Journal of Experimental Botany, Vol. 62, No. 15, pp. 5249–5266, 2011
doi:10.1093/jxb/err227 Advance Access publication 23 August, 2011
DARWIN REVIEW
The cytoskeleton in plasmodesmata: a role in intercellular
transport?
Rosemary G. White1,* and Deborah A. Barton2
1
2
Commonwealth Scientific and Industrial Research Organisation, Division of Plant Industry, Canberra, ACT 2601, Australia
School of Biological Sciences, University of Sydney, NSW 2006, Australia
* To whom correspondence should be addressed. E-mail: [email protected]
Received 19 May 2011; Revised 27 June 2011; Accepted 28 June 2011
Abstract
Actin and myosin are components of the plant cell cytoskeleton that extend from cell to cell through plasmodesmata
(PD), but it is unclear how they are organized within the cytoplasmic sleeve or how they might behave as regulatory
elements. Early work used antibodies to locate actin and myosin to PD, at the electron microscope level, or to
pitfields (aggregations of PD in the cell wall), using immunofluorescence techniques. More recently, a green
fluorescent protein (GFP)-tagged plant myosin VIII was located specifically at PD-rich pitfields in cell walls.
Application of actin or myosin disrupters may modify the conformation of PD and alter rates of cell–cell transport,
providing evidence for a role in regulating PD permeability. Intriguingly, there is now evidence of differentiation
between types of PD, some of which open in response to both actin and myosin disrupters, and others which are
unaffected by actin disrupters or which close in response to myosin inhibitors. Viruses also interact with elements of
the cytoskeleton for both intracellular and intercellular transport. The precise function of the cytoskeleton in PD may
change during cell development, and may not be identical in all tissue types, or even in all PD within a single cell.
Nevertheless, it is likely that actin- and myosin-associated proteins play a key role in regulating cell–cell transport,
by interacting with cargo and loading it into PD, and may underlie the capacity for one-way transport across
particular cell and tissue boundaries.
Key words: Actin, cell–cell transport, cytoskeleton, myosin, plasmodesma.
The plant cytoskeleton
The plant cytoskeleton is a dynamic network of protein
filaments that, together with protein motors and a wide
range of accessory proteins that modify its behaviour, is
involved in every aspect of cell growth and development. As
in animal cells, one major function is to shuttle cargo within
cells, a particularly important function in mature, vacuolate
plant cells. Actin filaments, assembled from actin monomers, are responsible for this cytoplasmic streaming, and in
the cortical regions of plant cells are extremely dynamic,
with individual filaments mostly short lived (<30 s; Staiger
et al., 2009). The second major component of the plant
cytoskeleton, microtubules, are hollow cylinders assembled
from ab tubulin heterodimers which form actively organized arrays in the cell cortex (Shaw et al., 2003) whose
primary function in interphase cells is to guide the direction
of cellulose microfibril deposition in the cell wall (Paredez
et al., 2006). Microtubules are also dynamic, with turnover
half-times of 30–90 s in interphase plant cells (Hush et al.,
1994; Shaw et al., 2003). Despite their stochastically
dynamic behaviour, actin filaments and microtubules form
tracks along which the motor proteins myosin and kinesin,
respectively, traffic cellular cargo. Cytoskeleton organization and turnover are constantly modified by a highly
diverse and still-expanding array of actin-binding proteins
(ABPs) and microtubule-associated proteins, including nucleating, severing, end-binding, capping, or bundling proteins, as well as proteins that form cross-bridges within and
between these filament systems (e.g. Wasteneys and Yang,
Abbreviations: ABP, actin-binding protein; BDM, butane dione monoxime; ER, endoplasmic reticulum; GFP, green fluorescent protein; MP, movement protein; NEM,
N-ethylmaleimide; PD, plasmodesma(ta); SEL, size exclusion limit; (T)EM, (transmission) electron microscopy; TMV, tobacco mosaic virus.
ª The Author [2011]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
5250 | White and Barton
2004; Collings et al., 2006; Hussey et al., 2006; Hamada,
2007; Staiger et al., 2010).
In addition to its primary role in intracellular transport,
the cytoskeleton is implicated in intercellular transport of
cytoplasmic molecules via plasmodesmata (PD), the membrane-lined channels linking plant cells across the cell wall
(Fig. 1). Both actin and myosin have been immunolocalized
to these intercellular channels (White et al., 1994; Blackman
and Overall, 1998; Radford and White, 1998; Reichelt et al.,
1999; Baluška et al., 2001a, 2004), and cytological evidence
indicates a role for both in cell–cell transport. Interestingly,
although there is no specific evidence for microtubules
or tubulin within PD, both microtubules and actin
filaments can direct certain cargo towards PD, a process
that appears to be high-jacked by viruses during infection
of plant cells (reviewed by Harries et al., 2010). Here, PD
structure is described, then similar intercellular connections in other kingdoms are considered. The evidence for
cytoskeletal proteins as components of PD is then reviewed
in detail and their role in cell–cell transport is assessed
(see also earlier reviews by Overall et al., 2000; Aaziz et al.,
2001).
Fig. 1. Ultrastructure of plant plasmodesmata (PD), revealed by TEM analysis of intact tissues. (A) Longitudinal section showing a young
Allium cepa root PD with electron-dense material (arrows) in the cell wall around the neck region (with kind permission from Springer
Science+Business Media: Protoplasma Callose deposition at plasmodesmata. 201, 1998. 30–37 Radford JE, Vesk M, Overall RL. Fig.
4b). (B) Longitudinal section of a single PD, showing cytoplasmic endoplasmic reticulum (ER) narrowing into the desmotubule (DT; large
arrowhead), which links the ER of two adjacent cells. The plasma membrane (PM) and cytoplasmic sleeve (small arrows) are also
continuous from cell to cell. (courtesy of JE Radford, unpublished). (C) Diagram highlighting PD components, showing where cytoskeletal
elements may fit into this structure. An alternative model is possible, with actin closest to the PM, and myosin attached to the DT.
(D) Longitudinal sections of some PD show constrictions only at the neck region, with the PM widely separated from the DT in the region
deep within the cell wall. Note electron-dense material (arrows) on the cytoplasmic side of the PM, and occasional strands (small arrow)
connecting DT and PM. (Hordeum vulgare root, courtesy of JE Radford, unpublished). (E) Longitudinal sections of PD from high-pressure
frozen and freeze-substituted tobacco leaf tissue showing open DT (arrowheads) with strands linking them to the PM (small arrows).
Microtubules (MT) can be seen adjacent to the PM. (courtesy of MEA Schoenwaelder, unpublished). (F) Transverse section of tobacco
leaf PD, showing particles possibly attached to or embedded in both desmotubule wall (DW) and plasma membrane (IPM, inner plasma
membrane) and connected by strands across the cytoplasmic sleeve (with kind permission from Springer Science+Business Media:
Protoplasma Substructure of freeze-substituted plasmodesmata. 169, 1992. 28–41. Ding B, Turgeon R, Parthasarathy MV. Fig. 4b).
(G) Transverse section in which particles (arrowhead) are mainly associated with the DT, and strands (arrows) appear to link the DT and
PM (double arrow). (Egeria densa leaf, JE Radford; reprinted from Trends in Plant Science 1, Overall RL, Blackman LM. A model of the
macromolecular structure of plasmodesmata. 307–311, 1996, with permission from Elsevier). Bars¼50 nm in A–E; 10 nm in F, G.
The cytoskeleton in plasmodesmata | 5251
Tunnels linking cells—plasmodesmata
Plasmodesmata are minute cell wall channels connecting the
plasma membrane and cytoplasm of neighbouring plant cells,
and contain a central core of tightly compressed endoplasmic
reticulum (ER), the desmotubule, which links the ER
networks of most cells (Fig. 1A–C). Newly formed PD
commonly have proteinaceous material at the neck (Fig. 1A)
and a narrow channel between the ER and enclosing plasma
membrane (Fig. 1A–C). In rapidly frozen and freezesubstituted tissue, this channel, and occasionally the desmotubule, may widen in places (Fig. 1D, E), with connections
between the two membranes (Fig. 1D–G). Cytoplasmic
components, including metabolites, signals, or developmental regulators such as transcription factors and RNA
molecules (reviewed in Xu and Jackson, 2010), move from
cell to cell through the small cylinder of cytoplasm between
the plasma membrane and desmotubule, and this narrow
space is where cytoskeletal components of PD are likely to
be found. Cytoskeletal activity at the neck regions of PD
may also direct and/or regulate movement of molecules into
the cytoplasmic channel. While small cytoplasmic molecules
[including the 27 kDa green fluorescent protein (GFP)]
diffuse freely between many plant cells, transport of larger
molecules is restricted and may require interaction with PDspecific proteins to facilitate movement (Roberts and
Oparka, 2003). This size exclusion limit (SEL) is higher
between cells in developing tissues and lower in more
mature tissues (reviewed in Xu and Jackson, 2010), suggesting a greater degree of intercellular transport regulation for
older PD. Deposition of callose in the cell wall at the neck
regions of PD (Fig. 1A) can modify their SEL and
ultimately close the cytoplasmic pathway. Actin and myosin
are also implicated in regulating the SEL of PD, but the
mechanics of this are as yet, unknown.
Small proteins in the ER lumen (Barton et al., 2011) and
both lipids (Grabski et al 1993; Martens et al., 2006) and
transmembrane proteins in the ER membrane (GuenouneGelbart et al., 2008) can move between cells via the
desmotubule. However, there appears to be no intercellular
exchange of either lipid or protein within the plasma
membrane through PD. As both actin and myosin are
associated with other ER domains in plant cells, it is
possible that they play a role in regulating intercellular
transport via the desmotubule. They may also help to
maintain the structure of the narrow cytoplasmic sleeve by
forming links between the ER and plasma membranes via
other PD proteins.
Tunnels linking cells—from Anabaena to
animals
The lower members of the plant kingdom also show a range
of PD-like connections (Cook et al., 1997; Cook and
Graham, 1999; Raven, 1997, 2005), many of which have
a central strand of ER surrounded by a connecting tube of
plasma membrane. Certain groups of fungi form cell-like
compartments linked by a diverse range of intercellular
connections, a small subset of which are identical in
structure to plant PD (Hawker et al., 1966; Markham,
1994; Lutzoni et al., 2004). Smaller and less complex
connections, termed ‘microplasmodesmata’, are found between cells in cyanobacterial filaments (Lang and Fay, 1971;
Giddings and Staehelin, 1981; Flores and Herrero, 2010).
In the animal kingdom, structures called tunnelling nanotubes (Rustom et al., 2004; Gerdes et al., 2007) are detected
between cells in an ever-increasing range of tissues. As in
plants, these intercellular connections are formed from
narrow tubes of plasma membrane and generally contain
a bundle of actin microfilaments. They allow direct transfer
of macromolecules from cell to cell, including intercellular
virus transfer, similar to PD (Sowinski et al., 2008).
Tunnelling nanotubes also traffic lipid-bound organelles
(Gurke et al., 2008), seen very rarely in PD (Fig. 4.11 in
Waters, 2004), and they are routes for migration of plasma
membrane lipids and proteins, which has not been detected
in PD.
These cell–cell channels differ in evolutionary origin and
composition, but converge on the same essential function,
which is to facilitate the exchange of large signalling or
information molecules, particularly nucleic acids, between
cells while protecting their contents from exposure to the
environment outside the cell membrane. It can be concluded, therefore, that direct cytoplasmic connections between neighbouring cells are essential to most multicellular
organisms. So far, cytoskeletal elements have been detected
only in PD and tunnelling nanotubes, but this may change
with further investigation.
Cytoskeletal components of plasmodesmata
A number of different approaches have been used to locate
cytoskeletal proteins in or near PD. Proteomic analysis of
relatively pure cell wall fractions has identified a number of
putative PD proteins (e.g. Epel, 1994; Epel et al., 1996;
Blackman et al., 1998; Faulkner et al., 2005; reviewed in
Faulkner and Maule, 2011), but cytoskeletal proteins are
not prominent in PD-enriched cell fractions, although
amino acid sequences with some similarity to animal
tropomyosins have been identified (Faulkner et al., 2009).
A more specific approach is to use antibodies against
targeted cytoskeletal proteins in pull-down assays, on protein blots, or in immunolocalization studies.
Actin
Actin was first associated with PD in transmission electron
micrographs by immunogold labelling (Fig. 2A, B, Table 1;
White et al., 1994; Blackman and Overall, 1998) using
a monoclonal antibody to chicken gizzard actin that
recognizes an epitope (amino acids 23–28; McLean et al.,
1990b) on the external surface of actin monomers and
filaments (Lessard, 1988; McLean et al., 1990b; Lorenz
5252 | White and Barton
Fig. 2. Distribution of actin, myosin, and other cytoskeletal proteins by immunofluorescence and immunoelectron microscopy. In C, G,
and H, PD are oriented horizontally on the page; in all other panels, PD are oriented vertically on the page. (A) ImmunoEM using an
antibody to chicken gizzard actin shows sparse labelling of PD in Hordeum vulgare root tips (with kind permission from Springer
Science+Business Media: Protoplasma, Actin associated with plasmodesmata, 180, 1994, 169–184. White RG, Badelt K, Overall RL,
Vesk M. Fig. 2d). (B) Chara corallina PD are also labelled by the same antibody to chicken gizzard actin (from Blackman LM, Overall RL.
1998. Immunolocalisation of the cytoskeleton to plasmodesmata of Chara corallina. The Plant Journal 14, 733–741, with permission from
Wiley-Blackwell) (C) Rhodamine–phalloidin labelling shows fluorescence at pitfields and in actin strands leading up to the pitfields in
H. vulgare coleoptiles (with kind permission from Springer Science+Business Media: Protoplasma, Actin associated with plasmodesmata, 180, 1994, 169–184. White RG, Badelt K, Overall RL, Vesk M. Fig. 8b,c). (D) Occasionally, single filaments are detected crossing
cell walls of Nicotiana plumbaginifolia suspension-cultured cells (with kind permission from Springer Science+Business Media:
The cytoskeleton in plasmodesmata | 5253
Table 1. Location of cytoskeletal proteins to cell plates, PD, and/or pitfields
Cytoskeletal
protein
Localization technique; antibody used; species tested:
Reference
Actin
IEM; C4 anti-chicken gizzard actin; Hordeum vulgare, Nephrolepis exaltata,
Azolla pinnata PD in roots and rhizomes
IEM; monoclonal C4 anti-chicken gizzard actin; Chara corallina PD
White et al. (1994)
Fluorescent phalloidins; PD in H. vulgare coleoptiles, Nicotiana plumbaginifolia suspension cells
IF; monoclonal C4 anti-chicken gizzard actin; C. corallina PD
Actin-related proteins IF; anti-Dictyostelium discoideum Arp3; Zea mays pitfields
Myosin
IEM; polyclonal anti-bovine smooth and skeletal; Allium cepa, H. vulgare, Zea mays root PD
IF; polyclonal anti-bovine smooth and skeletal; Z. mays root PD
IEM, IF; monoclonal anti-mung bean myosin; C. corallina PD
Tropomyosin
Centrin
IEM, IF; anti-myosin VIII; Lepidium sativum roots—cell plates
IF; Z. mays roots—cell plates
IF; anti-175 kDa myosin from BY-2 cells; Nicotiana tabacum BY-2 cell plates
FP; ATM1–GFP myosin in N. tabacum BY-2 cell plates
FP; GFP–myosin VIII (ATM1) in pitfields of Nicotiana benthamiana leaf epidermis
IF; polyclonal anti-chicken gizzard tropomyosin; C. corallina PD,
pitfields in Arabidopsis roots and Allium porrum leaf epidermis
IEM, IF; anti-Chlamydomonas centrin; roots and florets of Brassica oleracea, A. cepa roots—cell plates, PD,
pitfields
Blackman and Overall
(1998)
White et al. (1994)
Blackman and Overall
(1998)
van Gestel et al. (2003)
Radford and White (1998)
Radford and White (1998)
Blackman and Overall
(1998)
Reichelt et al. (1999)
Reichelt et al. (1999)
Yokota et al. (2009)
van Damme et al. (2004)
Golomb et al. (2008)
Faulkner et al. (2009)
Blackman et al. (1999)
IEM, immuno-EM: shows localization to individual PD; IF, immunofluorescence: labelled antibodies, labelled phalloidin show localization at
pitfields; FP, GFP-tagging: shows localization at pitfields.
et al., 1993). More general localization of actin to PD
clusters, or pitfields, was detected using fluorescent phalloidins [fungal toxins that bind specifically to filamentous actin
(Barden et al., 1987; Cooper, 1987; White et al., 1994); Fig.
2C, D], or by immunofluorescence, using an antibody to
human actin (C4 clone; Baluška et al., 2001a, 2004).
However, it is curious that in many publications showing
well-preserved, immunofluorescently labelled actin filaments
there is no obvious labelling of pitfields, even in elongate
cells with pronounced filaments (e.g. Collings and Wasteneys, 2005; Collings et al., 2006; Barton and Overall, 2010),
or when pitfields are specifically examined (Chaffey and
Barlow, 2001).
One reason for this may be that visualization of
fluorescently tagged actin in PD pushes against current
optical microscope resolution limits. In the first place, the
sizes of individual PD, at 40–50 nm diameter and 100 nm
long in young cell walls, and actin filaments, at 4–8 nm
diameter, are well below the resolution limits of conventional fluorescence or confocal microscopes. Therefore,
fluorescently labelled actin at or within PD may not be
obvious unless the PD are clustered into pitfields. Secondly,
Protoplasma, Actin associated with plasmodesmata, 180, 1994, 169–184. White RG, Badelt K, Overall RL, Vesk M. Fig. 9i,j,k). (E) PD
may be quite heavily labelled in immunoEM using a polyclonal antibody to smooth and skeletal myosin (from Radford JE, White RG.
1998. Localization of a myosin-like protein to plasmodesmata. The Plant Journal 14, 743–750, with permission from Wiley-Blackwell). (F)
C. corallina PD are labelled in immunoEM by an antibody to mung bean myosin (from Blackman LM, Overall RL. 1998.
Immunolocalisation of the cytoskeleton to plasmodesmata of Chara corallina. The Plant Journal 14, 733–741, with permission from
Wiley-Blackwell). (G) Immunofluorescence labelling with polyclonal antibody to animal myosin highlights pitfields and strands in the
cytoplasm in Zea mays (from Radford JE, White RG. 1998. Localization of a myosin-like protein to plasmodesmata. The Plant Journal 14,
743–750, with permission from Wiley-Blackwell). (H) Tropomyosin localizes to pitfields in cell walls, and remains associated with the wall
in plasmolysed Arabidopsis root cells (reprinted from European Journal of Cell Biology 88, Faulkner CR, Blackman LM, Collings DA,
Cordwell SJ, Overall RL. Anti-tropomyosin antibodies co-localise with actin microfilaments and label plasmodesmata. 357–369. 2009,
with permission from Elsevier). (I) Treatment with cytochalasin D widened the necks of these Nephrolepis exultata PD, but not others
(with kind permission from Springer Science+Business Media: Protoplasma, Actin associated with plasmodesmata, 180, 1994, 169–184.
White RG, Badelt K, Overall RL, Vesk M. Fig. 11d). (J) Application of BDM to PD caused narrower neck regions in Arabidopsis. (from
Radford JE, White RG. 1998. Localization of a myosin-like protein to plasmodesmata. The Plant Journal 14, 743–750, with permission
from Wiley-Blackwell). (K) Antibodies to Chlamydomonas reinhardtii centrin label only the neck region of PD (reprinted from European
Journal of Cell Biology 78, Blackman LM, Harper JDI, Overall RL. Localization of a centrin-like protein to higher plant plasmodesmata.
297–304. 1999, with permission from Elsevier). Bars¼50 nm in A, E, I, J; 100 nm in B, F; 5 lm in C, D, G, H; 2.5 lm inset in H;
25 nm in K.
5254 | White and Barton
since individual actin filaments are difficult to preserve in
plant cells using conventional aldehyde fixation treatments
(Goodbody and Lloyd, 1990; Baskin et al., 1996; Collings
et al., 2001; Meagher and Fechheimer, 2003; Barton and
Overall, 2010) they may be reduced or absent from PD in
aldehyde-fixed tissues. In some cases, immunolabelling
protocols incorporate a cell wall digestion step, which may
also disrupt pitfield actin. Thirdly, there are a number of
different actin isoforms in plants, including eight in
Arabidopsis (from 10 genes; reviewed in Meagher et al.,
1999; Meagher and Fechheimer, 2003), and it is conceivable
that actin in PD represents one or more specific isotypes in
vegetative or reproductive tissues (cf. McLean et al., 1990a),
so may not be recognized by antibodies raised against
animal actins. Even when plant-specific antibodies to highly
conserved epitopes are used, they may not recognize all
actin isovariants (Kandasamy et al., 1999; Meagher et al.,
1999; Meagher and Fechheimer, 2003). If the actin in PD is
in an unusual conformation, the epitope on the filament
surface may be hidden, and inaccessible for antibody
binding. However, before abandoning this approach, it
would be worthwhile to use plant- and isotype-specific
antibodies in immunoelectron microscopy (EM) analyses to
provide further information about the nature and location
of actin filaments at PD, especially since there has been only
one such study to date!
It is surprising that even with improved imaging technologies that push beyond the theoretical resolution limits,
such as super-resolution microscopy or electron tomography (Fitzgibbon et al., 2010; Bell and Oparka, 2011), we are
still ignorant as to how actin filaments physically associate
with PD. Are they present at PD neck regions only, or do
they traverse the length of the PD channel? If the latter, do
they form a continuous cytoskeletal pathway between cells?
Globular structures seen in the PD channel in transmission
electron microscopy (TEM) images (Overall et al., 1982;
Ding et al., 1992) appear similar to actin filaments in vitro
(Ding et al., 1991; Maciver et al., 1991). Together with the
immunolocation evidence above, this has given rise to
models incorporating actin filaments within PD channels
(e.g. White et al., 1994; Overall and Blackman, 1996;
Overall et al., 2000; Faulkner and Maule, 2011; Tilsner
et al., 2011), although some question whether any cytoskeletal elements could fit into the cytoplasmic sleeve (Bell and
Oparka, 2011; Tilsner et al., 2011). Actin microfilaments are
closely associated with cortical ER tubules (Quader et al.,
1987; Lichtscheidl et al., 1990; Boevink et al., 1998; Runions
et al., 2006) and microfilament removal disrupts the cortical
ER network (Wright et al., 2007), indicating a role for actin
in maintaining the ER network (Lichtscheidl and Weiss,
1988; Lichtscheidl et al., 1990; Boevink et al., 1998). Actin
filaments may bind to, and help maintain, the ER-derived
desmotubule within PD.
Alternatively, actin filaments may associate with the
plasma membrane (Ding et al., 1991), even within PD, since
it is unknown whether actin comprises the inner or outer
electron-dense or electron-lucent protein-sized structures
seen in TEM. Direct association of filamentous actin with
membranes occurs when the actin interacts with lipids at
specific sites (Gicquaud and Wong, 1994), and undergoes
a major conformational change to form highly ordered
arrays of actin filaments on the membrane surface
(Gicquaud, 1993, 1995). These arrays partially penetrate
the membrane surface (St-Onge and Gicquaud, 1990) and
would appear similar to interpretations of the ER- and
plasma membrane-associated proteins in PD (Ding et al.,
1992; Botha et al., 1993). If actin (or other proteins) in PD
were ‘recessed’ into one or other membrane, it would help
explain how all of the putative PD proteins might fit into
this tiny intermembrane space. A recent re-analysis of actin
filament structure suggests that it is quite flat (Oda et al.,
2009; Dominguez and Holmes 2011), and in this conformation filaments would be more likely to fit into PD, especially
at the neck. Occasionally, PD with very wide cytoplasmic
sleeves (but narrower necks) can be seen, especially in
freeze-substituted tissues (e.g. Dolzmann, 1965; Burgess,
1971; Tilney et al., 1991; Ding et al., 1992; Fig. 1b in
Overall et al., 2000), and here, perhaps, the actin is not
constrained to either membrane. At present, there are no
convincing data either way to indicate how actin is arranged
in PD.
Detection of fluorescently tagged actin within living PD
would provide more substantial evidence for its location
there, but development of stable lines expressing any plant
actin isoform tagged with a fluorescent protein has proven
intractable. Introduction of readily available derivatized
animal actins has not solved this problem, as Oregon greentagged animal actin injected into plant cells becomes
aggregated and does not integrate with plant actin filaments
(Jing et al., 2003). Instead, stable Arabidopsis lines expressing fluorescent protein-tagged actin-associated proteins,
such as fimbrin (Kovar et al., 2001; Wang et al., 2004),
actin-binding domain protein (McCurdy et al., 2001), mouse
talin (Kost et al., 1998), human talin (Takemoto et al.,
2003), or LIM-domain proteins (Papuga et al., 2010), all
highlight actin filaments and filament bundles within cells.
Interestingly, in none of these lines are there specific PD- or
pitfield-associated fluorescent structures, implying that if
actin is located there, it may not be in a standard
filamentous form with side-binding-associated proteins.
Currently there is no evidence for intercellular movement of
actin or fluorescently tagged ABPs between plant cells,
although rhodamine–phalloidin, and other fluorescently
tagged variants of these small actin-binding peptides, can
spread into neighbouring cells following injection (Cleary,
1995; Molchan et al., 2002).
A role for actin in plasmodesmal function?
With the assumption that cytoskeletal proteins in PD will
be regulated similarly to the cytoskeleton elsewhere in the
cell, a range of actin-disrupting or actin-stabilizing drugs
have been used to identify a role for actin filaments in
regulating PD transport between cells. Effects on transport
are measured by observing effects on cell–cell transmission
of cytoplasmic signals, such as electrical currents or
The cytoskeleton in plasmodesmata | 5255
fluorescent probes of different sizes. The time for the signal
to move into neighbouring cells, or the number of cells
traversed within a certain time, is compared following
different treatments and assumed to indicate the permeability of the PD cytoplasmic sleeve. Co-injecting the actin
disrupter, cytochalasin D, with a fluorescent 10 kDa
dextran into Nicotiana tabacum (Ding et al., 1996; Table 2)
or Nicotiana benthamiana (Su et al., 2010) mesophyll cells
substantially increased cell–cell transport of the dextran.
Conversely, co-injecting the actin-stabilizing fungal toxin,
phalloidin, with the dextran almost completely prevented its
movement. This suggests that actin filament dynamics may
play a role in regulating the pore size of PD between these
cells and, when filaments are disrupted, larger molecules can
move from cell to cell. When phalloidin favours filament
assembly and bundling over disassembly, perhaps the PD
pore is physically blocked, or binding of regulatory proteins
is perturbed, preventing transport. Supporting these findings, treatment of Nephrolepis exaltata root tips with
cytochalasin D induced widening of PD neck regions (Fig.
Table 2. Effects of cytoskeleton disrupters on cell–cell transport
Cytoskeletal inhibitor
Effects on PD or transport; specific tissue
tested
Actin filament disrupters; cytochalasin B or D; actin
polymerization inhibitors; latrunculin A or B
Widening of neck region; Nephrolepis exaltata
TEM analysis
rhizomes
Increased transport, Nicotiana tabacum mesophyll Dye transport
Increased transport; Nicotiana benthamiana
Dye transport
mesophyll
No effect; Setcreasea purpurea stamen hairs
Dye transport
No effect on structure; Hordeum vulgare roots, AzollaTEM analysis
pinnata roots
No effect; N. tabacum leaf trichomes
Dye transport
No effect; Tradescantia virginiana stamen hairs
Dye transport
No effect; Arabidopsis root tissues
Dye transport, GFP
transport
Dye transport
No effect; Egeria densa leaf tissues
Reduced transport due to reduced cytoplasmic
streaming; Chara sp.
Reduced transport; N. tabacum mesophyll
Profilin; actin-sequestering protein
VIGS against actin
Actin filament stabilizers; phalloidins
BDM, myosin inhibitor
Detection method
Increase or decrease depending on initial
conductance; Salvinia auriculata trichomes
Increased transport; N. tabacum mesophyll
Increased transport; N. benthamiana mesophyll
Reduced transport; N. tabacum mesophyll
Reduced transport; N. benthamiana mesophyll
Narrowing of neck region; Zea mays roots
Chloride transport
Virus transport
Cell–cell electrical
conductance
Dye transport
Virus transport
Dye transport
Dye transport in MPexpressing tissue
TEM analysis
Reduced transport; Arabidopsis root tissues, Elodea Dye transport, GFP
canadensis leaf epidermis
transport
No effect on transport; N. tabacum mesophyll
Virus transport
NEM, myosin inhibitor
Myosin antibodies
Increased transport; T. virginiana stamen hairs
Dye transport
Reduced transport; Nitella translucens
Reduced transport; T. virginiana stamen hairs
11
C and 14C transport
Dye transport
Increased transport; Arabidopsis root tissues, E.
Dye transport, GFP
canadensis leaf epidermis
transport
Increased transport; Arabidopsis root epidermis, N. Dye transport
tabacum mesophyll
Increased transport; T. virginiana stamen hairs
Dye transport
Reduced transport; Arabidopsis root epidermis
Dye transport
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Radford and White
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RG White et al.
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(2001)
Ding et al. (1996)
Liu et al. (2005)
Ding et al. (1996)
Su et al. (2010)
Radford and White
(1998)
RG White et al.
(unpublished)
Kawakami et al.
(2004)
Radford and White
(2011)
Dale et al. (1983)
Radford and White
(2011)
RG White et al.
(unpublished)
Volkmann et al.
(2003)
Radford and White
(2011)
RG White et al.
(unpublished)
5256 | White and Barton
2I; Table 2; White et al., 1994), suggesting that actin
filaments may be structural components of PD, perhaps
concentrated within this region.
More recent experiments in which the actin cytoskeleton
was perturbed have produced contrasting results. For
example, cytochalasin D had no noticeable effect on the
structure of PD in Azolla pinnata or barley (Hordeum
vulgare) root cells (White et al., 1994); latrunculin B
treatment did not increase transport of Lucifer yellow CH
across an active barrier to cytoplasmic movement at the
base of N. tabacum leaf trichomes (Christensen et al., 2009);
and neither cytochalasin D nor latrunculin B treatments
increased the percentage of cells exhibiting intercellular
movement of a fluorescent 3 kDa dextran in Tradescantia
virginiana stamen hairs (Radford and White, 2011), Elodea
canadensis leaf cells, or Arabidopsis root cells (RG White,
MJ Talbot, JE Yarnold, JE Radford, unpublished results).
Variation is also seen within tissues; application of cytochalasin B reduced cell–cell electrical conductance when the
initial conductance was high, and increased conductance if
it was initially low, in submerged trichomes of Salvinia
auriculata (Krasavina et al., 2001). Since cell–cell transport
varies depending on the plant species, tissue, and stage of
development of each system examined (Jones, 1976; Itaya
et al., 1998; Roberts et al., 2001; reviewed in Roberts and
Oparka, 2003), it is perhaps not surprising that there are
also variations in regulatory mechanisms.
However, one problem with these and most other
cytoskeleton-modifying chemicals is that they are usually
applied to entire tissues and, even when microinjected,
diffuse readily across cell boundaries, so there may be some
overlap between effects on cell metabolism and physiology
and more specific effects on PD. To avoid such side effects,
it may be possible to perturb actin assembly in single cells
by microinjecting RNAi, since virus-induced silencing
(VIGS) of actin reduced cell–cell transport of Tobacco
mosaic virus (TMV) in N. benthamiana (Liu et al., 2005).
Constitutive silencing of actin using VIGS causes stunting,
similar to that seen in Arabidopsis plants with mutations in
actin genes (Gilliland et al., 1998), or after long-term
latrunculin B treatment (Baluška et al., 2001b), so tissue- or
cell-specific knock-down in actin expression could be very
enlightening.
Introduction of protein antibodies is a fairly specific way
to inhibit protein function, and injection of actin antibodies
would be a useful supplement to the chemical studies.
Interestingly, microinjecting profilin, an actin monomersequestering protein, into N. tabacum mesophyll cells increased intercellular transport of large fluorescent dextrans
(Ding et al., 1996). This opens up the possibility that
endogenous actin-associated proteins regulate transport
through PD via their actin-binding or actin-modifying
capabilities.
Relevant to these considerations is mounting evidence
that viruses and/or their related proteins associate with actin
filaments during infection of plant cells (reviewed by
Harries et al., 2010). A number of viruses bind to actin in
vitro (e.g. McLean et al., 1995), or produce components
(TMV replication complexes, Liu et al., 2005; Cauliflower
mosaic virus P6 inclusion bodies, Harries et al., 2009a; viral
Hsp70h, Prokhnevsky et al., 2005) that move along microfilaments to PD. Application of actin inhibitors disrupts
these processes and can also reduce the cell–cell spread of
TMV replication complexes (Kawakami et al., 2004),
although TMV can move from cell to cell in the absence of
actin filaments (Hofmann et al., 2009). Recently Su et al.
(2010) demonstrated that the movement proteins (MPs) of
both TMV and Cucumber mosaic virus (CMV) sever actin
filaments to increase the SEL of N. benthamiana PD. Both
of these viruses track along actin microfilaments to reach
PD, and Su et al. (2010) showed that the MPs of TMV or
CMV appeared to sever actin filaments only at PD. Thus,
viruses can either discriminate between cytoplasmic and PD
actin, or the severing function is only active when their MPs
are bound within PD. Presumably, actin severing enables
cell–cell movement of the viruses. It is possible that MPinduced actin severing triggers part of the plant defence
response against virus invasion, since defence responses are
initiated following actin depolymerization upon fungal
invasion (Kobayashi and Kobayashi, 2007).
Actin-binding proteins
It is puzzling that despite the large number of ABPs in
plants, few have been localized to PD. While this could be
partly due to the difficulty in preserving and immunolabelling
actin in plants, it is also possible that PD-specific actinassociated proteins may not be detected by currently available antibodies, or by protein isolation or fluorescent tagging
techniques, leading to the impression of absence. Nevertheless, it can be speculated that actin-depolymerizing factors
or proteins such as villins that sever actin filaments
(Khurana et al., 2010) could be important in regulating
intercellular traffic, if located at PD. One ABP localized to
pitfields is the actin-related protein Arp3 (van Gestel et al.,
2003). The Arp2/3 complex nucleates actin filaments (Deeks
and Hussey, 2005; Fiserová et al., 2006) when activated by
SCAR/WAVE (Zhang et al., 2008), and its presence in PD
may indicate a role for actin filament turnover in their
dynamic regulation. Since virus MPs are located at PD and
show actin-severing capacity only in this location, there may
be additional endogenous protein(s) with similar functions
at PD. Research into actin-binding proteins is at an early
stage, and further investigation may identify specific associations with PD, especially with recent discoveries of a range
of new proteins involved in actin nucleation (e.g. Campellone
and Welch, 2010).
Plant myosins
The motor protein, myosin, is the most prominent ABP
found in PD. Of the 24 recognized types of myosin found in
eukaryotes, the four myosins VIII and 13 myosins XI are
found in plants, with myosin XIII found only in Acetabularia (Mooseker and Foth, 2008). They are multicomponent
complexes identified by their heavy chains, which comprise
The cytoskeleton in plasmodesmata | 5257
an actin-binding head, hinged neck, and variable length,
cargo-binding tail domain. These heavy chains are usually
accompanied by regulatory calmodulin-like myosin light
chains. The myosin head binds actin strongly in the absence
of ATP (rigor state), whereas ATP releases myosin from the
actin filament. The neck undergoes an immediate conformational change, the recovery stroke, and upon binding to
actin several monomers further along the filament, the
bound ATP is immediately hydrolysed, enabling somewhat
tighter binding. Following the release of ADP, release of Pi
causes the now tightly bound myosin head to undergo
a conformational change, the power stroke, which moves its
cargo relative to the actin filament, and the cycle begins
again (see Spudich, 2001; Holmes, 2008).
Like actin, myosin was first localized to PD with
immuno-EM using polyclonal antibodies to animal myosins
(Fig. 2E, F; Table 1; Blackman and Overall, 1998; Radford
and White, 1998), which recognize highly conserved motifs
in the myosin head, as well as an antibody to the C-terminal
tail of plant myosin VIII (Reichelt et al., 1999). Immunofluorescence analyses with the same antibodies showed labelling of individual, multilobed PD (Blackman et al., 1998),
pitfields (Radford and White, 1998; Reichelt et al., 1999;
Baluška et al., 2001a, 2004) and occasionally fine strands
crossing young cell walls (Fig. 2G; Radford and White,
1998).
Apart from these earlier studies, there have been no
further analyses of myosin (or actin) in or near PD at the
EM level. There is also a lack of any analysis of changes in
the location of myosin or other proteins as PD change their
structure from an initially simple pore to more complex
synapses linking neighbouring cells. With the wider availability of plant myosin-specific antibodies, it would now be
useful to identify whether myosin is located more towards
the neck region or deeper in the cytoplasmic sleeve, and
confirm, if possible, co-localization with actin. Other large,
multicomponent structures which regulate traffic across
double-membrane boundaries, such as the nuclear pore
complex, undergo substantial changes in composition
during cell development (Doucet et al., 2010), and there are
distinct differences in pore protein isoform composition
between cell types (Meier and Brkljacic, 2010). It remains to
be elucidated whether specific myosins are components of
PD at different developmental stages, and whether they are
also restricted to certain cell or tissue types.
Fluorescence analysis of living cells has expanded myosin
analysis, and several groups have used GFP fusions to parts
of the myosin protein to localize myosin in both transient
and stably expressing Arabidopsis and Nicotiana lines. These
analyses show that none of the fluorescent protein-tagged
myosin XI tail regions localizes to PD (Reisen and Hanson,
2007; Walter and Holweg, 2008; Avisar et al., 2009;
Sattarzadeh et al., 2009), and it seems likely that no myosin
XI plays a direct role as an internal component of PD. One
of two known myosin XI isotypes is strongly localized to
the coalescing cell plate in tobacco BY2 cells and appears
critical for localization of the ER there, suggesting a key
role in bringing and holding the trapped ER in place
between cell plate vesicles, although it is not localized to
PD in mature cell walls (Yokota et al., 2009). Intriguingly,
activity of the tobacco myosin XI, and not myosin VIII,
appears essential for ER tubule formation (Yokota et al.,
2011); perhaps one role is tightening the desmotubule in
nascent PD. Of the Arabidopsis myosin VIII family, one
GFP fusion with the IQ-tail zone of ATM1 appears to
localize to sites of ER attachment as well as pitfields when
expressed in N. benthamiana leaves (Golomb et al., 2008).
Furthermore, overexpression of the tail regions, including
the IQ motif, of three orthologues of Arabidopsis myosin
VIII sequenced from N. benthamiana prevented PD localization of a closterovirus hsp70 homologue, a viral protein
essential for cell–cell virus movement, whereas overexpression of myosin XI tails had no effect (Avisar et al., 2008).
However, when transiently expressed as tail–GFP fusions in
N. tabacum, three of the four Arabidopsis myosin VIII
family were detected at the plasma membrane, with the
fourth located in the nucleus (Avisar et al., 2009).
The actin-binding myosin head may be required to locate
the myosin correctly, if located at PD, or indeed anywhere
else in the cell. For example, a class I myosin requires the
head domain and correct conformation of the tail domain
for localization to actin-enriched membrane projections in
animal cells (Komaba and Coluccio, 2010). Furthermore,
free GFP or GFP-tagged myosin can impair actin–myosin
interactions in highly excitable muscle cells (Agbulut et al.,
2006, 2007; Nishimura et al., 2006; Sekar et al., 2007) where
actomyosin interactions occur in highly structured tissues.
Although there is no evidence that GFP–protein fusions
cause similar problems in other cell types, it is possible that
in the tightly packed PD neck region, the GFP–actomyosin
interaction may affect localization of the GFP–myosin tail
fusions. As for analysis of the role of actin, local cell or
tissue silencing would be useful to confirm roles for specific
myosins in regulation of PD.
Tropomyosin
Of particular interest is the recent localization of a tropomyosin-like protein in PD, and identification of a putative
endogenous plant tropomyosin in Chara and Arabidopsis
(Fig. 2H; Faulkner et al., 2009). In animals and yeast,
tropomyosin forms head–tail dimers that bind along, and
stabilize, actin filament structure and regulate its activity
and interaction with other ABPs, including myosin
(Meagher and Fechheimer, 2003; Gunning et al., 2008).
Tropomyosin research in plants has lagged because tropomyosins are almost entirely a-helical proteins and BLAST
homology searches cannot identify specific tropomyosin
sequences (Pruyne, 2008; Gardiner et al., 2011). The
sequence similarities identified in plant genome databases
tend to be short and are often to coiled-coil regions of
diverse other proteins (Meagher and Fechheimer, 2003;
Staiger and Hussey, 2004; Gunning, 2008), so there is
residual doubt that plants contain a true tropomyosin
homologue. However, since a major function of tropomyosin is to modulate the interaction between the motor
5258 | White and Barton
domain of the myosin head with filamentous actin (Perry,
2001; Gunning, 2008; Gunning et al., 2008), it is a puzzle
how plant myosins function in the absence of actin-bound
tropomyosin (and associated proteins such as troponins)
which provide calcium-regulated access to the actin filament. A different approach is needed; for example, yeast
tropomyosin was only identified through biochemical purification (Liu and Bretscher, 1989; reviewed in Pruyne,
2008), which may yet be a pathway for identification of
plant tropomyosins. Recently, Gardiner et al. (2011)
identified two putative Arabidopsis tropomyosin homologues using an algorithm that predicts protein homology.
It will be of interest to see whether these localize along actin
filaments or to PD in vivo.
In PD, a tropomyosin-like protein could stabilize actin
against turnover by agents that normally lead to disassembly of the highly dynamic cortical actin cytoskeleton
(Wachsstock et al., 1994; Staiger et al., 2009), or regulate
actin–myosin interactions there. PD are quite resistant to
proteases and detergents (Tilney et al., 1991; Turner et al.,
1994), and additional cytoskeleton-binding proteins may
underlie this stability. Tropomyosin is not indispensable
for myosin ‘walking’ along actin filaments or for movement
of (non-tropomyosin) stabilized actin filaments on a bed
of anchored myosin heads, but, without tropomyosin,
the sliding velocity is much lower (Higashi-Fujime and
Nakamura, 2009). Perhaps tropomyosin in PD is a stabilizing agent, not an accelerant. In animal epithelial cells, actin
filaments in lamellipodia lack tropomyosin and are highly
dynamic, whereas the more stable filament bundles further
from the leading edges of cells, in the lamella, are stabilized
with tropomyosin (Gupton et al., 2005). Tropomyosin is
also concentrated along the very robust peripheral actin
filament bundles in Chara cells (Faulkner et al., 2009),
which are locations requiring stable actin bundles as well as
high myosin activity. Arp2/3 binds more readily to actin
filaments lacking tropomyosin (van Gestel et al., 2003;
Gupton et al., 2005), and is found at the PD neck, but more
sparsely (if at all) down the length of PD (van Gestel et al.,
2003), providing some indirect support for tropomyosinstabilized actin within PD but not more generally in the
cytoplasm.
Myosin function in plasmodesmata
The role of myosin in PD has been less well characterized,
and, as with actin, the effects of chemical inhibitors vary
with the tissue examined. Inhibitors of myosin function
generally arrest myosin either attached to actin in the rigor
conformation, or detached from actin and unable to hydrolyse ATP prior to reattachment or the power stroke
(Spudich, 2001; Holmes, 2008). If myosin inhibitors interfere with actomyosin-based activities, such as cytoplasmic streaming, they are generally assumed to inhibit all
other processes relying on actomyosin dynamics.
Two drugs commonly used to inhibit myosin function are
2,3-butanedione monoxime (BDM) and N-ethylmaleimide
(NEM). BDM blocks myosin ATPase activity, such that the
myosin head is bound loosely to actin, and NEM binds to
a conserved cysteine residue near the myosin head, altering
the ATP-binding site, which prevents ATP binding and
prevents myosin detachment from actin after the power
stroke (Kohama et al., 1987, Patel et al., 1998). Although
BDM may not affect actomyosin activity in some plant
(McCurdy, 1999; Forer and Fabian, 2005) or animal (e.g.
Coluccio, 2008) tissues, the consensus is that both inhibitors
are useful probes of myosin function in plants (Tominaga
et al., 2000; Molchan et al., 2002; Funaki et al., 2004; Forer
and Fabian, 2005).
BDM induced constrictions in the neck region of Zea
mays and Hordeum vulgare PD (Fig. 2J; Radford and
White, 1998); it also reduced intercellular transport of
TMV replication complexes in tobacco mesophyll tissue
(Kawakami et al., 2004), and transport of fluorescent
tracers in Arabidopsis root cells (RG White, MJ Talbot, JE
Yarnold, JE Radford, unpublished results). The implication
is that constriction of the PD neck results in reduced
transport, yet myosin is found all along the cytoplasmic
sleeve, not solely at the neck, suggesting that myosin is only
active in this specific zone in PD. Furthermore, in animal
tissues, BDM detaches the myosin II (Packer et al., 1988;
Higuchi and Takemori, 1989; Bagni et al., 1992; Herrmann
et al., 1992; Steele and Smith, 1993; Backx et al., 1994;
Hajjar et al., 1994; Zhao et al., 1995) and V (Cramer and
Mitchison, 1995) heads from actin, suggesting that disengagement of actin and myosin, each putatively on different
PD membranes, causes PD closure. PD also open when
ATP levels are reduced (Tucker, 1993; Cleland et al., 1994;
Wright and Oparka, 1997) and close in the presence of
additional ATP (Radford and White, 2011), which would
produce either firmly attached or detached myosin, respectively, supporting the idea that detached myosin
correlates with closed PD. If the default status of PD is
open, and reduced SEL or complete closure requires energy,
then perhaps myosin forms ‘struts’ between the ER and
plasma membrane (one of which is lined with actin), which
are no longer active if myosin is detached.
The PD-located plant myosin VIII is more specifically
inhibited by injection of antibodies to its unique coiled-coil
tail region, which increased transport between Arabidopsis
root epidermis or tobacco mesophyll cells (Volkmann et al.,
2003). Similarly, injection of polyclonal anti-myosin antibodies, in this case presumably interfering with a conserved
site on the myosin head, also increased cell–cell transport
between T. virginiana stamen hairs (Radford and White,
2011). However, when the same antibody was injected into
Arabidopsis root epidermal cells, PD transport was reduced
(RG White, MJ Talbot, JE Yarnold, JE Radford, unpublished results). Furthermore, BDM treatment increased
and NEM reduced cell–cell transport in T. virginiana
(Radford and White, 2011), as seen previously in Nitella
(Dale et al., 1983), but opposite to Z. mays or H. vulgare
(Radford and White, 1998), again suggesting that interfering with the head and tail regions can produce different
results, and that cell- and tissue-specific factors regulate PD.
It would be profitable to screen other small molecules for
The cytoskeleton in plasmodesmata | 5259
their potential to inhibit specific plant myosins, as has been
done for skeletal myosin II (Cheung et al., 2002).
Myosin regulation of plasmodesmatal actin
A final consideration is that myosin inhibitors may indirectly cause changes in cell–cell transport by affecting
other aspects of cell physiology. For example, both NEM
(Karlsson and Lindberg, 1985; Menzel and Elsner-Menzel,
1989; Liebe and Quader, 1994; Quader and Liebe, 1995)
and BDM (Šamaj et al., 2000; Holweg et al., 2003) have
been shown to cause fragmentation, re-arrangement, and
bundling of actin, although 30 mM BDM, while inhibiting
cytoplasmic streaming, did not affect actin filaments in
T. virginiana stamen hairs (Molchan et al., 2002; Radford
and White, 2011). When muscle myosin II is bound to
actin the myosin head interacts strongly with regions on
two adjacent actin monomers (Milligan, 1996). This strong
interaction can stabilize the actin filament and even induce
the polymerization of actin monomers into filaments
(Carlier et al., 1994). Thus disruption of the actin–myosin
interaction with chemical treatments or antibody injection
could disrupt the actin filaments within PD (White et al.,
1994), which could then affect cell–cell transport (Ding
et al., 1996), at least in expanding Arabidopsis root cells.
Yarrow et al. (2003) found that BDM blocked actin
incorporation into sites of rapid actin turnover in cultured
animal cells, by delocalizing actin-polymerizing proteins at
the cell membrane. Myosin I proteins interact directly with
the Arp2/3 complex in yeast, Dictyostelium, and other
species to promote actin polymerization (Ridley, 2000),
and this interaction is disrupted by BDM (Yarrow et al.,
2003). If specific sites of rapid actin turnover in plant cells
also depend upon a suite of plasma membrane-associated,
BDM-sensitive polymerization factors, and such active
sites include PD, then BDM may have a secondary effect
on PD through this mechanism.
Related cytoskeletal proteins; centrin, calmodulin,
tubulin
Antibodies to the contractile protein, centrin, label developing cell plates (Del Vecchio et al., 1997; Blackman
et al., 1999; Harper et al., 2000) and the neck region of PD
(Fig. 2K; Blackman et al., 1999), leading to speculation that
a centrin-like protein may link the ER and plasma
membrane. Centrins decorate repeated sites on centrinbinding proteins, and initiate twisting and contraction of
these long proteins when they bind calcium and change
their conformation (Salisbury, 2004; Gogendeau et al.,
2007). Centrins are also components of the nuclear pore
complex in yeast (Rout et al., 2000) and vertebrates
(Resendes et al., 2008), where they play an essential role in
export of mRNA and protein. Nuclear pores are useful
comparisons with PD, as they have similar features,
particularly the regulated transport of macromolecules
through a pore which traverses a double membrane.
Perhaps, similar to the actin-severing role of some virus
MPs, centrin may act in the regulation and turnover of PD
proteins at the neck region, similar to its role in yeast (Chen
and Madura, 2008).
Since changes in calcium levels clearly regulate PD
permeability, it is surprising that other calcium-binding
regulatory proteins, such as calmodulin, are not localized
there (Blackman and Overall, 1998), and this absence
suggests that the calmodulin-like myosin light chains may
not regulate rapid myosin dynamics at PD. Antibodies to
tubulin label PD rather sparsely, only slightly more than in
the cytoplasm (Blackman and Overall, 1998), and treatment
with a microtubule inhibitor, colchicine, had little effect
on cell–cell conductivity in Trianea bogotensis roots or
S. auriculata trichomes (Krasavina et al., 2001). Microtubules can deposit cargo, especially viruses, at PD (Harries
et al., 2009b), but they do not seem to continue into the
cytoplasmic channel. Other viruses traffic along actin
filaments within cells (Harries et al., 2009b, 2010). Perhaps
the common element is myosin, since some myosins, for
example myosin V, can bind to either actin filaments or
microtubules, and while the true motor function operates
only with actin, this myosin can slide for some distance
along microtubules (see Mooseker and Foth, 2008). Further
investigation may reveal a similar dual function in one or
more of the plant myosins XI or VIII, which share features
in common with myosin V (Yamamoto, 2008).
Cytoskeleton organization in and regulation of
plasmodesmata
Ultrastructural elements seen in and around PD, revealed
by TEM staining and thought to be individual proteins,
suggest a spiral arrangement of single filament(s) around
the desmotubule (White et al., 1994; Overall and Blackman,
1996; Blackman et al., 1999; Overall, 1999; Benitez-Alfonso,
2010), although supporting evidence is sparse. In comparison, it was thought that helically arranged elements
decorating microtubules in developing photoreceptor cells
were actin derived (Obata and Usukura, 1992), but this is
probably centrin decoration (Wolfrum, 1995; Trojan et al.,
2008), with the actin filaments playing a role adjacent to the
centriole-derived microtubules (Obata and Usukura, 1992;
Trojan et al., 2008). In any event, it is difficult to envisage
how myosin could traffic along actin filaments in the
confined space of the cytoplasmic sleeve, especially at the
neck region.
Most plant actin is highly dynamic (Staiger et al., 2009),
and actin filaments are probably not stiff enough to shape
the desmotubule by themselves (Kole et al., 2004, 2005;
Gupton et al., 2005). In bacteria, MreB and Mbl are two of
several bacterial homologues of actin which form spiral
structures on the inner surface of the bacterial inner
membrane, and are essential for maintenance of cell shape
and assembly of the bacterial peptidoglycan matrix (Gupton
et al., 2005; Divakaruni et al., 2007; White et al., 2010).
MreB filaments form thick, spiral bundles that require
auxiliary cross-linking factors to reinforce their stiff cortexlike structure and to anchor these assemblies to the inner
5260 | White and Barton
membrane (Esue et al., 2005; White et al., 2010). If plant
actin is similarly stabilized and associated with a membrane,
association with the plasma membrane rather than the
desmotubule would produce a looser actin spiral, similar to
those seen in bacteria.
However, instead of actin monomers or filaments, the
particles associated with the desmotubule may be myosin
heads, arranged in a spiral as they are in muscle, with the
heads pointing outwards to actin lining the plasma membrane. Myosin heads may interact with plasma membranebound actin only at the PD neck, whereas, through the
cytoplasmic sleeve, myosin may interact with actin or actinlike proteins on certain specific types of cargo. In any event,
further high resolution analyses of PD ultrastructure are
clearly needed to reveal the position of actin filaments and
myosin in PD, especially at the neck where cargo transfers
from the cytoplasmic to the PD cytoskeleton.
Conclusions and prospects
Exploration of the role of cytoskeletal proteins in PD
structure and regulation has only just begun, and future
progress will depend on targeting these proteins in specific
cell types, rather than using general inhibitors. For example,
injection of RNAi to actin (cf. VIGS; Liu et al., 2005) or
myosin could be very informative, together with celltargeted inducible expression of RNAi, and complementation analyses (e.g. Abu-Abied et al., 2006). Perhaps then we
will be able to answer the questions raised here and shed
a brighter and clearer light on the intercellular cytoskeleton
in plants.
Acknowledgements
We thank Drs Janine Radford and Monica Schoenwaelder
for providing the unpublished micrographs in Fig. 1.
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