundling
protein (amino acids 124-295) with a C-terminal myc epitope tag.
![]() |
References |
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