Cell Motility and the Cytoskeleton 54:29 – 40 (2003)
Dynamic Localization of Myosin-I to
Endocytic Structures in Acanthamoeba
E. Michael Ostap,1* Pamela Maupin,2 Steven K. Doberstein,2 Ivan C. Baines,3
Edward D. Korn,3 and Thomas D. Pollard4
1
The Pennsylvania Muscle Institute and The Department of Physiology, University of
Pennsylvania School of Medicine, Philadelphia
2
Department of Cell Biology and Anatomy, Johns Hopkins University School of
Medicine, Baltimore, Maryland
3
Laboratory of Cell Biology, National Heart, Lung, and Blood Institute, National
Institutes of Health, Bethesda, Maryland
4
Department of Molecular, Cellular, and Developmental Biology Yale University,
New Haven, Connecticut
We used fluorescence microscopy of live Acanthamoeba to follow the time course
of the concentration of myosin-I next to the plasma membrane at sites of macropinocytosis and phagocytosis. We marked myosin-I with a fluorescently labeled
monoclonal antibody (Cy3-M1.7) introduced into the cytoplasm by syringe loading. M1.7 binds myosin-IA and -IC without affecting their activities, but does not
bind myosin-IB. Cy3-M1.7 concentrates at two different macropinocytic structures: large circular membrane ruffles that fuse to create macropinosomes, and
smaller endocytic structures that occur at the end of stalk-like pseudopodia. These
dynamic structures enclose macropinosomes every 30 – 60 s. Cy3-M1.7 accumulates rapidly as these endocytic structures form and dissipate rapidly after they
internalize. Double labeling fixed cells with Cy3-M1.7 and polyclonal antibodies
specific for myosin-IA, -IB, or -IC revealed that all three myosin-I isoforms
associate with macropinocytic structures, but individual structures vary in their
myosin-I isoform composition. Myosin-I and actin also concentrate transiently at
sites where amoebae ingest yeast or the pseudopodia of neighboring cells (heterophagy) by the process of phagocytosis. Within 3 min of yeast attachment to the
amoeba, myosin-I concentrates around the phagocytic cup, yeast are internalized,
and myosin-I de-localizes. Despite known differences in the regulation of macropinocytosis and phagocytosis, the morphology, protein composition, and dynamics of phagocytosis and macropinocytosis are similar, indicating that they
share common structural properties and contractile mechanisms. Cell Motil.
Cytoskeleton 54:29 – 40, 2003. © 2003 Wiley-Liss, Inc.
Key words: motility; cytoskeleton; actin; macropinocytosis; phagocytosis; heterophagy
Contract grant sponsor: NIH; Contract grant number: GM-26132;
Contract grant sponsor: Cancer Research Fund of the Damon RunyonWalter Winchell Foundation Fellowship; Contract grant number:
DRG-1294; Contract grant sponsor: NIH; Contract grant number:
GM-57247.
*Correspondence to: E.M. Ostap, Department of Physiology, University of Pennsylvania School of Medicine, B400 Richards Building,
3700 Hamilton Walk, Philadelphia, PA 19104-6085.
E-mail: [email protected]
Received 3 June 2002; Accepted 22 July 2002
I.C. Baines’s current address is Max-Planck-Institute of Molecular Cell
Biology and Genetics, Pfotenhauerstrasse 108, Dresden, Germany.
S.K. Doberstein’s current address is Xencor Inc., 111 W. Lemon Ave.,
Monrovia, CA 91016.
© 2003 Wiley-Liss, Inc.
30
Ostap et al.
INTRODUCTION
Phagocytosis and macropinocytosis, two forms of
clathrin-independent endocytosis, share some morphological features but differ in their functions and regulation. In phagocytosis, interaction of plasma membrane
receptors with molecules on the surface of a particle
triggers ingestion of the particle [for a review, see Rabinovitch, 1995]. Macropinocytosis is the constitutive,
non-selective uptake of extracellular fluid and solutes in
large vesicles ⬎0.5 ␮m in diameter [for a review, see
Swanson and Watts, 1995]. Both processes depend on
large-scale rearrangements of the plasma membrane and
the underlying actin cytoskeleton to create large intracellular vesicles. However, the molecular mechanisms of
these processes are not understood.
Several studies have implicated actin filaments and
myosins in phagocytosis and macropinocytosis: both actin filaments and multiple myosin-I isoforms concentrate
in the cortex of phagocytic cups and ruffling membranes
[e.g., Fukui et al., 1989; Baines and Korn, 1990; Zhu and
Clarke, 1992; Yonemura and Pollard, 1992; Baines et al.,
1992; Ruppert et al., 1995; Allen and Aderem, 1995;
Baines et al, 1995; Swanson et al., 1999; Schwarz et al.,
2000; Diakonova et al., 2002]; and disruption of actin
filaments [e.g., Holm et al., 1993; Kubler and Riezman,
1993; Lamaze et al., 1997] or deletion of myosin-I genes
[Jung and Hammer, 1990; Titus et al., 1993; Wessels et
al., 1991; Novak et al., 1995; Jung et al., 1996; Geli and
Riezman, 1996; for a review, see Ostap and Pollard,
1996a] inhibit endocytosis. Additionally, deletion of myosin-VII in Dictyostelium inhibits phagocytosis [Titus,
1999] by disrupting particle adhesion [Tuxworth et al.,
2001].
The morphology of endocytosis suggests that the
actin cytoskeleton first drives the extension of the plasma
membrane to surround a particle or throw up ruffles to
surround medium. Later in the process, the actin cytoskeleton may constrict the opening to the outside to
promote membrane fusion required to form the endosome. Actin polymerization is presumed to contribute to
the extension of the plasma membrane, as postulated for
the extension of the leading edge of motile cells [for a
review, see Pollard et al., 2000]. The myosin-I class of
membrane-binding motors may also contribute to the
extension [Jung et al., 2001] as well as to the constriction
phases of endocytosis [Swanson et al., 1999]. Nevertheless, research has not progressed to the point where the
mechanisms of membrane movement and fusion are understood.
The existence of multiple myosin-I isoforms in
cellular slime molds [Novak et al., 1995; Jung et al.,
1989, 1996; Titus et al., 1989], budding yeast [Geli and
Riezman, 1996], amoeba [Lynch et al., 1989; Maruta et
al., 1979], and vertebrates [Sokac and Bement, 2000]
complicates the assignment of physiological functions.
For example, deletion of individual or multiple myosin-I
genes reduces the rates of pinocytosis and phagocytosis,
but does not stop either process [Jung and Hammer,
1990; Titus et al., 1993; Wessels et al., 1991; Novak et
al., 1995; Jung et al., 1996]. It is not known which type
of pinocytosis is inhibited or which steps in either process are faulty in particular mutants. The results do
suggest that multiple myosin-Is participate in endocytosis and that they have partially overlapping, but complementary functions.
Here we report new morphological observations
that address myosin-I dynamics and myosin-I isoform
involvement in macropinocytosis and phagocytosis. We
find that myosin-I concentrates rapidly and transiently
near the plasma membrane at sites of endocytosis. All
three myosin-I isoforms participate in both processes.
The mixture of myosin-I isoforms varies in individual
ruffles ingesting fluid by macropinocytosis, suggesting
heterogeneity in the process.
MATERIALS AND METHODS
Strains
An axenic strain of Acanthamoeba castellanii (Neff
strain) was grown in aerated suspension culture [Pollard
and Korn, 1972] or in plastic tissue culture bottles (Falcon). Adherent cells were harvested by shaking from the
surface. Cells were pelleted at 1,000 –3,000g for 1–3 min
and resuspended in growth medium at the appropriate
density.
Immunoblotting
Monoclonal antibody M1.7 was raised against a
mixture of Acanthamoeba myosin-I isoforms [Hagen et
al., 1986; Kiehart et al., 1984] and purified from ascites
by ammonium sulfate precipitation and ion exchange
chromatography [Kiehart et al., 1984]. Purified Acanthamoeba myosin-IA, -IB, and -IC [Lynch et al., 1991]
were resolved by SDS gel electrophoresis and electroblotted onto nitrocellulose. The blots were blocked with
1% fetal calf serum in phosphate buffered saline (150
mM NaCl, 20 mM Na phosphate, pH 8.0) and probed
with M1.7 tissue culture supernatant for 30 min. The
nitrocellulose was washed three times for 5 min each in
PBS plus 1 mM EDTA and 0.2% Triton X-100, once in
2 M urea, 100 mM glycine, 1% Triton X-100, pH 8.0, for
5 min, and once again in PBS/EDTA/Triton X-100 for 5
min. The bound monoclonal antibody was detected by
incubating the nitrocellulose with a 1:10,000 dilution of
goat-anti-mouse IgG conjugated to horseradish peroxidase (Hyclone Laboratories, Logan, UT) in PBS for 30
min, washed as described above, and developed using the
ECL chemiluminescence detection system (Amersham
Myosin-I Localization
Fig. 1. Immunoblot of purified Acanthamoeba myosin-IA, myosinIB, and myosin-IC probed with the monoclonal antibody M1.7 showing isoform specificity.
Corp., Arlington Heights, IL). M1.7 binds Acanthamoeba myosin-IA and myosin-IC, but not myosin-IB,
on immunoblots (Fig. 1).
Immunofluorescence Microscopy of Fixed Cells
For immunofluorescence microscopy using only
M1.7, or M1.7 and rhodamine phalloidin, we fixed amoebae with formaldehyde and methanol by the method of
Fukui at al. [1986] without agar overlay and stained
according to Yonemura and Pollard [1992]. Staining was
qualitatively similar when amoebae were fixed and
stained using the two-step technique of Yonemura and
Pollard [1992]. When M1.7 was used with polyclonal
isoform-specific antibodies in double-labeling experiments, amoeba were fixed and stained according to
Baines and Korn [1990]. We used (1) Texas-Red-labeled
goat-anti-mouse IgG (Molecular Probes, Eugene, OR)
that was preadsorbed against fixed whole amoebae
[Yonemura and Pollard, 1992], (2) Cy3-labeled goatanti-mouse IgG, or (3) Cy2-labeled goat-anti-mouse IgG
(Amersham Corp., Arlington Heights, IL) to detect
bound monoclonal antibodies. We used fluorescein-conjugated goat-anti-rabbit IgG (Molecular Probes) to detect
bound polyclonal antibodies.
For quantitative analysis of cells double stained
with M1.7 and one of the isoform specific polyclonal
antibodies [Baines et al., 1995], we selected fields of
cells with 1–5 endocytic structures on 10 to 50% of the
cells. We identified endocytic structures by their characteristic ring-shaped fluorescent staining and, where feasible, confirmed the identification by phase contrast microscopy to rule out the misidentification of contractile
vacuoles or digestive vacuoles. Since we used fluorescence to screen for endocytic structures, in experiments
with cells probed with M1.7 and anti-MIA or anti-MIC,
endocytic structures that contained only myosin-IB
would not have been stained and, thus, not included in
the quantitation.
31
To investigate phagocytosis, we added Saccharomyces cerevisiae, previously fixed with 4% formaldehyde and 0.5% glutaraldehyde, in 50 mM NaCl to adherent amoebae. After a 2-min incubation, we processed
the samples for immunofluorescence as described above.
We acquired digital fluorescence and DIC images
of fixed cells using a Zeiss 100⫻, 1.3 NA PlanNeofluar
oil-immersion objective with epi-illumination with a 100
W HBO mercury lamp. A Photometrics PXL cooled
CCD camera with a Kodak KAF 1400 chip was run at
⫺25°C with IP LAB Spectrum acquisition software. For
double staining, we used either (1) a Leitz Orthoplan
microscope with vertical fluorescence illuminator or a
Zeiss Axiovert 135 inverted microscope, 63⫻ objective
and Zeiss LSM410 confocal attachment, or (2) the EPR
3D Deconvolution System (Scanalytics, Fairfax, VA).
Syringe Loading
Antibody M1.7 at a concentration of 65 ␮M in
phosphate buffered saline (PBS; 10 mM NaPO4, 150 mM
NaCl, 1 mM EGTA, pH 7.5) was incubated for 30 min at
25°C with the lysine-reactive, Cy3 monofunctional dye
(Cy3; Biological Detection Systems, Pittsburgh, PA) at
350 ␮M. Unreacted dye was removed by gel filtration
chromatography using a Sephadex G-50-medium column
equilibrated in PBS, pH 7.5. Cy3-M1.7 was dialyzed
extensively against PBS and stored in the dark at 4°C in
0.02% NaN3. Two Cy3-M1.7 preparations were used in
this study; one had a Cy3 to M1.7 ratio of 0.73, and the
second had a ratio of 0.97.
Log phase Acanthamoeba cells grown in aerated
culture were harvested and resuspended at 106 cells ml-1
in 1–5 ␮M Cy3-M1.7 in loading buffer (50 mM alanine,
50 mM proline, 24 mM glutamate, 1 mM EGTA, 2 mM
MgCl2, 40 mM ␥-aminobutyric acid, pH 6.5) [Sinard and
Pollard, 1989]. Cells were syringe-loaded during six
slow passes (three in and three out) through a 30-gauge
needle [Clarke and McNeil, 1992; Doberstein et al.,
1993]. The cells were immediately centrifuged for 1 min
at 1,000g, washed in growth medium, centrifuged again,
and resuspended in growth medium at 105 cells ml-1.
Five to 30% of the cells were loaded with Cy3-M1.7, as
determined by visual inspection or by fluorescence activated cell sorting.
Fluorescence and Differential Interference
Contrast (DIC) Microscopy of Live Cells
Live cells were plated on a glass coverslip in a flow
cell [Berg and Block, 1984] and allowed to attach for
5–15 min. The cells were perfused at 4 ml/h with airequilibrated Acanthamoeba growth medium for 30 min
before and during microscopic observation to prevent
oxygen depletion.
DIC images of live cells were acquired with a Zeiss
Axiovert 135 microscope using a Zeiss 100⫻, 1.3 NA
32
Ostap et al.
PlanNeofluar oil-immersion objective and DIC optics.
The images were magnified by a 1.6⫻ optovar and a
4.0⫻ TV lens and collected with a Newvicon camera
(C2400-07; Hamamatsu Photonics, Bridgewater, NJ),
and processed in real time for background subtraction,
and contrast enhancement using a Hamamatsu Argus 10
image processor. Images were recorded on S-VHS tape
and digitized using NIH Image.
Fluorescence images were acquired with a cooled
CCD camera and the Zeiss Axiovert 135 microscope
using an exposure of 0.1 to 0.3 s, based on the fluorescence intensity of the cell. To minimize photodamage, a
computer-controlled shutter limited fluorescence excitation to the sample to the time of image acquisition.
Myosin-I ATPase Activity and In Vitro Motility
The steady-state ATPase rate of 70 nM phosphorylated myosin-IA plus or minus 700 nM Cy3-M1.7 was
determined in low ionic strength buffer (2 mM ATP, 2
mM MgCl2, 1 mM EGTA, 5 mM KCl, 1 mM DTT, 15
mM imidazole, pH 7.5) in the presence of 75 ␮M actin
[Pollard, 1982]. The presence of Cy3-M1.7 did not
change the actin-activated ATPase rate of phosphorylated MIA (7 s-1), confirming previous measurements
with unlabeled M1.7 [Hagen et al., 1986].
Actin filament gliding in the in vitro motility assay
[Kron et al., 1991; Zot et al., 1992; Bresnick et al., 1996]
was used to assess the effect of M1.7 on the motor
activity of myosin-IC. M1.7 was added to phosphorylated myosin-IC bound to the nitrocellulose coverslip
after blocking with 5 mg/ml casein. The rate of filament
gliding in the absence of M1.7 was 0.09 ⫾ 0.03 ␮m s-1,
and the rate of filament gliding in the presence of M1.7
was 0.18 ⫾ 0.03 ␮m s-1.
Scanning Electron Microscopy
Cells at a density of 105 cells/ml of culture medium
were allowed to attach to sterile glass coverslips (cleaned
with ethanol and acetone) for 10 min at room temperature. After a 30-min incubation in 50 mM NaCl at room
temperature, adherent cells were fixed with 3% glutaraldehyde in 100 mM sodium cacodylate, 5% sucrose, 10
mM CaCl2, pH 6.8, for 1 h at room temperature. After 3
washes in 100 mM sodium cacodylate, 5% sucrose, 10
mM CaCl2, pH 6.8, for 10 min each, cells were treated
for 1 h with 2% OsO4 in 100 mM sodium cacodylate,
washed twice with deionized water for 5 min each and
dehydrated by successive 5-min washes in 25, 50, 70,
and 90% ethanol followed by three 10-min washes in
100% ethanol. Dehydrated cells were dried in a Balzers
CPD-10 critical point dryer (Balzers-Union, Liechtenstein) with liquid CO2, and the coverslips were stored in
desiccators until sputter coated with gold for 60 s at 20
mA using either a Polaron E5100 SEM coating unit or a
Denton Vacuum DESK II sputter coating unit. The sam-
ples were observed and photographed in an Amray 1810
scanning electron microscope (Amray Electron Optics,
Bedford MA) using Polaroid Type 55 film.
Transmission Electron Microscopy
Amoebae were plated onto a surface prepared so
that they could be observed by fluorescence microscopy,
photographed, positionally mapped, embedded in epoxy,
rephotographed, and thin sectioned for transmission electron microscopy. A grid for mapping the position of
amoebae was created by evaporating a thin layer of
carbon over a 50 mesh copper EM grid that was placed
on the glass surface of a MatTek dish (35 mm dish with
a no. 0 glass coverslip glued under a 1.5-cm diameter
hole cut in the bottom of the dish; MatTek, Ashland,
MA). The EM grid was removed and the petri dish was
sterilized by washing with 70% ethanol. Amoebae from
suspension culture were plated and maintained with 9 ml
of fresh media daily for 1–2 days.
Amoebae were fixed in 1% glutaraldehyde in 20%
Dulbecco’s PBS (DPBS) with 0.2% Triton X-100 for 15
min at room temperature, rinsed twice in DPBS, blocked
with 0.1% BSA in DPBS (PBS-BSA) for 10 min, incubated with rhodamine phalloidin (Molecular Probes) in
PBS-BSA, and rinsed twice in PBS-BSA. Amoebae of
interest were located, mapped on the grid, and photographed using an Olympus IMT-2 inverted microscope
with phase and fluorescence optics with a 100⫻ Olympus objective.
In preparation for electron microscopy, dishes were
rinsed with DPBS twice, fixed with 2% glutaraldehyde
and 0.4% tannic acid in DPBS for 15 min at room
temperature, rinsed 2 ⫻ 5 min with DPBS, fixed with
0.1% osmium tetroxide in DPBS for 15 min, rinsed with
3 ⫻ 3 min of DPBS, and dehydrated with a series of
ethanol concentrations (50, 70, 95, and three changes of
100%) for 10 min each. Three layers of Epon were
applied. Each application was separated by an hour. The
Epon was allowed to cure for 48 h at 60°C. The amoebae
of interest were located on the carbon grid and were
photographed using transmitted light. The embedded
amoebae were removed from the culture dish by freezing
the sample in liquid nitrogen.
Selected amoebae were located within the carbon
grid lines by bright field microscopy, marked, excised,
and remounted on Epon blanks. Serial 70-nm sections
were collected (LKB Ultramicrotome), stained with
0.1% lead citrate, and examined with either a
JEOL100CX or a Zeiss EMIOA electron microscope.
Features of interest were traced onto clear sheets and
digitized. Stereo renderings were produced with NIH
Image, version 1.6.
Myosin-I Localization
33
Fig. 2. Left: DIC micrographs immunofluorescence micrographs of
three different Acanthamoeba fixed while undergoing fluid-phase endocytosis and labeled with (center) rhodamine phalloidin and (right)
M1.7 and Cy2 conjugated second antibody. Scale bar ⫽ 5 ␮m.
RESULTS
Dynamic Localization of Myosin-I and Actin to
Macropinocytic Structures
Actin and myosin-IA and/or myosin-IC concentrate
in the cortex of cells adjacent to two types of plasma
membrane specializations: (1) invaginations formed by
large membrane ruffles (Fig. 2, rows 1 and 2); and (2)
smaller cups at the end of stalk-like membrane regions
(Fig. 2, row 3). The morphology of these structures
suggests that they participate in fluid-phase endocytosis
producing vesicles larger than 0.5 ␮m. The M1.7 monoclonal antibody used to identify myosin-I also concentrates around contractile vacuoles, structures of similar
size [Yonemura and Pollard, 1992]. However, structures
like those in Figure 2 are not contractile vacuoles, since
they do not react with monoclonal antibody N2 [Fok et
al., 1993] specific for the vacuolar proton ATPase (VATPase) found in contractile vacuoles [Fok et al., 1993]
(data not shown).
To correlate unambiguously the morphological features observed in fixed cells with specific physiological
events, we localized myosin-I in live cells with Cy3M1.7 (Figs. 3 and 4).We introduced the fluorescent antibody into living Acanthamoeba by syringe loading.
This antibody binds tightly to myosin-IA and -IC (Kd ⫽
0.2 nM) [Hagen et al., 1986] without inhibiting the
actin-activated ATPase or actin gliding in the in vitro
motility assay (see Methods). We used this indirect labeling approach because labeling myosin-IA and myosin-IC directly with lysine- or sulfhydryl-directed fluorescent dyes compromised the actin-activated ATPase.
Most of the Cy3-M1.7 loaded into live cells spread
diffusely through the cytoplasm (Figs. 3 and 4). The
Fig. 3. Time course of macropinocytosis illustrated by fluorescence
micrographs of a live Acanthamoeba syringe loaded with Cy3-M1.7.
Cy3-M1.7 concentrated near the membrane and in the cortex of the
dynamic ruffle. The amoeba was migrating towards the top and to the
right. The time after the first micrograph is given in seconds. Bottom
right: Scanning electron micrograph of a fixed amoeba with a similar
endocytic structure. Scale bar ⫽ 5 ␮m.
Fig. 4. Fluorescence micrographs of a live Acanthamoeba syringeloaded with Cy3-M1.7 performing endocytosis. Cy3-M1.7 concentrated around the macropinocytic cup at the end of the stalk-like
pseudopod (arrow). The time after the first micrograph is given in
seconds. Bottom right: Scanning electron micrograph of a fixed
amoeba with a similar endocytic structure (arrow). Scale bar ⫽ 5 ␮m.
following evidence suggests that the cytoplasmic Cy3M1.7 was bound to myosin-IA and/or myosin-IC rather
than free. At the concentration of Cy3-M1.7 (5 ␮M) used
in the loading buffer, the intracellular concentration of
34
Ostap et al.
TABLE I. Quantitative Analysis of Myosin I Isoform Distribution in Macropinocytic Structures
That Stain With Myosin I Antibodies*
Structures stained by (%)
Experiment
1
2
3
Antibodies
Isoform specific
antibody only
M1.7 only
Both antibodies
Anti-MIA, M1.7
Anti-MIB, M1.7
Anti-MIC, M1.7
NA
29
NA
40
52
25
60
19
75
*The analyses were performed on single cells or groups of cells selected specifically for the presence of macropinocytic structures. Experiment
1 had 25 total cells containing 50 macropinocytic structures as identified by staining with antibodies; experiment 2 had 21 cells containing 52
macropinocytic structures; and experiment 3 had 30 cells containing 76 macropinocytic structures. NA, Any structure that stained with anti-MIA
or anti-MIC would also stain with M1.7 because this antibody recognizes both MIA and MIC. From experiment 2: 29% of total (antibody
positive) macropinocytic structures contain only myosin-IB; 19% contain myosin I-B plus myosin-IA/IC; 52% contain no myosin-IB; and 71%
contain myosin-IA/IC. From experiments 1 and 2: 28% (40% of 71%) of total structures contain myosin-IC but not myosin-IA; and 43% (60%
of 71%) contain myosin-IA. From experiments 3 and 2: 18% (25% of 71%) of total structures contain myosin-IA but not myosin-IC; and 53%
(75% of 71%) contain myosin-IC. Therefore, 25% of total structures (71–28 –18%) contain both myosin-IA and myosin-IC.
M1.7 should be less than half of the intracellular concentration of myosin-IA and -IC [Doberstein et al., 1993;
Baines et al., 1992]. Varying the concentration of Cy3M1.7 in the loading buffer between 0.5 and 5.0 ␮M did
not change the fluorescence distribution as one would
expect if the antibody concentration exceeded the myosin-I concentration. Also, fluorescence-recovery-afterphotobleaching (FRAP) experiments showed the diffusion of cytoplasmic Cy3-M1.7 (D ⫽ 10-9 cm2 s-1) was
significantly slower than expected for free antibody (D ⬎
10-8 cm2 s-1; data not shown). These observations are
consistent with quantitative immune-electron microscopy results that showed most myosin-IA is in the cytoplasm [Baines et al., 1995].
Amoeba, like other cells, use dynamic plasma
membrane ruffles to internalize large vacuoles containing
the extracellular medium (Fig. 3). Each dynamic membrane ruffle extended from the cell surface, became circular as it folded back and fused upon itself to form an
endosome (Fig. 3, 0 –12 s). The top of the circular ruffle
appeared to close around the endosome with a pursestring or contractile-ring-like motion (Fig. 3, 12– 42 s).
Within 3–5 s after the internalization of the macropinosome, a new ruffle formed at the same site, and the
process repeated. Acanthamoeba form only one of these
large endocytic structures at a time, usually near the
trailing end of migrating cells. In scanning electron micrographs (Fig. 3, bottom right), these circular membrane
ruffles are similar to actin-rich endocytic structures
called crowns in Dictyostelium [Hacker et al., 1997] and
kidney cell lines [Dowrick et al., 1993]. Scanning (data
not shown) and transmission electron micrographs [Bowers and Korn, 1968; Ryter and Bowers, 1976] of amoebae
fixed while growing in suspension culture do not show
these large endocytic structures.
Cy3-M1.7 concentrated near the plasma membrane
of ruffles that take in medium by macropinocytosis (Fig.
3). Despite some photobleaching of the Cy3 (Fig. 3,
0 –12 s), the fluorescence intensity increased as the ruffle
enclosed the endosome (Fig. 3, 18 – 42 s). This fluorescence increase is most likely due to concentration of
myosin-I in the cortex as the ruffle’s cytoskeleton contracted. The fluorescence dissipated from the macropinosome within 20 s of internalization, as determined by
examining multiple focal planes.
Cy3-M1.7 also concentrated near the plasma membrane of smaller endocytic structures at the end of one or
more stalk-like membrane pseudopods on stationary cells
(Fig. 4, arrows). Endosomes formed by cupping of the
plasma membrane at the end of the pseudopods and
moved toward the cell center at about 0.2– 0.3 ␮m per
second (Fig. 4). Multiple endocytic events can occur
serially at the end of such a stalk-like region. Cy3-M1.7
remained concentrated around the newly formed endosome just after the endocytic cup pinched from the cell
surface (Fig. 4). Once an endosome entered the cell body,
any associated fluorescence was obscured by the high
fluorescence intensity of the cytoplasm. A scanning electron micrograph (Fig. 4, bottom right) shows that such a
stalk-like region can extend 4 – 8 ␮m from the cell body,
terminating in a ruffled region that may be responsible
for dynamic endosome creation.
Myosin-I Isoforms Are Differentially Localized in
Macropinocytosis
Double staining cells with M1.7, as a marker for
macropinocytic structures that contain myosin-IA and
-IC, and polyclonal antibodies specific for either myosinIA, -IB, or -IC [Baines et al., 1995] revealed that large
membrane ruffles involved in macropinocytosis are heterogeneous with respect to their myosin-I antibody staining. As expected, all macropinocytic structures stained
positively with either or both M1.7 and anti-myosin-IB
and all of the macropinocytic structures that stained with
either anti-myosin-IA or anti-myosin-IC also stained
with M1.7 (Table I).As described in Table I, 18% of all
Myosin-I Localization
35
Fig. 6. Fluorescence micrographs of a live Acanthamoeba syringeloaded with Cy3-M1.7 shown during phagocytosis of a yeast. The
arrow shows the initial binding site of the yeast particle, since it is not
visible in the fluorescence micrograph. The time after the first micrograph is given in seconds.
Fig. 5. DIC (left) and immunofluorescence (right) micrographs of
Acanthamoebae fixed (top, middle) while ingesting yeast (Y) and
(bottom) while ingesting each other. Cells were labeled with M1.7 and
Cy3 conjugated second antibody. Scale bars ⫽ 5 ␮M.
endocytic structures detected with myosin-I antibodies
contained myosin-IA, but not myosin-IB, 28% contained
myosin-IB but not myosin-IA, 25% contained both myosin-IA and myosin-IC, 29% contained myosin-IB only,
and 19% contained both myosin-IB and myosin-IA
and/or myosin-IC.
Myosin-I and Actin Localize to Phagocytic
Structures
M1.7 concentrated around the phagocytic cup and
early phagosomes (Fig. 5) during the ingestion of yeast.
Internalized yeast in phagosomes separated from the
plasma membrane did not strongly stain for myosin-I
(Fig. 5, middle). However, punctate staining around
some internalized structures is apparent.
We followed the time course of myosin-I redistribution during phagocytosis with syringe-loaded Cy3M1.7 (Fig. 6). Binding of the yeast to the plasma membrane initiated phagocytosis (Fig. 6, arrow). Within 10 s,
Cy3-M1.7 concentrated at the point of yeast contact.
During the next 60 s, Cy3-M1.7 continued to concentrate
in the cortex surrounding the yeast as it is internalized.
Within 2 min, the fluorescence around the internalized
phagosome was indistinguishable from the background
fluorescence.
Fig. 7. DIC micrographs of a live Acanthamoeba showing the dynamics of cannibalistic phagocytosis. The upper amoeba was attempting to phagocytose a pseudopod of the lower amoeba. The internalized
membrane of the lower amoeba appeared to be extending an endocytic
cup within the top amoeba. As the amoebae pulled apart, membrane
tethers broke between the amoebae. These cell-to-cell contacts are
common between amoebae crowed together on a solid substrate.
Bottom right: Transmission electron micrograph of a thin section
showing the cortical actin filaments excluding organelles in the zone of
contact between two cells with cell-to-cell contacts similar to those
observed in the live cells.
In high-density cultures, amoebae ingest parts of
their neighbors. This cannibalistic behavior is initiated
when a “victim” amoeba extends a pseudopod that
touches a second amoeba (Fig. 7). In most cases, the
pseudopod is withdrawn without provoking an attack, but
occasionally the second amoebae becomes an aggressor.
36
Ostap et al.
Fig. 8. Electron micrographs of thin sections showing cannibalistic phagocytosis. A–F: Tracings of sequential sections of a high-density culture of Acanthamoeba showing the high concentration of actin filaments near
the membranes of (black stipple) phagocytic and (black) “victim” amoebae. One of these amoebae is both a
perpetrator and victim of phagocytosis. G: Electron micrograph of a thin section of the amoebae used for the
tracings (tracing B). The space between the cells is due to shrinkage during preparation for thin sectioning. The
plasma membranes are outlined in black for clarity. Scale bar ⫽ 10 ␮M.
It surrounds the victim’s membrane projection with a
phagocytic cup and actively pulls on the membrane (Fig.
7). In high-density cultures, single amoeba may be simultaneously phagocytic and a victim of phagocytosis
(Fig. 8). These sucker-like structures of Acanthamoeba
and Naegleria fowleri (a pathogenic free-living ameboflagellate) have been called amoebastomes [John et al.,
1984, 1985; Diaz et al., 1991).
Phagocytic structures labeled intensely with rhodamine phalloidin and M1.7 (Fig. 5, bottom row; Figs. 9
and 10). Electron micrographs of thin sections showed a
high concentration of actin filaments around the phagocytic membrane (Figs. 8 and 10). Actin filaments concentrated in three areas around the membranes of large
phagocytic structures (Fig. 10, left): the tips of the phagocytic cup (Fig. 10, left, top arrow); the very bottom of the
structure (Fig. 10, left, bottom arrow); and around the
middle of the invagination in a contractile-ring like structure (Fig. 10, asterisks). Aggressor amoebae can amputate and ingest a whole pseudopod from a victim amoeba
(Fig. 11) indicating that the phagocytic structure can
adhere to and exert considerable force upon neighboring
cells.
The pseudopods of the victim also contain high
concentrations of myosin-I and actin filaments (Fig. 5,
bottom row, Figs. 9 and 10) during and after ingestion by
the aggressor (Fig. 11). We do not know if the high
concentration of the contractile proteins in the victim’s
membrane is a response to being phagocytosed (possibly
a response to tension), or if these proteins participated in
the initial extension of the pseudopod. We could not
observe such cannibalistic phagocytic events with living
Acanthamoeba syringe-loaded with Cy3-M1.7. Cells
must be plated at high density for 6 –12 h before a
significant number of cell-cell phagocytic structures develop. On this time scale, Cy3-M1.7 is concentrated in
vesicles, probably targeted for digestion.
DISCUSSION AND CONCLUSIONS
Role of Myosin-I and Actin in Endocytosis
The plasma membrane and cortex surrounding
large macropinocytic and phagocytic structures in Acanthamoeba are enriched in myosin-I and actin filaments
(this study), subunits of the Arp2/3 complex [Kelleher et
al, 1995; Mullins et al., 1997], actophorin [Quirk et al.,
1993], and alpha-actinin (Kaiser and Ostap, unpublished
observations). All of these proteins likely participate in
the extension and withdrawal of these structures, as well
as the formation and transport of newly formed endosomes. The remarkably similar morphology and protein
Myosin-I Localization
37
retraction of the endocytic structures might result either
from the catastrophic depolymerization of supporting
actin filaments or from the active force produced by
myosin-I [Novak et al., 1995; Tang and Ostap, 2002].
Myosin-IB associated with the plasma membrane is
phosphorylated (and as a consequence activated) at sites
of membrane protrusions (such as during the formation
of pseudopods, filopodia, or endocytic cups), while phosphorylated myosin-IA is recruited into the actin-rich cortical cytoplasm at the same sites [Baines et al., 1995].
Thus, the localization of phosphomyosin-IB is consistent
with a role in membrane dynamics, including membrane
protrusion, membrane retraction, endosome formation,
and vesicle motility. The localization of phosphomyosin-IA is consistent with mediating movement of cytoplasm into or out of the nascent protrusion, or functioning as a regulator of the elastic modulus of the cortical
actin network, influencing the extent to which the cortex
can change shape [Condeelis et al., 1988; Jung et al.,
1993; Ostap and Pollard, 1996a,b].
Heterogeneity of Myosin-I Distribution
Fig. 9. Cannibalistic phagocytosis. Top: Scanning electron micrograph. Bottom: Confocal fluorescence micrograph of amoebae labeled
with (green) bodipy-phalloidin and (red) Cy3-M1.7.
compositions of the macropinocytic structures and the
phagocytic structures (this study) [Baines et al., 1992,
1995] suggest that cytoskeletal participation in both macropinocytosis and phagocytosis is similar, in spite of the
fact that these processes are regulated differently [Swanson and Baer, 1995].
Since myosin-II [Yonemura and Pollard, 1992],
kinesin (data not shown), and dynein (data not shown)
are excluded from the endocytic structures, it is likely
that the force for extension is produced by either actin
polymerization, as proposed for the extension of the
leading edge of motile cells and filopodia [Pollard et al.,
2000], or by the action of cortical myosin-I (or unidentified members of the myosin family) on actin filaments.
The formation and transport of the endosome may also be
mediated by membrane-bound and cortical myosin-I.
The contractile-ring-like structure of actin filaments seen
surrounding the phagocytic membranes suggests a role
for myosin-I and actin in endosome creation. Similarly,
All three known Acanthamoeba myosin-I isoforms
concentrate in macropinocytic structures, but none is
detected in all of these structures (Table I). We considered possible explanations for this heterogeneity. (1) All
three isoforms concentrate in all macropinocytic structures but their accessibility to the antibody varies as these
endocytic structures mature. (2) The myosin-I isoforms
accumulate at different times during the formation and
withdrawal of all of the structures. (3) The myosin-I
composition of endocytic structures is heterogeneous.
The first two explanations are less likely, because
Cy3-M1.7 accumulates continuously around endocytic
structures in live cells. Although the fluorescence distribution of the Cy3-M1.7 within the structure changes
(Figs. 3 and 4), the Cy3-M1.7 never leaves the structure
during the lifetime of an endocytic event (30 – 60 s).
Myosin-I heterogeneity seems the more likely explanation, since the endocytic structures themselves are
heterogeneous with two types for macropinocytosis
(Figs. 3 and 4) and one involved in phagocytosis (Fig. 6).
Specific combinations of myosin-I isoforms may be required for each of these endocytic events. However, it is
also possible that myosin-I isoform expression levels
vary from cell to cell resulting in apparent differences in
protein heterogeneity. We have not yet tested these hypotheses by quantitating myosin-I isoforms in the different endocytic structures, because these structures are
difficult to distinguish with certainty in fixed cells compared with live cells or scanning electron microscopy.
The heterogeneity of myosin-I localization to the
endocytic structures may help explain the phenotypes of
Dictyostelium mutants lacking multiple myosin-I isoforms. These mutants have impaired fluid-phase endocy-
38
Ostap et al.
Fig. 10. Cannibalistic phagocytosis. Left: Electron micrograph of a
thin section of an amoeba phagocytosing a pseudopod of a “victim”
amoeba. Plasma membranes are highlighted with a black line. Inset:
Fluorescence micrograph of the same field. The cells were labeled with
rhodamine-phalloidin before the sample was processed for electron
microscopy. Three areas of high actin concentration are shown (ar-
rows and asterisk). Center and right: Stereo rendering of tracings of
16 serial sections tilted ⫺20° and ⫹20°, respectively. The victim
amoeba is shown in white. The phagocytic amoeba is black, but
outlined in white. Actin filaments in the aggressor amoeba are drawn
as white stipple. Scale bar ⫽ 3 ␮m.
Ostap and Pollard, 1996a]. However, no combination of
losses of three out of the six known myosin-Is completely abolished pinocytosis and phagocytosis. These
defects in endocytosis do not appear to result from the
inhibition of endosome or lysosome transport and processing, but are most likely due to components of the
internalization step in the actin-rich cell cortex [Temesvari et al., 1996]. Therefore, complementary internalization mechanisms may contribute to endocytosis with
multiple myosin-I isoforms participating in each pathway.
Substrate Attachment and Endocytosis
Fig. 11. a, b: Electron micrographs with (c, d) corresponding fluorescence micrographs of amoebae stained with rhodamine-phalloidin
showing ingested pseudopods from victim amoebae. Plasma membranes are outlined in black. Scale bar ⫽ 3 ␮m.
tosis and phagocytosis. Single, double, and triple myosin-I knockouts progressively inhibit the rate of
pinocytosis, and the losses of activity in the multiple
knockouts are greater than the sum of individual losses
[Jung et al., 1996; Novak et al., 1995; for a review, see
Since neither we nor previous investigators [Baines
et al., 1992; Baines and Korn, 1990; Bowers and Korn,
1968; Ryter and Bowers, 1976] observed large ruffles on
Acanthamoeba fixed in suspension, substrate attachment
may stimulate macropinocytosis. Acanthamoeba grown
in suspension culture have numerous endosomes 120 nm
in diameter, a size consistent with clathrin-mediated endocytosis, and few endosomes 0.2–2.5 ␮m in diameter,
consistent with macropinocytosis [Bowers and Olszewski, 1972].
This dependence of macropinocytosis on adherence
may be related to the phenotypes of Dictyostelium lines
with myosin-I null mutations. Dictyostelium double-
Myosin-I Localization
knockout mutants (myoA⫺/myoB⫺ and myoB⫺/myoC⫺)
overcome their defects in pinocytosis when attached to a
substrate [Novak et al., 1995]. One explanation is that
these myosin-I isoforms are not required for a subset of
pinocytic mechanisms that are activated by substrate
attachment, such as the macropinocytic ruffles of Acanthamoeba. In contrast, Dictyostelium mutants with another set of null mutations of one (myoB⫺), two
(myoB⫺/ myoD⫺), or three (myoB⫺/ myoC⫺/myoD⫺)
myosin-I genes [Jung et al., 1996] did not recover from
their defects in pinocytosis on a solid substrate. Thus, a
process like ruffling macropinocytosis may depend on a
specific subset of myosin-I isoforms [Ostap and Pollard,
1996a]. More detailed analysis will be required to sort
out the participation of particular combinations of myosin-I isoforms in endocytosis.
Heterophagy
A striking finding is that Acanthamoeba cells
grown at high density undergo heterophagy (phagocytosis of neighboring cells). Heterophagy is known to occur
in metazoan cells as a response to developmental cell
death [Clarke, 1990] and in specialized tissues, like the
phagocytosis of photoreceptor outer segments by the
retinal pigment epithelium [Bok, 1993]. However, it remains to be determined if Acanthamoeba heterophagy is
an artifact of high-cell-density culture growth, or if it has
a physiological purpose (e.g., clearing of unhealthy cells
or transfer of genetic material).
ACKNOWLEDGMENTS
We thank D.A. Kaiser (Salk Institute) for purifying
M1.7 and M. Delannoy (Johns Hopkins) for help with
scanning electron microscopy. We are very grateful to
D. B. Murphy (Johns Hopkins) and the Sanger Laboratory (University of Pennsylvania) for helpful discussions
and for the use of their microscope. We thank Dr. A. Fok
(University of Hawaii) for monoclonal antibody, N2.
This work was supported by a grant from the NIH
(GM-26132) to T.P. E.M.O. was supported by grants
from the Cancer Research Fund of the Damon RunyonWalter Winchell Foundation Fellowship (DRG-1294)
and NIH (GM-57247).
REFERENCES
Allen LH, Aderem A. 1995. A role for MARCKS, the alpha isozyme
of protein kinase C and myosin I in zymosan phagocytosis by
macrophages. J Exp Med 182:829 – 840.
Baines IC, Korn ED. 1990. Localization of myosin IC and myosin II
in Acanthamoeba castellanii by indirect immunofluorescence
and immunogold electron microscopy. J Cell Biol 111:1895–
1904.
Baines IC, Brzeska H, Korn ED. 1992. Differential localization of
Acanthamoeba myosin I isoforms. J Cell Biol 119:1193–1203.
39
Baines IC, Corigliano-Murphy A, Korn E.D. 1995. Quantification and
localization of phosphorylated myosin I isoforms in Acanthamoeba castellanii. J Cell Biol 130:591– 603.
Berg HC, Block SM. 1984. Miniature flow cell designed for rapid
exchange of media under high-power microscope objectives.
J Gen Microbiol 130:2915–2920.
Bok D. 1993. The retinal pigment epithelium: a versatile partner in
vision. J Cell Sci Suppl 17:189 –195.
Bowers, B, Korn ED. 1968. The fine structure of Acanthamoeba
castellanii. I. The trophozoite. J Cell Biol 39:95–111.
Bowers B, Olszewski, TE. 1972. Pinocytosis in Acanthamoeba castellanii: Kinetics and morphology. J Cell Biol 53:681– 694.
Bresnick, AR, Wolff-Long VL, Baumann O, Pollard TD. 1986. Phosphorylation on threonine-18 of the regulatory light chain dissociates the ATPase and motor properties of smooth muscle
myosin II. Biochemistry 34:12576 –12583.
Clarke MS, McNeil PL. 1992. Syringe loading introduces macromolecules into living mammalian cell cytosol. J Cell Sci 102:533–
541.
Clarke PGH. 1990. Developmental cell death: morphological diversity
and multiple mechanisms. Anat Embryol 181:195–213.
Condeelis J, Hall A, Bresnick AR, Warren V, Hock R, Bennet H,
Ogihara S. 1988. Actin polymerization and pseudopod extension during amoeboid chemotaxis. Cell Motil Cytoskeleton
10:77–90.
Diakonova M, Bokoch G, Swanson JA. 2002. Dynamics of cytoskeletal proteins during Fc␥ receptor-mediated phagocytosis in
macrophages. Mol Biol Cell 13:402– 411.
Diaz J, Osuna A, Rosales MJ, Cifuentes J, Mascaro C. 1991. Suckerlike structures in two strains of Acanthamoeba: scanning electron microscopy study. Int J Parisitol 21:365–367.
Doberstein SK, Baines IC, Wiegand G, Korn ED, Pollard TD. 1993.
Inhibition of contractile vacuole function in vivo by antibodies
against myosin-I. Nature 356:841– 843.
Dowrick P, Kenworthy P, McCann B, Warn R. 1993. Circular ruffle
formation and closure lead to macropinocytosis in hepatocyte
growth factor/scatter factor-treated cells. Eur J Cell Biol 61:
44 –53.
Fok AK, Clarke M, Ma L, Allen, RD. 1993. Vacuolar H-ATPase of
Dictyostelium discoideum. J Cell Sci 106:1103–1113.
Fukui Y, Yumura S, Yumura TK, Mori H. 1986. Agar overlay method:
high resolution immunofluorescence for the study of the contractile apparatus. Methods Enzymol 134:573–580.
Fukui, Y, Lynch TJ, Brzeska H, Korn ED. 1989. Myosin I is located
at the leading edges of locomoting Dictyostelium amoebae.
Nature 341:328 –331.
Geli MI, Riezman H. 1996. Role of type I myosins in receptormediated endocytosis in yeast. Science 272:533–535.
Hacker U, Albrecht R, Maniak M. 1997. Fluid-phase uptake by macropinocytosis in Dictyostelium. J Cell Sci 110:105–112.
Hagen SJ, Kiehart DP, Kaiser DA, Pollard TP. 1986. Characterization
of monoclonal antibodies to Acanthamoeba myosin-I that
cross-react with both myosin-II and low molecular weight
nuclear proteins. J Cell Biol 103:2121–2128.
Holm PK, Hansen SH, Sandvig K, van Deurs B. 1993. Endocytosis of
desmosomal plaques depends on intact actin filaments and
leads to a nondegradative compartment. Eur J Cell Biol 62:
362–371.
John DT, Cole Jr TB, Narciano-Cabral FM, 1984. Sucker-like structures on the pathogenic amoeba Naegleria fowleri. App Env
Micro 47:12–14.
John DT, Cole TB, Bruner RA. 1985. Amoebostomes of Naegleria
fowleri. J Protozool 32:12–19.
40
Ostap et al.
Jung G, Hammer III JA. 1990. Generation and characterization of
Dictyostelium cells deficient in a myosin-I heavy chain isoform. J Cell Biol 110:1955–1964.
Jung G, Saxe CL, Kimmel AR, Hammer JA. 1989. Dictyostelium
discoideum contains a gene encoding a myosin-I heavy chain.
Proc Natl Acad Sci USA 86:6186 – 6190.
Jung G, Fukui Y, Martin B, Hammer III JA. 1993. Sequence, expression pattern, intracellular localization, and targeted disruption
of the Dictyostelium myosin ID heavy chain isoform. J Biol
Chem 268:14981–14990.
Jung G, Wu X, Hammer III JA. 1996. Dictyostelium mutants lacking
multiple classic myosin I isoforms reveal combinations of
shared and distinct functions. J Cell Biol 133:305–323.
Jung G, Remmert K, Wu X, Volosky JM, Hammer III JA. 2001. The
Dictyostelium CARMIL protein links capping protein and the
Arp2/3 complex to type I myosins through their SH3 domains.
J Cell Biol 153:1479 –1497.
Kelleher JF, Atkinson SJ, Pollard TD. 1995. Sequences, structural
models, and cellular localization of the actin-related proteins
Arp2 and Arp3 from Acanthamoeba. J Cell Biol 131:385–97.
Kiehart DP, Kaiser DA, Pollard TD. 1984. Monoclonal antibodies
demonstrate limited structural homology between myosin
isozymes from Acanthamoeba. J Cell Biol 99:1002–1014.
Kron SJ, Toyoshima YY, Uyeda TQ, Spudich JA. 1992. Assays for
actin sliding movement over myosin-coated surfaces. Methods
Enzymol 196:399 – 416.
Kubler E, Riezman, H. 1993. Actin and fimbrin are required for the
internalization step of endocytosis in yeast. EMBO J 12:2855–
2862.
Lamaze C, Fujimoto LM, Yin HL, Schmid SL. 1997. The actin
cytoskeleton is required for receptor-mediated endocytosis in
mammalian cells. J Cell Biol 272:20332–20335.
Lynch TJ, Brzeska H, Miyata H, Korn ED. 1989. Purification and
characterization of a third isoform of myosin I from Acanthamoeba castellanii. J Biol Chem 264:19333–19339.
Lynch TJ, Brzeska H, Baines IC, Korn ED. 1991. Purification of
myosin-I and myosin-I heavy chain kinase for Acanthamoeba
castellanii. Methods Enzymol 196:12–23.
Maruta H, Gadasi H, Collins JH, Korn ED. 1979. Multiple forms of
Acanthamoeba myosin-I. J Biol Chem 254:3624 –3630.
Mullins RD, Stafford WF, Pollard TD. 1997. Structure, subunit topology, and actin-binding activity of the Arp2/3 complex from
Acanthamoeba. J Cell Biol 136:331– 43.
Novak KD, Peterson MD, Reedy MC, Titus MA. 1995. Dictyostelium
myosin I double mutants exhibit conditional defects in pinocytosis. J Cell Biol 131:1205–1221.
Ostap EM, Pollard TD. 1996a. Overlapping functions of myosin
isoforms? J Cell Biol 133:221–224
Ostap EM, Pollard TD. 1996b. Biochemical kinetic characterization of
the Acanthamoeba myosin-I ATPase. J Cell Biol 132:1053–
1060.
Pollard TD. 1982. Assays for myosin. Methods Enzymol 85:123–130.
Pollard TD, Blanchoin L, Mullins RD. 2000. Molecular mechanisms
controlling actin filament dynamics in nonmuscle cells. Annu
Rev Biophys Biomol Struct 29:545–576.
Pollard TD, Korn ED. 1972. A protein cofactor required for the actin
activation of a myosin-like ATPase of Acanthamoeba castellanii. Fed Proc 31:502.
Quirk S, Maciver SK, Ampe C, Doberstein SK, Kaiser DA, VanDamme J, Vandekerckhove JS, Pollard TD. 1993. Primary
structure of and studies on Acanthamoeba actophorin. Biochemistry 32:8525– 8533.
Rabinovich, M. 1995. Professional and non-professional phagocytosis:
an introduction. Trends Cell Biol 5:85– 87.
Ruppert C, Godel J, Muller RT, Kroschewski R, Reinhard J, Bähler M.
1995. Localization of the rat myosin I molecules myr1 and
myr2 and in vivo targeting of their tail domains. J Cell Sci
108:3775–3786.
Ryter A, Bowers B. 1976. Localization of acid phosphatase in Acanthamoeba castellanii with light and electron microscopy during
growth and after phagocytosis. J Ultrastruct Res 57:309 –321.
Schwarz EC, Neuhaus EM, Kistler C, Henkel AW, Soldati T. 2000.
Dictyostelium myosin IK is involved in the maintenance of
cortical tension and affects motility and phagocytosis. J Cell Sci
113:621– 633.
Sinard JH, Pollard TD. 1989. Microinjection into Acanthamoeba castellanii of monoclonal antibodies to myosin-II slows but does
not stop cell locomotion. Cell Motil Cytoskeleton 12:42–53.
Sokac AM, Bement WM. 2000. Regulation and expression of metazoan unconventional myosins. Int Rev Cytol 200:197–304.
Swanson JA, Baer SC. 1995. Phagocytosis by zippers and triggers.
Trends Cell Biol 5:89 –93.
Swanson JA, Johnson MT, Beningo K, Post P, Mooseker M, Araki N.
1999. A contractile activity that closes phagosomes in macrophages. J Cell Sci 112:307–316.
Swanson JA, Watts, C. 1995. Macropinocytosis. Trends Cell Biol
5:424 – 428.
Tang N, Ostap, EM. 2001. Motor domain-dependent localization of
myo1b (myr-1). Curr Biol 11:1131–1135.
Temesvari LA, Bush JM, Peterson MD, Novak KD, Titus MA,
Cardelli JA. 1996. Examination of the endosomal and lysosomal pathways in Dictyostelium discoideum myosin I mutants.
J Cell Sci 109:663– 673.
Titus MA. 1999. A class VII unconventional myosin is required for
phagocytosis. Curr Biol 9:1297–1303.
Titus MA, Warrick HM, Spudich, JA. 1989. Multiple actin-based
motor genes in Dictyostelium. Cell Regul 1:55– 63.
Titus MA, Wessels D, Spudich JA, Soll DR. 1993. The unconventional
myosin encoded by the myoA gene plays a role in Dictyostelium motility. Mol. Biol. Cell 4:233–246.
Tuxworth RI, Weber I, Wessels D, Addicks GC, Soll DR, Gerisch G,
Titus MA. 2001. A role for myosin VII in dynamic cell adhesion. Curr Biol 11:218 –329.
Wessels D, Murray J, Jung G, Hammer JA, Soll DR. 1991. Myosin IB
null mutants of Dictyostelium exhibit abnormalities in motility.
Cell Motil Cytoskeleton 20:301–315.
Yonemura S, Pollard TD. 1992. The localization of myosin I and
myosin II in Acanthamoeba by fluorescence microscopy. J Cell
Sci 102:629 – 642.
Zhu Q, Clark M. 1992. Association of calmodulin and unconventional
myosin with the contractile vacuole complex of Dictyostelium.
J Cell Biol 118:347–358.
Zot HG, Doberstein SK, Pollard TD. 1992. Myosin-I moves actin
filaments on a phospholipid substrate: Implications for membrane targeting. J Cell Biol 116:367–376.