MICROBIOLOGY AND MOLECULAR BIOLOGY REVIEWS, Mar. 2002, p. 21–38
1092-2172/02/$04.00⫹0 DOI: 10.1128/MMBR.66.1.21–38.2002
Copyright © 2002, American Society for Microbiology. All Rights Reserved.
Vol. 66, No. 1
Cytoskeleton of Apicomplexan Parasites
Naomi S. Morrissette* and L. David Sibley
Department of Molecular Microbiology, Washington University School
of Medicine, St. Louis, Missouri 63110
asite invasion of new cells account for much of the tissue
damage associated with apicomplexan infections. Although
apicomplexans are haploid for the bulk of their life cycles, they
have complex life cycles, involving differentiation to forms that
invade distinct tissues and hosts (Fig. 1). Differentiation can
generate gametes that undergo fusion to generate a transient
diploid zygote. The zygote immediately undergoes meiosis to
reestablish haploid organisms. In some cases, differentiation
also permits infection of organisms (such as mosquitoes or
ticks) that serve as vectors to transmit parasites from host to
host (2, 26, 39, 60, 67, 90, 157).
Apicomplexan parasites share a variety of morphological
traits that are considered diagnostic for this phylum (Fig. 2).
These protists have an elongated shape and a conspicuous
specialization of the apical region (2, 3, 26, 157). Many of the
distinct characteristics constitute a collection of unique organelles termed the apical complex. These organelles include
the rhoptries, the micronemes, the apical polar ring, and the
conoid. Rhoptries and micronemes are unique secretory organelles that contain products required for motility, adhesion
to host cells, invasion of host cells, and establishment of the
parasitophorous vacuole (2, 22, 24, 26, 41, 139). The conoid is
a small cone-shaped structure composed of a spiral of unidentified filaments (120, 138). It is thought to play a mechanical
role in invasion of host cells and is present in only some
apicomplexans. The apical polar ring is a hallmark organelle of
all members of the Apicomplexa (120, 134). It serves as one of
the three microtubule-organizing centers (MTOCs) in these
parasites; spindle pole plaques and centrioles/basal bodies are
the other MTOCs (26, 157). In addition to the apical complex,
THE APICOMPLEXA
Infection by parasitic protozoa of the phylum Apicomplexa causes incalculable morbidity and mortality to humans
and agricultural animals (2, 3, 19, 26, 28, 39, 90). Apicomplexan parasites include Plasmodium spp., the agents of malaria; Toxoplasma gondii, a significant opportunistic pathogen
in immunocompromised individuals; Eimeria spp., pathogens
of chicken and cattle; Theileria spp., tick-borne parasites of
cattle in Africa; and Cryptosporidium, an animal parasite as
well as an opportunistic pathogen of humans. The phylum also
includes gregarines, parasites of the guts of invertebrates including cockroaches and shrimp. This review is restricted to
apicomplexan parasites of medical and agricultural importance
since the bulk of the research has been done in this area.
All apicomplexans are obligate intracellular parasites. Most
apicomplexan parasites grow and replicate within the parasitophorous vacuole, a nonphagosomal, membrane bound compartment that is segregated from most cellular trafficking pathways (78, 110, 143, 186). Proliferation of these organisms
occurs by invasion of a host cell and is followed by parasite
growth and cell division until the host cell is lysed by the
replicating parasites. Parasites released by host cell lysis do not
grow or undergo cell division extracellularly and must rapidly
reinvade other host cells in order to survive. Repeated cycles of
host cell invasion, parasite replication, host cell lysis, and par* Corresponding author. Mailing address: Department of Molecular
Microbiology, Washington University School of Medicine, 660 South
Euclid Ave., St. Louis, MO 63110. Phone: (314) 362-8874. Fax: (314)
362-3203. E-mail: naomi@borcim.wustl.edu.
21
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THE APICOMPLEXA..................................................................................................................................................21
MICROTUBULES IN THE APICOMPLEXA ..........................................................................................................23
Organization ..............................................................................................................................................................23
Conoid ........................................................................................................................................................................25
Role in Apicomplexan Replication .........................................................................................................................25
Tubulin and Microtubule-Associated Proteins .....................................................................................................28
SUBPELLICULAR NETWORK OF THE APICOMPLEXA ..................................................................................28
Proteins of the Subpellicular Network...................................................................................................................28
Organization of the Inner Membrane Complex and the Subpellicular Network ............................................28
ACTIN AND MYOSIN IN THE APICOMPLEXA ...................................................................................................29
Properties and Localization of Actin .....................................................................................................................29
Motility and Invasion ...............................................................................................................................................29
Actin and Actin Binding Proteins ..........................................................................................................................31
Myosin ........................................................................................................................................................................31
MANIPULATION OF THE HOST CYTOSKELETON BY APICOMPLEXAN PARASITES ...........................32
Reorganization of the Microvilli of Intestinal Epithelia by Cryptosporidium ...................................................32
Plasmodium Modification and Mimicry of Erythrocyte Cytoskeletal Proteins.................................................32
Theileria Exploitation of Host Cell Microtubules.................................................................................................33
CONCLUSIONS ...........................................................................................................................................................33
ACKNOWLEDGMENTS .............................................................................................................................................35
REFERENCES ..............................................................................................................................................................35
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MICROBIOL. MOL. BIOL. REV.
the apicomplexans have other unique structural features, such
as an essential chloroplast-like organelle called the apicoplast
(87, 99, 169, 194). The parasites are bounded by the pellicle, a
composite structure consisting of the plasma membrane and
the closely apposed inner membrane complex (IMC) (2, 4, 26,
43, 44, 58, 103, 104, 177, 183). The pellicle is intimately associated with a number of cytoskeletal elements, including actin,
myosin, microtubules, and a network of intermediate filamentlike proteins (Fig. 3).
Apicomplexan protozoa share a number of cytoskeletal elements (microtubules, actin, myosin, and intermediate filament-like proteins) with other “typical” eukaryotic systems
used to study the cytoskeleton. Nonetheless, studies of the
cytoskeletons of apicomplexans have revealed startling differ-
ences from model organisms that are worth mentioning at the
outset. For example, the singlet subpellicular microtubules of
apicomplexans are unusually stable and withstand the high
pressure, cold, and detergents typically used to isolate them,
conditions incompatible with survival for most microtubules
(113). In contrast, the microfilaments of apicomplexans are
thought to be exceedingly transient. Microfilaments are observed only after treatment with jasplakinolide (a drug that
drives actin polymerization), and the bulk of actin (⬎98%) is
sequestered in globular form (12, 36, 151). Apicomplexan myosins are unconventional, constituting a new class of unusually
small “neckless” motors (68–70, 73, 123). In summary, apicomplexan protozoa constitute an ancient and diverse phylum with
peculiar cell biological traits that make these parasites an in-
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FIG. 1. Life cycle of apicomplexan parasites. For the faint of heart or those not interested in details, the center circle illustrates a generic
apicomplexan life cycle. The rapidly proliferating haploid form has the capacity to differentiate into gametes that fuse to produce a diploid
zygote. Meiosis reestablishes the haploid state and leads to sporozoites, the form most widely associated with establishment of new infections
after inoculation of a naive host by an infected vector. The outer two circles represent the specific life cycles for T. gondii and P. falciparum.
The bradyzoite form of Toxoplasma (ⴱ) is responsible for reactivation of latent infection and is an obligatory stage between tachyzoites and
gametes. Plasmodium spp. do not have an analogous stage, although “hypnozoites” (latent sporozoites) are implicated in reactivation by P.
vivax and P. ovale.
VOL. 66, 2002
CYTOSKELETON OF APICOMPLEXAN PARASITES
triguing topic for study. This review will survey our current
understanding of the cytoskeleton of apicomplexan parasites.
MICROTUBULES IN THE APICOMPLEXA
Organization
The haploid forms of apicomplexan parasites have two discrete populations of microtubules: the subpellicular microtubules and the spindle microtubules (Fig. 3A and a). The subpellicular microtubules radiate from the apical polar ring and
run down the cytosolic face of the pellicle, ending in the region
below the nucleus (approximately two-thirds of the length of
the parasite [1, 12, 13, 26, 31, 120, 131]). These spirally arranged microtubules closely follow the serpentine body shape
of apicomplexans. Subpellicular microtubules confer both
elongated shape and apical polarity to apicomplexan parasites.
Replicative forms or drug-treated parasites that lack subpellicular microtubules are nonpolar, nonmotile, and noninvasive
(72, 168). Subpellicular microtubules are organized by lateral
association with the apical polar ring (APR), a circular MTOC
(120, 134). Attachment of subpellicular microtubules to the
APR is supported by blunt projections of the APR, which form
a cogwheel pattern in transverse views. The plus end of the
subpellicular microtubules is distal to this MTOC, and the ends
of the subpellicular microtubules do not appear to be physically capped (134).
The arrangement of subpellicular microtubules varies
among apicomplexan species, but the number, length, and organization are absolutely stereotyped within the life cycle stage
of a species. The number of subpellicular microtubules ranges
from a band of 3 or 4 in the tiny Plasmodium falciparum
merozoites to ⬃60 in the considerably larger Plasmodium ookinetes (2, 3, 12, 26, 136, 156, 157). In coccidian parasites
(Toxoplasma and Eimeria), the subpellicular microtubules are
evenly spaced beneath the periphery of the pellicle; however,
in Plasmodium species, most of the microtubules occupy twothirds of the circumference and one microtubule is centered
within the latter one-third of the pellicle (3, 157, 180). P.
falciparum merozoites are an exception to this spacing generalization; they have a reduced number (three or four) of subpellicular microtubules termed f-MAST (falciparum merozoite-associated assemblage of subellicular microtubules) that
extend down one side of the merozoite membrane from the
apex toward the posterior (12, 59). Other P. falciparum life
cycle stages (sporozoites and ookinetes) have a full complement of subpellicular microtubules, as do merozoites of other
Plasmodium species. Theileria sporozoites also deviate from
this pattern. Although the tick-borne kinete stage of Theileria
contains subpellicular microtubules and an IMC, Theileria
sporozoites lack subpellicular microtubules and the IMC altogether and enter host cells in a distinct fashion (see below) (49,
50, 147, 148, 152).
Replicating parasites employ spindle microtubules during
mitosis. Nuclear division proceeds without nuclear membrane
breakdown (12, 26, 59, 79, 130, 161). Spindle microtubules are
nucleated from electron-dense, amorphous plaques associated
with nuclear invaginations and embedded in the nuclear membrane (12, 26, 82, 109, 140, 146, 157). This spindle-organizing
structure has been variously referred to as the centrocone, the
centriolar plaque, the spindle pole body, or the centriolar
equivalent. We have chosen to characterize this structure as a
“spindle pole plaque” to distinguish it from the adjacent centrioles that are sometimes present in members of the Apicomplexa. Centrioles are apparently not required for spindle assembly, since Plasmodium merozoites and Theileria sporozoites
lack these structures and construct spindles by using only the
spindle pole plaques (5, 12, 51, 140, 150, 157, 159). However, it
is also possible that centrioles exist in these organisms and are
obscured by inclusion in the electron-dense spindle pole
plaque.
In many apicomplexans, centrioles are located in the cytoplasm close to but separate from the spindle pole plaques
(26, 31, 42, 82, 100, 116, 157). Centrioles are highly ordered
MTOCs typically consisting of a 9⫹0 structure of nine triplet
microtubule blades organized in a turbine fashion. Apicomplexan centrioles have an unconventional form consisting of a
central single microtubule surrounded by nine singlet microtubules, a deviation from the canonical structure (31, 40, 42,
82, 157, 165, 182). It is curious that apicomplexans apparently
contain both spindle pole plaque structures and centrioles in
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FIG. 2. The morphology of apicomplexan parasites. Apicomplexans are highly polarized cells containing a collection of organelles that
are specific to the phylum. Rhoptries, micronemes, and dense granules
are secretory organelles that contain products required for motility,
invasion, and establishment of the parasitophorous vacuole. The
conoid is a small cone-shaped spiral of unidentified filaments. It is
thought to play a mechanical role in invasion and can be protruded
from or retracted into the apical polar ring. The apical polar ring
serves as an MTOC for the subpellicular microtubules. The spirally
arranged subpellicular microtubules closely follow the serpentine body
shape of apicomplexans. The parasites are bounded by the pellicle, a
composite structure consisting of the plasma membrane and the
closely apposed IMC. The endoplasmic reticulum surrounds the nucleus, and the Golgi body is immediately above it. The apicoplast is
immediately adjacent to the Golgi body.
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CYTOSKELETON OF APICOMPLEXAN PARASITES
Conoid
Some apicomplexan parasites (Toxoplasma, Eimeria, and
Sarcocystis) contain an additional cytoskeletal structure, the
conoid, not found in other apicomplexans (Plasmodium and
Theileria). It has been suggested that the conoid plays a
mechanical role in invasion by parasites that must penetrate
the robust barrier of the intestinal epithelium of vertebrates
(26, 108, 121, 136, 138). The conoid consists of a set of counterclockwise spiraling filaments that create a pointed or coneshaped structure at the extreme apex of these parasites (26, 64,
136, 138, 153). Extrusion of the conoid can be stimulated by
ionomycin-triggered calcium influx, whereas pretreatment with
cytochalasin D inhibits conoid extrusion (108). The filamen-
tous subunits of the conoid resemble microtubules but are
curled into an extremely tight coil, suggesting that if they are
microtubules they are unusually constructed or deformed. (Recent work by K. Hu, D. Roos, and J. Murray [submitted for
publication] suggests that the Toxoplasma conoid is constructed from tubulin organized into a novel polymer form
consisting of a comma-shaped sheet of nine protofilaments.)
The conoid is approximately 250 nm in diameter and can be
extended beyond or retracted into the apical polar ring. There
are two ⬃400-nm-long, closely associated microtubules in the
middle of the conoid. These microtubules are tightly bound to
each other and are eccentric to the longitudinal axis of the
conoid. This arrangement may be due to contact with the
conoid and preconoidal rings via lateral projections (120). The
course of the conoid subunits parallels the counterclockwise
spiral of the subpellicular microtubules.
Role in apicomplexan replication
Apicomplexan parasites replicate by internal budding to
create either two daughter cells or multiple progeny. Nuclear divisions within the Apicomplexa are cryptomitoses
(the nuclear membrane remains intact throughout), and
kariokinesis occurs without chromosomal condensation (26,
79, 130, 157). Replication in Toxoplasma and Neospora occurs
by endodyogeny, which creates two daughter parasites (63, 71,
91, 154). Replication by Plasmodium, Theileria, Eimeria, and
Babesia occurs by schizogeny, which can create 64 daughter
parasites (3, 12, 72, 79, 150, 157). The processes of endodyogeny and schizogeny are quite similar, differing mainly in the
preservation or loss of maternal cell specialization. In
endodyogeny, two daughter parasites are formed within an
intact, fully polarized mother parasite (Fig. 4A). This preserves
the ability of replicating tachyzoites to invade throughout the
cell cycle. The internal daughter cells are delimited by an IMC
and associated subpellicular microtubules, and each contains
(in addition to the nucleus, mitochondrion, Golgi, and plastid)
a complete set of apical organelles (100, 154, 183). When the
daughter cells are fully mature, the maternal apical complex is
disassembled and the daughter parasites bud from the mother,
adopting her plasma membrane. In schizogeny, after host cell
invasion, the parasite subpellicular microtubules and apical
complex are disassembled (Fig. 4B). After growth and multiple
nuclear divisions, polarized parasites are regenerated when the
nuclei move to the schizont periphery and associate with the
assembling IMC, subpellicular microtubules, and apical organelles (3, 12, 72, 157). The newly repolarized parasites then
bud out of the maternal cell as merozoites. Due to their large
FIG. 3. Cytoskeletal elements in the Apicomplexa. (A) The subpellicular microtubules radiate out of the apical polar ring and run down the
cytosolic face of the IMC. The spindle microtubules are nucleated from spindle pole plaques within the nuclear membrane. Centrioles (consisting
of nine singlet microtubules surrounding a central singlet microtubule) are adjacent to the spindle pole plaques. (a) Isolated, negatively strained
conoid (co), apical polar ring (apr), and subpellicular microtubules from T. gondii (113). (B) A network of intermediate filaments, the subpellicular
network, underlies the length of the IMC. The lower right inset illustrates the pattern of IMPs revealed by freeze fracture of the IMC. These
particles may represent the transmembrane domains of receptors that link the subpellicular network to the cytoplasmic face of the inner membrane
complex. (b) Glycerol- and detergent-extracted, freeze-dried replicas of Toxoplasma tachyzoites illustrate the regular array of the subpellicular
network filaments. (Image of the Toxoplasma network kindly provided by J. Heuser.) (C) Actin is localized between the plasma membrane and
the IMC. When the plasma membrane is separated from the IMC, actin remains associated with the IMC and myosin is associated with the plasma
membrane. (c) Actin filaments protrude beyond the conoid (co) of Toxoplasma tachyzoites treated with jasplakinolide, a drug that induces actin
polymerization. (Panel c reprinted from reference (151) with the permission of the publisher.)
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addition to the apical polar ring to organize microtubules. It
may be that the centrioles are maintained throughout the asexual life cycle in order to serve as a template for construction of
basal bodies that nucleate flagellar axonemes in the male gametes. The centriole is also associated with inheritance of the
apicoplast during replication in Toxoplasma (169). One intriguing possibility is that the centrioles function as a “super-organizing center” coordinating the apical polar ring MTOC and
the spindle pole plaque MTOC.
Although most parasite replication occurs by asexual division, apicomplexans also differentiate to gametes that fuse to
form a diploid zygote. The male gamete (microgamete) is
flagellated and swims to the female gamete (macrogamete) to
carry out fertilization. The anterior end of apicomplexan microgametes is pointed and contains three basal bodies in close
proximity to the apical pole (116, 137, 158, 181). These basal
bodies nucleate two or three flagella that extend past the nucleus and away from the apical end. Additional microtubules
originate in the basal apparatus zone and extend to the posterior end of the microgamete. Two of the flagella are long and
are free from association with the gamete body. The third
flagellum is shorter and is attached to surface of gamete at its
anterior end. In some species this third flagellum is present
only in rudimentary form as a band of microtubules that extends along the length of the gamete. In contrast to the atypical
centrioles observed in other stages, the basal body of male
gametes has a typical triplet microtubule structure with ninefold symmetry and the flagellar axoneme contains a conventional 9⫹2 arrangement of doublet microtubules surrounding
the central pair of microtubules (137, 158–160, 181). In Plasmodium, genesis of the basal bodies involves an intermediate
9⫹1 singlet form similar to the centrioles in other apicomplexans (157).
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MICROBIOL. MOL. BIOL. REV.
round shape and lack of apical specialization, replicating parasites are noninvasive during schizogeny.
P. falciparum replication has been studied by observing microtubule structures during replication in red cells (130). Premitotic intracellular parasites have a diffuse distribution of
unpolymerized tubulin. The first visible structure is a spindle
pole plaque that is localized within the nuclear membrane.
This nucleates microtubules to give rise to a hemispindle,
which partitions into two. Partitioning of the spindle pole
plaque has also been observed by electron microscopy. A band
of striated fibers termed the couche fibrillaire spans the separating plaques (42, 140, 161). This structure may represent
centrin fibers, since similar structures are observed in algae
and yeast (89, 164, 192). The spindle halves segregate in opposite directions by migration within the nuclear membrane to
ultimately produce a full spindle. The multiple nuclear divisions of schizogeny are not completely synchronous; spindles in
various stages of assembly are often present in a single latestage schizont. Distinct postmitotic microtubule structures appear in late schizogeny. Each structure is associated with an
individual nucleus and consists of extranuclear microtubules
arranged in a regular radial array like the spokes on a cartwheel. These microtubules extend beyond the nucleus toward
the center of the schizont. Remains of this microtubule organization persist as a rod-like structure in budding merozoites.
This remnant may represent f-MAST, the reduced set of three
or four subpellicular microtubules that occurs in P. falciparum
merozoites (12, 59).
The microtubules of extracellular apicomplexans are not
dynamic and are therefore impervious to microtubule-disrupting drugs (135). Assembly of microtubules occurs in the course
of replication; therefore, intracellular parasites are susceptible
to microtubule-depolymerizing drugs (Table 1) (168). In Toxoplasma and Plasmodium, the spindle and subpellicular microtubule populations are differentially stable to disruption by
oryzalin or colchicine (14, 115). Lower concentrations (0.5 M
oryzalin or 1.0 mM colchicine) shorten microtubules. Under
these conditions, Toxoplasma and Plasmodium continue to undergo nuclear division and budding but lack functional subpellicular microtubules and are incapable of invading new host
cells. When removed from 0.5 M oryzalin, Toxoplasma recovers normal morphology and is invasive (115). In contrast,
higher concentrations of drug (2.5 M oryzalin or 5.0 to 10.0
mM colchicine) disrupt both subpellicular and spindle microtubules (149, 168). Parasites under these conditions are incapable of nuclear division or budding, although cell growth,
DNA synthesis, and centriole replication continue unchecked
(115, 149, 168). When removed from 2.5 M oryzalin, Toxoplasma tachyzoites attempt to bud as crescent-shaped parasites. Since the polyploid nuclear mass cannot be correctly
segregated, daughter parasites are made that lack nuclei altogether (115).
Assessment of the activity of microtubule-destabilizing drugs
on intracellular parasites is complicated by the activity of these
drugs on host cells. In many cases, toxicity to host cells antedates any effects on parasite microtubules. In this situation,
parasites fail to replicate, but this is due to adverse effects on
the host cell rather than to direct inhibition of parasite func-
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FIG. 4. Replication by the Apicomplexa. (A) Endodyogeny proceeds without loss of maternal cell shape and apical polarity. Formation of a
curved nucleus is coupled to construction of two buds composed of nascent daughter IMC and subpellicular microtubules. The daughter cells
develop surrounded by their own subpellicular microtubules and IMC in the fully polarized mother cell. Once the daughter cells are mature, they
bud out of the remnants of the mother cell. (B) Schizogeny proceeds with loss of apical polarity. Invasion of a polarized elongated parasite is
followed by disassembly of the subpellicular microtubules and IMC and by extensive cell growth and nuclear divisions. To reassemble polarized
daughter cells, the multiple nuclei align with individual sets of apical organelles, subpellicular microtubules, and IMC at the periphery of the
replicating cell.
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27
TABLE 1. Drug disruption of cytoskeletal elements in the Apicomplexa
Drug and cytoskeletal element
(drug mechanism)
Actin microfilaments
Cytochalasin D (disrupter)a
Latrunculin (disrupter)b
Jasplakinolide (stabilizer)c
Concn
0.1–0.2 M
10 nM–5 M
50 nM–2 M
Action
Comments
Inhibits actin-based gliding motility
(extracellular) and invasion
Inhibits gliding motility in extracellular
parasites; use resistant host cells to avoid
host cell effects.
Will also show host cell actin effects.
Acts on extracellular parasites; will also act
on host cell actin in intracellular settings.
Inhibits motility and invasion
Filament polymerization inhibits
gliding and invasion
Microtubules
Oryzalin (disrupter)d
0.5 M
Parasites lack apical polarity
Spindle microtubules
2.5 M
Parasites are incapable of
replication
Colchicine (disrupter)e
Subpellicular microtubules
Spindle microtubules
10 M–1 mM
5–10 mM
Parasites lack apical polarity
Colcemid (disrupter)f
50 M
Parasites are incapable of
replication
Inhibits intracellular growth
Vinblastine (disrupter)g
15–100 nM
Inhibits intracellular growth
Vincristine (disrupter)h
7 nM
Tubulozole (disrupter)i
10 M
Parasites are killed
Taxol (stabilizer)j
0.1–0.5 M
Parasites are incapable of budding
Docetaxel/taxotere (stabilizer)k
10 nm–50 M
Inhibits intracellular growth
Parasites are incapable of
replication
Epithalone A (stabilizer)l
0.1–0.5 M
Parasites are incapable of budding
Myosin
BDM (ATPase inhibitor)m
20–40 mM
Inhibits actin-based gliding motility
(extracellular) and invasion
Inhibits actin-based gliding motility
(extracellular) and invasion
KT5926 (LC kinase inhibitor)n
1–5 M
No drug effect on extracellular parasites;
use resistant host cells to avoid host cell
disruption (no microtubules in RBCs).
As for subpellicular microtubules.
No cell biological observations accompany
study.
No cell biological observations accompany
study.
No cell biological observations accompany
study.
May inhibit parasite protein synthesis
rather than microtubules
No drug effect on extracellular parasites;
host cell microtubules will also be
affected (no microtubules in RBCs).
No drug effect on extracellular parasites;
host cell microtubules will also be
affected (no microtubules in RBCs).
No drug effect on extracellular parasites;
host cell microtubules will also be
affected (no microtubules in RBCs).
Will also act on host cell myosins (not
present in red cells).
Most probably inhibits secretion of TRAP
family adhesins rather than myosin.
a
Data from references 8, 25, 28, 35, 37, 39, 54, 56, 57, 64, 69, 85, 103, 104, 129, 131, 140, 143–145, 148, 152, 156, 163, and 170.
Data from reference 145.
Data from references 123, 145, and 147.
d
Data from references 11, 17, 22, 59, 78, 96, 110, 145, and 164.
e
Data from references 15, 23, 59, 85, 131, 156, 173, 185, 188, and 189.
f
Data from reference 16.
g
Data from references 16, 35, 120, 131, 156, 174, 188, and 189.
h
Data from references 35 and 174.
i
Data from references 16, 35, and 36.
j
Data from references 59, 69, 85, 144, 145, 159, and 168.
k
Data from references 137 and 159.
l
Data from reference 168.
m
Data from references 37, 56, 57, 64, and 119.
n
Data from reference 37.
b
c
tions. For these reasons, the most comprehensive examination
of the effects of microtubule-disrupting drugs has been carried
out with the P. falciparum erythrocytic stages in red cells
(which lack microtubules). In vitro studies have established
that replicating P. falciparum merozoites are susceptible to
colchicine and colcemid, to vinblastine and vincristine, to the
dinitroanilines trifluralin and pendimethalin, and to tubulozoles, including tubulozole-T, an isomer that is inactive in
mammalian systems (15, 33, 34, 178). In contrast, merozoites
show only low sensitivity to the benzimidazoles (albendazole,
thiabendazole, mebendazole, and omeprazole) (33, 163). The
results for benzimidazole are consistent with the deduced
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Subpellicular microtubules
Also other dinitroanilines such as
trifluralin, ethafluralin
No effect on extracellular parasites; no
effect on host cell microtubules, but
intracellular parasites are affected.
As for subpellicular microtubules.
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Tubulin and Microtubule-Associated Proteins
Apicomplexan tubulin genes have been cloned from T. gondii, Eimeria tenella, Babesia bovis, P. yoelii, P. berghei, C. parvum, and P. falciparum (8, 20, 21, 25, 29, 30, 74, 75, 118, 119,
145, 179, 197). In most of these apicomplexans, the ␣-tubulin
and -tubulin genes appear to be unlinked, single-copy genes
containing up to three introns. The introns are in the same
location and are similar in sequence in E. tenella, C. parvum,
and T. gondii. The ␥-tubulin gene has also been sequenced in
P. falciparum. It is a single-copy gene and lacks introns (95).
Curiously, P. falciparum and P. yoelii each have two ␣-tubulin
genes, which are located on different chromosomes (8, 129).
The ␣-tubulin-I gene is expressed throughout the parasite differentiation cycle, but the ␣-tubulin-II gene is specifically expressed in male gametes. The ␣-tubulin-II-specific monoclonal
antibody 5E7 specifically labels stage III through mature male
gametocytes and exflagellating and free male gametes (129).
Immunoelectron microscopy using this antibody labels the axonemes of male gametes.
Microtubule-associated proteins (MAPs) are clearly critical to the highly organized structure of apicomplexan parasites. Bridges connecting the subpellicular microtubules to
the inner membrane complex have been observed in thin
sections of parasites (3, 131, 195). Isolated frozen-hydrated
microtubules of T. gondii have a distinct 32-nm periodicity
along their length as revealed by Fourier analysis (113). The
periodicity most probably results from a MAP that heavily
decorates these microtubules and that may account for their
unusual stability after isolation. This MAP may coordinate the
close interaction of the subpellicular microtubules with the
IMC. The existence of a group of monoclonal antibodies that
labels the subpellicular microtubules in Toxoplasma and crossreacts with Plasmodium suggests that the MAPs may be conserved within the Apicomplexa (112, 114). The Plasmodium
and Cryptosporidium genome databases and the Toxoplasma,
Eimeria, and Neospora expressed sequence tag (EST) projects
have sequences annotated as encoding putative kinesins and
dyneins.
SUBPELLICULAR NETWORK OF THE APICOMPLEXA
Proteins of the Subpellicular Network
Deoxycholate extraction of Toxoplasma reveals a network of
filamentous material that extends from the APR to the posterior of the parasite (98). The filaments have a diameter of 8 to
10 nm, and the network has the same general shape as the
parasite, suggesting that these filaments may play a role in
generating and maintaining cell shape. A polyclonal serum
generated from the extracted pellicles has been used to identify
two novel proteins that localize to the subpellicular network
(98). These proteins, TgIMC-1 and TgIMC-2, are not homologous to any known proteins but are predicted to form regions
of coiled coils and have weak similarity to the coiled-coil domains of cytoskeletal proteins such as myosin. TgIMC-1 is rich
in valine and glutamic acid (together they constitute almost
30% of the protein). Interestingly, TgIMC-1 is similar to the
articulins, a group of proteins that form a membrane skeleton
in Euglena and other protists. Both the articulins and TgIMC-1
have a 12-amino-acid VPV repeat. Gold labeling with antisera
to either TgIMC-1 or TgIMC-2 labels the cytoplasmic face of
the pellicle between the subpellicular microtubules and the
IMC (Fig. 3B). A homolog of TgIMC-1 is present in the P.
falciparum genome. PfIMC-1 has an additional 220-amino-acid
region near the C terminus, containing seven copies of a repeating sequence. The TgIMC-1 antiserum cross-reacts with
Plasmodium, labeling late-stage schizonts during the formation
of merozoites.
Organization of the Inner Membrane Complex
and the Subpellicular Network
The IMC lies directly below the parasite plasma membrane
and is closely associated with it, creating a three-layered pellicle characteristic of the Apicomplexa (26, 104, 183). In Plasmodium sporozoites, the IMC is constructed from a single
large flattened vesicle joined by a single suture line traversing
the long axis of the parasite (44). In other apicomplexans, it is
made of many flattened vesicles aligned in longitudinal rows
and joined in a patchwork fashion by sutures (43, 124). Glycerol-extracted, negatively stained pellicles from Sarcocystis
ovifelis, Besnotia jellisoni, and Eimeria falciformis all have a
mesh-like pattern that runs along the full length of the IMC
(32). This mesh-like structure is similar to the network of
IMC-associated filaments observed in Toxoplasma. Freeze
fracture of Toxoplasma, Sarcocystis, Eimeria, and Plasmodium
has shown that apicomplexans share a striking organization of
the IMC (6, 43, 44, 104, 124). The membranes of the IMC are
characterized by parallel alignment of intramembranous particles (IMPs). The lines of IMPs extend down the long axis of
the parasite and are organized as single rows interspersed with
double rows. The double rows correspond in number and arrangement to the underlying subpellicular microtubules (Fig.
3B, inset). The rows show continuity across the plates of the
IMC, which is remarkable because each plate is a topologically
distinct vesicle. Fourier analysis of the IMPs shows that they
have a distinct 32-nm longitudinal repeat creating a two-dimensional lattice, with the second dimension at an angle of
approximately 75° to the rows (113). The integrity of the particle lattice is not destroyed by disruption of actin or microtu-
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amino acid sequences of apicomplexan -tubulins that lack
Glu 198 and Phe 200, predictors of sensitivity to these compounds (44a, 81a). Plasmodium merozoites are also sensitive to
microtubule-stabilizing drugs such as taxol, docetaxel, and
epothilone A, which inhibit schizont nuclear division and budding (141, 162, 172). Colchicine, trifluralin, and taxol also perturb gametocyte and ookinete differentiation in Plasmodium
(80, 88). Parallel experiments to examine the role of microtubules in other apicomplexans have been carried out with Eimeria, Toxoplasma, and Cryptosporidium by using dinitroaniline compounds such as oryzalin or trifluralin (10, 16, 168).
These drugs specifically inhibit the microtubules of plants and
protists and do not destabilize vertebrate microtubules. To
assess the effects of other drugs such as colchicine and taxol,
resistant host cells can be used. When colchicine- or taxolresistant cells are used as host cells for Toxoplasma replication,
the effects of colchicine and taxol are akin to what is observed
in Plasmodium merozoites (115).
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CYTOSKELETON OF APICOMPLEXAN PARASITES
29
bules; this suggests the existence of additional cytoskeletal
filaments. Glycerol- and detergent-extracted freeze-dried replicas of Toxoplasma tachyzoites reveal a filament network with
striking similarity to the IMP lattice (Fig. 3b). The recent
discovery of intermediate filament-like proteins that localize to
the subpellicular network most probably identifies the latticegenerating network elements. We hypothesize that the IMP
lattice may represent the transmembrane domains of receptors
that anchor the subpellicular network to the IMC. This lattice
may also anchor the subpellicular microtubules since they are
decorated with a MAP that binds at 32-nm intervals (see
above) (113).
parasite and can be up to 2 M long (151). Jasplakinolideinduced filaments decorate with myosin subfragment 1, demonstrating that they are indeed actin; unfortunately, the filaments are too close together to determine polarity. Anti-actin
antibodies label the apical projections in jasplakinolide treated
parasites. In untreated parasites, heterologous actin antibodies
label the apical region or appear as a diffuse distribution in the
cytosol (47, 187, 196). A polyclonal antiserum directed against
Toxoplasma actin labels a circumferential pattern in the
tachyzoites, extending below the apical region (36). In hypotonically swelled parasites, a Toxoplasma-specific antiactin
monoclonal antibody labels the region between the plasma
membrane and the IMC (35).
ACTIN AND MYOSIN IN THE APICOMPLEXA
Properties and Localization of Actin
Motility and Invasion
A large fraction of the actin in parasites of the Apicomplexa
appears to be in monomeric rather than polymerized form.
Experiments with Toxoplasma have established that tachyzoites have strikingly small amounts of assembled actin (36).
Approximately 98% of Toxoplasma actin is globular, and
this distribution is not shifted by the addition of agents that
drive actin polymerization, such as phalloidin, MgCl2, exogenous actin, spermine, or phosphatidylinositol-4,5-bisphosphate
(PIP2). In fact, although some researchers have observed phalloidin binding (127, 187) or filament stabilization with phalloidin (126), others have concluded that phalloidin does not bind
to apicomplexan microfilaments (27, 36, 55, 56, 111, 151). Until
quite recently, apicomplexan microfilaments had not even
been observed by electron microscopy (12). However, in the
presence of the microfilament-polymerizing drug jasplakinolide, Toxoplasma will form microfilaments (Fig. 3C and c) (126,
151). Jasplakinolide induces actin polymerization, most notably at the apical end of extracellular Toxoplasma tachyzoites
(Table 1). The actin filaments in these membrane-enclosed
apical projections are aligned parallel to the long axis of the
Most members of the Apicomplexa are motile. In these
species, locomotion is intimately associated with host cell invasion and probably employs the same underlying cellular machinery. In fact, gliding locomotion and invasion often appear
continuous and occur without a discernible change in speed.
Apicomplexans (including gregarines) move by gliding motility
(38, 52, 77, 84, 133). This movement does not exploit a discrete
organelle (such as a flagellum) or result from amoeboid deformations of the cell. It is, however, substrate dependent.
Motility has been analyzed in T. gondii and consists of three
behaviors: circular gliding, upright twirling, and helical gliding
(61, 66). When a crescent-shaped parasite lies on its right side,
it moves in counterclockwise circles. Twirling occurs when the
parasite is attached to the substrate by its posterior end, producing a clockwise spinning. Lastly, helical gliding is similar to
twirling but occurs in horizontal parasites (Fig. 5A). This biphasic behavior consists of an 180° clockwise revolution (resulting in a corkscrewing forward movement) coupled to a
parasite flip to return the parasite to its original face, so that it
can initiate helical motion anew. Helical gliding is the only
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FIG. 5. Motility and invasion by the Apicomplexa. (A) Helical gliding is the only behavior associated with motility that leads to long-distance
movement across a substrate. During this biphasic behavior, the parasite moves forward a body length and then repositions to repeat this cycle.
This sequence illustrates the path taken by a gliding Toxoplasma tachyzoite to move forward one complete cycle. The parasite travels forward by
moving a contact zone from its apex to its posterior in a helical path. Because the parasite is crescent shaped, it moves in a “corkscrew” fashion
(frames 1 to 5). The crescent shape of the tachyzoite prevents the continuation of this motion because the parasite cannot contact the substrate
with its concave face. To rectify this, the parasite rotates 180° without forward motion (frames 6 and 7) before initiating another cycle of
‘corkscrewing’ translational movement (frames 8 and 9). (Modified from reference (66) with the permission of the publisher.) (B) Apicomplexan
invasion consists of attachment and apical orientation, induction of a parasitophorous vacuole, and translocation of the parasite into the vacuole.
Apically secreted adhesins are capped along the moving junction to the posterior of the parasite. The moving junction is associated with a
constriction of parasite shape that moves from the apex to the posterior.
30
MORRISSETTE AND SIBLEY
cytic uptake of parasites because invasion of host cells (including macrophages) does not induce host cell membrane ruffling,
actin microfilament reorganization, or tyrosine phosphorylation, which are all indicative of phagocytosis (111). Invasion is
three to four times faster than phagocytosis (occurring within
25 to 40 s) and is characterized by parasite penetration into a
tight-fitting vacuole formed by invagination of the plasma
membrane. In contrast, phagocytosis of Toxoplasma involves
membrane ruffling and the parasite is captured in a loosefitting phagosome that forms over 2 to 4 min (78, 111, 121).
Phagocytosis involves both reorganization of the host cytoskeleton and tyrosine phosphorylation of host proteins (111).
Experiments with cytochalasin D-resistant T. gondii have
definitively established that invasion of Toxoplasma is critically
dependent on actin filaments in the parasite but not in the host
cell (37). Invasion of cytochalasin D-resistant host cells by
wild-type (cytochalasin-sensitive) Toxoplasma tachyzoites is
blocked by cytochalasin D. As observed with Plasmodium, attachment and apical orientation of Toxoplasma is normal in the
presence of cytochalasin D. Cytochalasin D-resistant Toxoplasma mutants were isolated by chemical mutagenesis and
selection for growth in cytochalasin D in resistant host cells.
These resistant parasites have a point mutation in the singlecopy actin gene ACT1 (A136G) and can invade wild-type host
cells in the presence of cytochalasin D. Transformation of the
mutant act1 allele into wild-type Toxoplasma confers cytochalasin D-resistant motility and invasion either as an allelic replacement or as a nonhomologous integration, generating a
pseudodiploid parasite (37).
Apicomplexan invasion consists of three phases: (i) attachment with apical orientation, (ii) induction of a parasitophorous vacuole, and (iii) translocation of the parasite into the
vacuole (Fig. 5B). Parasites attach to host cells and form an
intimate connection through apical end contact (37, 53, 106).
This results in sequential secretion from the parasite micronemes and rhoptries. Adhesins from the micronemes are
translocated along the parasite length and are shed at the site
of the moving junction; parasitophorous vacuole components
from the rhoptries are secreted into this forming compartment
(24). Both tight adhesion to the host cell and secretion into the
host cell occur when parasites are immobilized early in invasion by cytochalasin treatment (37, 53, 65, 106).
The “moving junction” which forms during invasion is a
circumferential zone of attachment at the orifice of the host
cell invagination (7, 105). It is characterized by a markedly
thickened host cell membrane with increased electron density
and is frequently accompanied by a constriction in the parasite
body. The parasite enters the nascent parasitophorous vacuole
by capping the moving junction down its body. Ultimately, the
parasite becomes enclosed within a cavity delimited by the
invaginated host cell membrane. Formation of a moving junction that is capped to the posterior during invasion is likely to
be a feature of invasion shared by many apicomplexans (Theileria is an exception [discussed below]). The moving junction is
a highly specialized interface of the parasite with the host cell,
presumably exploiting cytoskeletal proteins, signaling molecules, and receptors. Very little is known about this interface.
P. falciparum MCP-1 (merozoite-capping protein 1) is a 60kDa merozoite protein that moves from anterior to posterior
with the moving junction during merozoite invasion of red cells
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behavior that results in long-distance movement across a substrate. Actin-disrupting or -stabilizing drugs (cytochalasin D
and jasplakinolide), as well as myosin inhibitors (butanedione
monoxime [BDM]), disrupt Toxoplasma motility and invasion
(35, 37, 126). This suggests that an actomyosin-based mechanism underlies these behaviors (Table 1). Cytochalasin inhibition of gliding motility and/or invasion has been demonstrated
in C. parvum, T. gondii, E. tenella, E. acervulina, and Plasmodium species (37, 56, 77, 106, 133, 135). Motility and invasion
are associated with translocation of secreted adhesive proteins
from the apex to the parasite posterior and their shedding or
deposition into a slime trail. Both adhesins and parasite surface antigens are deposited into a trail by gliding parasites (11,
22, 37, 48, 166, 167, 171). The presence of surface proteins in
slime trails is likely to reflect the artificially sticky substrate
used to capture adhesin trails.
Conserved adhesins have been found in Plasmodium, Toxoplasma, Eimeria, Neospora, and Cryptosporidium spp. (22, 24,
92, 132, 155, 171, 185). The thrombospondin-related anonymous protein (TRAP) family adhesins have common structural
motifs and presumably an underlying mechanism of action.
They contain a conserved adhesive domain consisting of a
thrombospondin type 1 repeat that occurs in different numbers
and locations within the individual apicomplexan adhesins.
TRAP family proteins are localized to micronemes and are
apically secreted during motility and invasion (22, 171). TRAP
proteins are located on the surface of motile parasites in a
transmembrane form and are capped from anterior to posterior in gliding parasites. The short cytoplasmic tail of these
proteins is conserved and is thought to interact with the actomyosin cytoskeleton in order to translocate the protein backward. Adhesins are released as soluble protein by proteolytic
cleavage at the posterior end of locomoting parasites (23).
When the sporozoite-specific TRAP gene is knocked out in P.
berghei merozoites, parasites behave normally until they differentiate to sporozoites within the mosquito. TRAP knockout
sporozoites are immotile and noninvasive (171). Moreover,
expression of a TRAP construct missing the last 14 amino acids
of the cytoplasmic tail permits surface localization of the protein in knockout sporozoites (81). However, these sporozoites
also display atypical behavior, repeatedly gliding one-third of a
circle and snapping back to their original position. This suggests that the inability of TRAP to be appropriately translocated and/or released at the posterior end of locomoting parasites prevents their continued forward movement (81, 101).
Menard has recently written a comprehensive review of gliding
motility and the adhesins in the Apicomplexa (101).
The existence of an actomyosin-based mechanism for invasion was first implied by observations of the effect of cytochalasin B on Plasmodium invasion. Cytochalasin B blocks Plasmodium knowlesii merozoite invasion of red cells (106).
Merozoites will attach irreversibly to red cells and form a
vestigial parasitophorous vacuole but are inhibited from moving into the cell. Since red cells are unequivocally nonphagocytic, the effects of cytochalasin on Plasmodium invasion are
likely to represent drug disruption of parasite (not host cell)
microfilaments. However, for other apicomplexans, cytochalasin inhibition of invasion was sometimes attributed to inhibition of induced phagocytosis by the host cell. Active invasion of
Toxoplasma tachyzoites can be distinguished from the phago-
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CYTOSKELETON OF APICOMPLEXAN PARASITES
(86). MCP-1 lacks both a signal sequence and transmembrane
domain and is located in the parasite cytosol. The precise role
of MCP-1 in invasion remains obscure. The protein has an
amino-terminal domain that is conserved in bacterial and eukaryotic oxidoreductases (76). There are also a group of microneme proteins that may play a role in this junction that
localize to the moving junction and the (exposed) posterior
region during invasion (24, 57, 62).
Actin and Actin Binding Proteins
characterized in T. gondii (9). The single-copy gene encodes a
13.4-kDa protein. Toxoplasma ADF has a high degree of sequence similarity to other ADF homologs, particularly Acanthamoeba actophorin and plant ADFs. Toxoplasma ADF localizes to cytoplasm, especially under the plasma membrane.
Recombinant Toxoplasma ADF purified from E. coli binds
actin monomers and depolymerizes microfilaments in a pHindependent, concentration-dependent fashion.
One surprising finding is that a homolog of the actin monomer binding protein profilin has not been found in the Apicomplexa. In its place, Toxoplasma has apparently substituted
a novel protein, toxofilin (125). Toxofilin is a 27-kDa actin
monomer binding protein that was originally isolated from
Toxoplasma extracts on G-actin affinity columns. In pyrene
actin assays, toxofilin inhibits actin polymerization, acting as an
actin-sequestering protein. It also slows microfilament disassembly through a filament end-capping activity. A single-copy
gene encodes toxofilin. The protein has a pI of 9.63 and two
coiled-coil domains and lacks consensus motifs or any similarity to known proteins. Overexpression of green fluorescent
protein (GFP)-tagged toxofilin in vertebrate cells disrupts
stress fibers and reduces microfilament levels by half. Toxofilin
localizes to the apical cytoplasm in intracellular Toxoplasma
but is found at the posterior of invading parasites. In motile
parasites, toxofilin is localized throughout the entire cytoplasm.
Myosin
Evidence for myosins in the Apicomplexa was first suggested
by observations that T. gondii gliding motility and host cell
invasion are reversibly inhibited by the myosin inhibitors BDM
(an ATPase inhibitor) and KT5926 (a myosin light-chain kinase inhibitor) (35). BDM also blocks motility and invasion in
Plasmodium and in Cryptosporidium (56, 123). The effect of
BDM is likely to be myosin specific; however, KT5926 blocks
host cell attachment and motility by inhibiting the secretion of
adhesins, proteins required for motility and cell attachment
(Table 1). Immunofluorescence with heterologous myosin antibodies or antisera that recognizes the highly conserved myosin peptide LEAF localizes to the anterior pole or to a circumferential pattern that overlaps with the distribution of actin in
T. gondii and P. falciparum (35, 144, 187). Immunoelectron
microscopy of hypotonically swelled parasites (the plasma
membrane is separated from the IMC) demonstrates that actin
is associated with the IMC and that myosin is associated with
the plasma membrane (Fig. 3C) (35). The LEAF peptide antiserum identifies a 90-kDa band in T. gondii lysates, and heterologous myosin antibodies label a Plasmodium protein of 86
kDa (187).
Apicomplexan myosins are highly atypical and were ultimately cloned in Toxoplasma by using degenerate PCR of
conserved regions of the motor domain (70). A similar strategy
has been used to identify myosins in Plasmodium, Neospora,
Eimeria, Sarcocystis, Babesia, and Cryptosporidium (69). All
apicomplexan myosins are extremely similar, suggesting that
the diversity of myosins in these parasites is extremely limited
(69). Phylogenetic analysis of the myosins places these motors
in a novel, highly divergent class (XIV) in the myosin superfamily (69, 70). Apicomplexan myosins range from 91 to 125
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Genes encoding actin have been cloned in several members
of the Apicomplexa (36, 83, 119, 189–191). In T. gondii and C.
parvum, actin is encoded by a single-copy gene (36, 83, 119). In
Cryptosporidium, the actin gene is intronless, and in Toxoplasma it has one intron. In P. falciparum, the situation is akin
to that for ␣-tubulin. There are two genes encoding actin
(189–191). One gene (actin I) is intronless and is expressed
throughout the parasite life cycle. In contrast, the P. falciparum
actin II gene has an intron and is transcribed only in the sexual
stages (191). The amino acid sequence of actin II is divergent
from that of previously characterized actins. Additionally, relative to the high degree of conservation shown by most actins,
the 79% amino acid sequence similarity between Plasmodium
actin I and actin II is quite low. Actin-related proteins (arps)
have not been characterized in the Apicomplexa yet, but sequences in the Plasmodium genome are annotated as putative
arps.
F-actin affinity chromatography has been used to isolate
actin binding proteins from P. knowlesii and P. falciparum
merozoites and from Toxoplasma tachyzoites (54, 125, 173). In
P. knowlesii, five major proteins with molecular masses of 75,
70, 48, 40, and 32/34 kDa are eluted from F-actin columns
(173). The 70-kDa protein has been identified as heat shock
protein 70 (HSC70). The 32/34-kDa doublet coelutes with
HSC70 from columns or in gel filtration chromatography; however, the identity of these proteins remains unknown. Highly
enriched fractions of the Plasmodium HSC70-HSC32-HSC34
complex inhibit rabbit skeletal muscle actin polymerization in
vitro. Biochemical experiments have established that this is due
to a capping activity that is Ca2⫹ independent and is inhibited
by PIP2.
Homologs of two widely conserved actin-associated proteins, coronin and actin-depolymerizing factor (ADF), have
been characterized in the Apicomplexa. Coronin is a WD repeat containing actin binding protein that was first characterized in Dictyostelium discoideum, where it is essential for
phagocytosis and motility. WD repeats (a tryptophan-aspartic
acid motif) are found in diverse proteins; this motif is thought
to mediate protein-protein interactions. Homologs of coronin
are found in a large variety of eukaryotes, ranging from humans to C. elegans to yeast. A coronin homolog has been
described in P. falciparum and is encoded by a single-copy gene
(174). Compared to Dictyostelium coronin, the Plasmodium
protein has conserved residues throughout the entire protein.
A monoclonal antibody to D. discoideum coronin detects a
42-kDa protein in Triton X-100-insoluble extracts of P. falciparum schizonts. ADF/actophorin/cofilin is a widely conserved
low-molecular-weight actin monomer-sequestering protein
with filament-severing activity. An ADF homolog has been
31
32
MORRISSETTE AND SIBLEY
MANIPULATION OF THE HOST CYTOSKELETON
BY APICOMPLEXAN PARASITES
Reorganization of the Microvilli of Intestinal
Epithelia by Cryptosporidium
Like other apicomplexans, C. parvum resides in a parasitophorous vacuole within the host cell, in this case the intestinal
epithelium. However, the Cryptosporidium intracellular vacuole is extracytoplasmic, remaining at the apical surface of infected cells, in the region of the microvilli. The parasite induces
host cell cytoskeletal rearrangement including the formation of
branched microvilli clustered around the parasitophorous vacuole (55, 93). Moreover, Cryptosporidium induces the formation of a junctional complex that lies between it and the cytoplasm of the infected epithelial cell, keeping the parasite in the
region of the microvilli. The junctional complex is associated
with a plaque containing host cell actin. Accumulation of host
cell actin, arp2, arp3, neural Wiskott-Aldrich syndrome protein
(N-WASP), vasodilator-stimulated phosphoprotein (VASP),
and ␣-actinin begins before entry is complete and these proteins localize beneath the invading parasite (45, 46). The
plaque does not contain other actin binding proteins found in
the intestinal epithelium, such as the catenins, zyxin, or plakoglobin (45). As parasites grow within the host cell, ␣-actinin is
lost from the plaque, but the plaque size continues to increase
to accommodate the increasing size of the replicating parasites.
In addition to the above results, other have described localization of phosphotyrosine and villin at the site of parasite attachment (55). Expression of dominant negative constructs of
Scar1 or N-WASP in host cells blocks Cryptosporidium invasion, suggesting that parasite-induced host cell actin reorganization is required for invasion (45, 46).
Plasmodium Modification and Mimicry of
Erythrocyte Cytoskeletal Proteins
Infection with Plasmodium merozoites results in dramatic
changes to the shape and biochemical properties of the parasitized red cell. Red cells infected with Plasmodium have increased phosphorylation of band 4.1, and a cysteine protease
from the parasite cleaves red cell ankyrin (96, 128). P. falciparum induces knob formation of the surface of red cells, and the
normal discoid shape of these cells becomes spherical (176).
These structural alterations contribute to sequestration of infected red cells in organ capillaries, preventing their circulation
and exposure to the spleen (107). The Plasmodium proteins
RESA, MESA, and HRP-1 are anchored to the red cell membrane by association with spectrin, band 4.1, and the band 3
binding domain of ankyrin, respectively (17, 94, 97, 122). P.
falciparum growth is decreased in human erythrocytes containing abnormal spectrin or band 4.1 (142). Parasites invade these
cells normally, suggesting that an intact red cell membrane
skeleton is required for parasite growth.
Plasmodium erythrocytic stages also synthesize proteins
which are similar to ankyrin and spectrin and which are hypothesized to play a role in reorganization of the cytoskeleton
of red cells. Plasmodium chabaudi ROPE (repetitive organellar protein) has a structure similar to that of spectrin (188).
This 229-kDa protein is localized to the apical end of merozoites, possibly in the rhoptries. ROPE has characteristics of a
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kDa and include the smallest myosins characterized thus far
(70, 73). There is a high degree of sequence conservation
among all apicomplexan myosins throughout their whole
length. All have short tails that do not have homology to any
other myosin tails. However, these tails do contain a highly
basic charge distribution similar to myosin I family members,
suggesting that the apicomplexan myosins may interact with
membranes. Generally, myosin motors have three domains;
the amino terminus contains the motor domain, the central
“neck” region binds light chains and acts as a lever arm, and
the tail is diverse, carrying out different functions such as
targeting to subcellular regions or binding to cargo (18, 102).
The apicomplexan myosins do not contain the strictly conserved glycine residue at the fulcrum point of the lever arm and
generally lack IQ motifs that bind calmodulin and calmodulinrelated proteins (68, 70, 73). Additionally, the Toxoplasma
myosins do not follow the TEDS rule, i.e., the presence of an
acidic or phosphorylatable residue at a precise site close to the
actin binding region (70). In lower eukaryotes, this residue is
crucial for stimulation of the ATPase of class I myosins, but
other exceptions to this rule have been described. Both the
absence of an IQ motif and the nonadherence to the TEDS
rule suggest that these motors may be regulated in a novel
fashion.
T. gondii expresses five class XIV myosins: TgM-A, TgM-B,
TgM-C, TgM-D, and TgM-E (68–70, 73). TgM-A is 93 kDa
and lacks a discernible neck domain and IQ motifs (70).
Epitope-tagged TgM-A localizes beneath the plasma membrane (73). Mutational analysis has established that a pair of
arginine residues is essential to target TgM-A to the periphery
(73). Since ectopically expressed TgM-A in HeLa cells does
not target to the plasma membrane, peripheral localization in
parasites may require a membrane-associated receptor. The
P. falciparum homolog of TgM-A (PfM-A/Pf-myo1) is synthesized in mature schizonts and is present in merozoites but
vanishes after the parasite enters the red cell (123). PfM-A is
associated with the particulate parasite fraction, and immunofluorescence and immunogold analysis shows that PfM-A localizes to the periphery of mature schizonts and merozoites.
TgM-B and TgM-C are the products of differential RNA
splicing and are 114 and 125 kDa respectively (70). They are
identical throughout their head and neck domains and diverge
in their distal tail structures. Both contain a single IQ motif.
TgM-B has not been localized, but TgM-C localizes to a juxtanuclear region toward the apical pole of the parasite, consistent with an association with the Golgi apparatus (70, 73).
TgM-D is a 91-kDa protein that has a punctate peripheral
localization (73). TgM-E is the most recently discovered myosin and is currently being characterized (69, 73). Biochemical
studies have established that the myosins bind actin in the
absence but not the presence of ATP and that they are tightly
associated with membranes (68, 73). The peripheral localization of TgM-A and of the GFP–TgM-A tail fusion is not
dependent on an intact F-actin cytoskeleton (73). Truncation
of the tail domains of TgM-A or TgM-D abolishes their peripheral localization and tight membrane association; fusion of
the TgM-A or TgM-D tail to GFP is sufficient to confer plasma
membrane localization (73).
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CYTOSKELETON OF APICOMPLEXAN PARASITES
cytoskeletal protein. A 364-amino-acid repetitive region based
on 32 11-mer repeats suggests that the protein forms an helical
coiled-coil triple helix containing a leucine-histidine zipper.
Strikingly, this three-dimensional arrangement resembles the
structure of spectrin. It has been postulated that ROPE may be
involved in invasion, by interacting with the erythrocyte cytoskeleton via molecular mimicry of spectrin. P. falciparum
expresses an 88-kDa phosphoprotein that is nearly identical to
the amino-terminal region of ankyrin, a region of the protein
that binds band 3 (170). This protein may also help the parasite
reorganize the membrane skeleton via molecular mimicry.
Theileria Exploitation of Host Cell Microtubules
CONCLUSIONS
Although researchers studying apicomplexan parasites have
amassed many data regarding the cytoskeleton, we still are
missing explicit evidence linking cytoskeletal components to
cellular properties. The following discussion suggests possible
links between the cytoskeletal elements and behavioral traits
that are common to the members of the Apicomplexa. One
obvious generalization is that in these cells, the subpellicular
network and the subpellicular microtubules are critical to cell
shape while actin and myosin are essential for motility and
invasion. This is not to say that future work will not uncover
additional functions for the cytoskeleton.
Although apicomplexan parasites are profoundly deformed
during host cell invasion, they retain membrane integrity during host cell entry. This may be ascribed to their robust arrangement of plasma membrane, IMC, and subpellicular network. The newly identified proteins IMC-1 and IMC-2 are
implicated in subpellicular network formation (98). In addition
to the obvious questions of how these proteins form filaments
and how filament assembly is regulated is the issue of how this
network associates with the IMC. The lattice of intramembranous particles observed after freeze fracture of the pellicle
could reflect the transmembrane domains of receptors for the
subpellicular network; however, the identity of these highly
organized particles remains undetermined, as does any physical connection between the particles and the lattice proteins
(43, 113, 124).
Apicomplexan parasites multiply by endodyogeny or
schizogeny. As described above, these processes require independent regulation of spindle and subpellicular microtubules.
Perhaps consequently, subpellicular microtubules and spindle
microtubules are organized by different MTOCs: the apical
polar ring and the spindle pole plaque/centrioles, respectively.
In endodyogeny, parasites must discriminate between maternal
and daughter apical polar rings and between maternal and
daughter subpellicular microtubules. In schizogeny, daughter
cell budding is induced after movement of multiple nuclei to
the periphery of a maternal cell that lacks subpellicular microtubules. It is likely that the spindle pole plaques then induce or
coordinate the formation of the apical polar rings, coupling
each nucleus with a set of subpellicular microtubules. Daughter cell budding is distinct from vertebrate cytokinesis. In fact,
inhibitor studies suggest that parasite scission may not utilize
microfilaments such as are required at the vertebrate cleavage
furrow (149). However, a recent study (28a) of the alternatively
spliced myosins MyoB and MyoC in Toxoplasma demonstrates
that overexpression of MyoB causes defects in cell division,
and the parasites make extremely large residual bodies.
Tagged MyoB localizes in a punctate cytosolic pattern and
tagged MyoC localizes to the apical and posterior polar rings
of tachyzoites. These latter observations suggest that MyoB
and MyoC may play a role in parasite cell division, implicating
an acto-myosin ring in parasite scission. Recent microtubule
inhibitor studies show that subpellicular microtubule assembly
can be disconnected from nuclear division, creating Toxoplasma tachyzoites that lack nuclei, although budding and scission
from the maternal mass is completed (115). Multiple MTOCs
permit apicomplexans to control nuclear division independently from cell polarity and cytokinesis. Although this grants
greater cell cycle flexibility to these parasites, it abolishes the
checks for coregulation of nuclear division and cytokinesis that
are found in other eukaryotes.
Rigidity, cell shape, and apical polarity are provided by the
subpellicular microtubules, and the apical polar ring organizes
these microtubules (120, 134, 168). The apical polar ring represents a MTOC that is unique to the Apicomplexa. We know
very little about its genesis and nothing about its component
proteins. If the apical polar ring controls daughter cell budding, it must be replicated in a highly regulated fashion. It will
also be informative to understand how it nucleates the subpellicular microtubules. The number of subpellicular microtubules and their organization are invariable within a life cycle
stage of a particular species of apicomplexan parasite. The
apical polar ring is a highly ordered structure and may contain
signals that determine the number and placement of subpellicular microtubules.
Like the subpellicular network, the subpellicular microtu-
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With respect to behavior and morphology, Theileria parva
sporozoites are certainly the most distinct members of the
Apicomplexa and do not conform to many of the general
properties of the Apicomplexa described in this review. Theileria sporozoites are bounded by a simple plasma membrane
structure and lack both an inner membrane complex and subpellicular microtubules. Host cell entry does not require apical
orientation of the Theileria sporozoite and occurs by a zippering rather than a moving-junction mechanism (49, 50, 79, 147,
148, 152). Once inside the host cell, the parasite escapes from
the parasitophorous vacuole and takes up residence in the
cytoplasm. The parasite plasma membrane becomes coated
with a number of host cell-derived microtubules organized in
arrays tangential to the sporozoite surface (49, 79, 117, 147,
175). Sporozoite-associated microtubules are highly resistant
to nocodazole disruption (147, 175). These parasites also activate NF-B, specifically inducing clonal expansion of infected
cells (193). Infected lymphocytes will proliferate indefinitely in
culture until antiparasitic drugs halt unchecked replication
(193). Sporozoites undergo nuclear divisions to form a
multinucleate schizont. The host microtubules associated with
the schizont are captured by the spindle of the proliferating
host lymphocytes, pulling fragments of the schizont into each
daughter lymphocyte (117, 175). The coupling of induction of
host cell proliferation and association with host cell microtubules ensures that infected cells are specifically expanded and
that the resulting progeny continue to harbor the parasite.
33
34
MORRISSETTE AND SIBLEY
The precise mechanism by which parasites use an acto-myosin motor to generate motility is unclear. Since actin filaments
are rare and since the apicomplexan myosins lack typical regulatory domains, it has been suggested that the movement of
myosin is limited by filament generation. Consistent with this,
jasplakinolide-treated parasites show increased motility, although drug treatment inhibits rather than enhances parasite
invasiveness (151). For the myosin motors to have force-generating movement, the microfilaments must be tethered so that
they remain in place. Actin filaments may be immobilized by
interactions with the IMC, and the rigidity provided by the
subpellicular network and the subpellicular microtubules could
provide the extra stability required for myosin movement to
transport adhesins to the posterior end of the parasite. Myosin
movement along the actin filaments would lead to capping of
adhesins down the length of the parasite and ultimately to
gliding motility or invasion. Gliding motility is a trait shared
with gregarines, apicomplexan parasites of invertebrates that
are quite distinct from other members of the phylum described
here (84, 85, 184).
This review has generalized the behavior of apicomplexans
as a group. By doing so, we have undoubtedly glossed over
differences, particularly with the more divergent members of
this phylum. However, in many cases, atypical attributes are
particular to the life cycle stage of the parasite rather than
characteristic of the organism as a whole. This is most clearly
illustrated with P. falciparum merozoites and Theileria sporozoites. Although most apicomplexans have many subpellicular
microtubules and use an acto-myosin mechanism to glide and
to invade cells, Plasmodium merozoites have reduced these
traits and Theileria sporozoites have eliminated them. P. falciparum merozoites have a drastically scaled-back set of subpellicular microtubules that may reflect the greatly reduced size of
merozoites relative to liver and insect stage parasites; both
traits may be dictated by the smaller size of the host red cells.
Merozoites do not display gliding motility, although they actively invade red cells in an actin-dependent fashion. Other
Plasmodium stages are larger, have prominent subpellicular
microtubules, and are clearly motile. Similarly, Theileria sporozoites lack subpellicular microtubules and infect lymphocytes
after entering in a novel and nonmotile fashion. Tick-stage
Theileria kinetes are larger and have subpellicular microtubules. Although very little work has been done on this stage, we
assume that they are motile and actively invade host cells.
Nonetheless, there are clearly dangers in overgeneralizing the
behavior of apicomplexan parasites. Other exceptions to these
generalizations will undoubtedly be found within this diverse
group of parasites.
At present, the P. falciparum genome project is nearing
completion and several genome projects are under way for
other Plasmodium species. The Cryptosporidium genome is also
nearing completion; genome projects for Toxoplasma, Theileria, and Babesia are under way; and there are ongoing EST
projects for Toxoplasma, Neospora, Eimeria, Sarcocystis, and
Plasmodium. In the past few years, researchers have developed
techniques to transfect Toxoplasma and Plasmodium, to make
targeted deletions, and to create gene replacements. With the
amassed information about the cytoskeleton and these new
resources and tools, we are truly poised to understand the
mechanisms underlying cytoskeletal functions in the apicom-
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bules also have intimate connections with the inner membrane
complex (3, 113, 131, 195). To understand these interactions, it
will be necessary to identify and characterize MAPs. MAPs
may dictate subpellicular microtubule length and position under the pellicle. It is unclear why some apicomplexans distribute their microtubules uniformly beneath the pellicle while
others center one microtubule beneath one-third of the circumference and evenly space the remainder below the other
two-thirds of the pellicle. In studies of motility, it is clear that
the convex and concave sides of the parasite are not equivalent.
The asymmetry of microtubules in some apicomplexans may
simply reflect areas that are more or less closely involved in
force generation during motility or other essential functions.
Subpellicular microtubules may contribute to motility by providing tracks that direct the acto-myosin-based capping activity. Toxoplasma tachyzoites and Plasmodium merozoites with
shortened subpellicular microtubules (due to drug treatment)
are noninvasive, supporting this notion (14, 115). However, it
is not absolutely clear how microtubules could serve as tracks
for the acto-myosin system, since actin and myosin are believed
to act between the plasma membrane and the IMC while
microtubules localize to the cytoplasmic face of the IMC (35,
120).
Many studies have implicated actin in apicomplexan motility, although apicomplexan microfilaments are apparently
quite labile under most circumstances. Polymerized actin is
observed only in the presence of jasplakinolide, and in untreated cells nearly all the actin is found as G-actin (36, 151).
The apical actin filaments observed after treatment of Toxoplasma with jasplakinolide may reflect the location of actin
regulators that nucleate or otherwise facilitate filament polymerization (151). Alternately, the apical localization of an Factin projection after jasplakinolide treatment may represent
the “path of least resistance” since the apical region is the only
area of the pellicle not surrounded by three unit membranes
and the subpellicular network. The short-lived nature of microfilaments suggests that actin assembly and disassembly are
closely regulated. The Plasmodium HSC70 complex caps Factin, limiting filament growth, and the apicomplexan homologs of ADF/cofilin are likely to sever filaments and sequester monomers, facilitating rapid disassembly of actin filaments
(9, 173). Additionally, in Toxoplasma, actin may be kept monomeric by sequestration by toxofilin, a novel monomer binding protein (125). BLAST searches of the Cryptosporidium and
Plasmodium genomes do not identify homologs of toxofilin,
suggesting that distinct proteins may provide this function in
other apicomplexans (unpublished data).
Myosin motors are also implicated in motility and invasion
(35, 56, 66, 123). The apicomplexan myosins are quite divergent from myosins in other organisms, constituting a new class
of motors in the myosin family (68–70, 73, 123). Apicomplexan
myosins are all quite similar but have different subcellular
localizations. Myosin-A is most likely to be involved in motility
since it is found beneath the plasma membrane, whereas myosin-B is located to the Golgi and myosin-D is found on vesicles, consistent with roles for these latter motors in membrane
traffic (68, 73, 123). Ectopic expression of myosin-A has shown
that it does not localize to the plasma membrane in nonapicomplexans and therefore must be targeted to this region in
parasites by additional proteins (73).
MICROBIOL. MOL. BIOL. REV.
VOL. 66, 2002
CYTOSKELETON OF APICOMPLEXAN PARASITES
plexan parasites. Disease caused by these protozoa has tremendous medical and economic impact worldwide. For the cell
biologist, the unique biology of the Apicomplexa represents an
intriguing departure from standard eukaryotic behaviors; for
the clinician, these distinctions may represent unique drug
targets.
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We thank Antonio Barragan, Audra Charron, John Cooper, Susan
Dutcher, Olivia Giddings, Dan Goldberg, Wallace Marshall, Alissa
Weaver, and Dawn Wetzel for commenting on the manuscript.
N.S.M. is supported by Individual NRSA fellowship F32 GM2048401A1 and was previously supported by an NIH training grant in Infectious Disease held by the Washington University School of Medicine. (T32; AI-0717221).
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