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Recent Advances in Understanding Plant Myosin Function: Life in the Fast Lane
Molecular Plant
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Volume 4
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Number 5
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Pages 805–812
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September 2011
REVIEW ARTICLE
Recent Advances in Understanding Plant Myosin Function: Life in the Fast Lane Imogen Sparkes1 School of Life Sciences, Oxford Brookes University, Gipsy Lane, Oxford, OX3 0BP, UK
Key words:
Myosin; actin; endoplasmic reticulum; organelle movement; plant development.
INTRODUCTION The advent of live cell imaging and fluorescent probes has confirmed the highly dynamic intracellular nature of plant cells. Whilst plants maybe sessile, intracellularly they certainly live life in the fast lane, with cytoplasmic streaming reaching rates of 100 lm s 1 in Chara (Shimmen and Yokota, 2004). The general phenomenon of streaming was attributed to bulk flow of organelles resulting in a hydrodynamic flow of the cytoplasm (Esseling-Ozdoba et al., 2008). However, quantification of these events has identified what appear to be directed types of movement displayed by several classes of organelle. Organelles do not simply move at the same speed in the same direction; they stop and go, move bi-directionally, and can reach speeds of up to 7–8 lm s 1. It is still a mystery as to why and how plant organelles display such seemingly random movements; however, insights gleaned from organelle positioning and movement in response to biotic and abiotic factors indicate that they do have a functional role. For example, repositioning of the nucleus and chloroplasts in response to light (Wada et al., 2003; Iwabuchi et al., 2007) increased peroxisome movement in response to cadmium (Rodriguez-Serrano et al., 2009), and drastic intracellular rearrangements are seen in response to pathogen attack and mechanical stimuli (Takemoto et al., 2003; Lipka et al., 2005; Hardham et al., 2008). Therefore, intracellular dynamics is analogous to a subcellular roadmap with routes of fast movement on intracellular ‘highways’ and more random meandering movements on the ‘byways’, all of which are implicated in delivery, collection, and shuttling of cargo, metabolites, and signaling intermediates for cellular homeostasis.
Plant myosins are classified into two groups: class XI and class VIII. Class XI is similar to mammalian class V owing to comparable protein domains including the dilute domain in the carboxy terminus, and they also share a common function in organelle movement. Plant myosins contain an amino terminal motor head domain responsible for actin binding and ATPase activity, a regulatory neck region containing IQ motifs, and a carboxy terminal implicated in cargo binding (Figure 1). Genome sequencing has shown that there are 9, 3, 13, and 12 class XI myosins and 2, 5, 4, and 2 class VIII myosins in brachypodium, moss, Arabidopsis, and rice, respectively (Reddy and Day, 2001; Jiang and Ramachandran, 2004; Vidali et al., 2010; Peremyslov et al., 2011). Algae also appear to encode for both class XI and class VIII myosins (Odronitz and Kollmar, 2007; Peremyslov et al., 2011). Recently, there has been an explosion in data relating to plant myosin location (see Table 1), function, and mechanistics (see reviews Shimmen and Yokota, 2004; Li and Nebenfu¨hr, 2008b; Sparkes, 2010, and references therein). Myosins have been implicated in endocytosis (Baluska et al., 2004; Golomb et al., 2008; Sattarzadeh et al., 2008), organelle movement (see below), cytoplasmic streaming and stiffness (Shimmen and Yokota, 2004; Esseling-Ozdoba et al., 2008; van der Honing et al., 2010),
1 To whom correspondence should be addressed. E-mail isparkes@brookes. ac.uk, tel. +44 (0)1865 483639, fax +44 (0)1865 483955.
ª The Author 2011. Published by the Molecular Plant Shanghai Editorial Office in association with Oxford University Press on behalf of CSPB and IPPE, SIBS, CAS. doi: 10.1093/mp/ssr063, Advance Access publication 19 July 2011 Received 4 May 2011; accepted 20 June 2011
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ABSTRACT Plant myosins are required for organelle movement, and a role in actin organization has recently come to light. Myosin mutants display several gross morphological phenotypes, the most severe being dwarfism and reduced fecundity, and there is a correlation between reduced organelle movement and morphological defects. This review aims to discuss recent findings in plants relating to the role of myosins in actin dynamics, development, and organelle movement, more specifically the endoplasmic reticulum (ER). One overarching theme is that there still appear to be more questions than answers relating to plant myosin function and regulation.
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cytokinesis (Collings et al., 2003; Nebenfu¨hr, 2007), plasmodesmata function (Radford and White, 1998, 2011), and viral movement (Avisar et al., 2008a; Harries et al., 2009). Here, this short review will focus on recent discoveries relating to the potential role of myosin in actin organization, organelle movement, more specifically ER remodeling, and plant development. Stromule movement and the role of myosin XI-F are covered in the accompanying review by Kong and Wada (2011) in this special issue.
PLANT MYOSINS AND ACTIN ORGANIZATION
Figure 1. Schematic Representation of Myosin Structure. Schematic representation of a class XI myosin highlighting the amino terminal head domain that binds ATP and actin (blue), the neck region containing six calmodulin binding IQ motifs (green) and coiled coil regions (red), and a carboxy globular tail containing a Dilute domain (black). The globular tail is composed of alpha helices proposed to form two interacting globular tail subdomains (GT1, GT2, see Table 1; Li and Nebenfu¨hr, 2007). Note, this schematic is based on myosin XI-K.
Table 1. Subcellular Location of Arabidopsis Myosin Fusions. Organelle
Myosin
Reference
Peroxisome
FP-XI-2 (1053–1505 aa)a
Reisen and Hanson, 2007
FP-GT1 OR GT2 XI-1 (1099–1301 aa, 1283–1520 aa)b
Li and Nebenfu¨hr, 2007
FP-GT2 XI-2 (1279–1505 aa)b
Li and Nebenfu¨hr, 2007
GT1+GT2; XI-1 (1099–1301 aa, 1283–1520 aa); XI-2 (1099–1297 aa, 1279–1505 aa); XI-I (1115–1290 aa, 1280–1522 aa); XI-K (1124–1325 aa, 1307–1545 aa)c
Li and Nebenfu¨hr, 2007
CCGT+CCGT XI-1 (867–1520 aa)c
Li and Nebenfu¨hr, 2008a
Golgi
FP-GT2 XI-1 (1283–1520 aa)*b
Li and Nebenfu¨hr, 2007
Chloroplast
FP-XI-F (1272–1313 aa)a
Sattarzadeh et al., 2009
Endosomes
FP-ATM1IQtail (838–1167 aa)a
Golomb et al., 2008
a
Golomb et al., 2008
Plasmodesmata
FP-ATM1IQtail (838–1167 aa)
Nuclear envelope
FP-XI-I IQtail/tail (752–1523 aa, 881–1523 aa)a
Avisar et al., 2009
Nucleolus
FP-VIIIB IQtail/tail (813–1127 aa, 905–1127 aa); FP-ATM2 tail (774–1031 aa)a
Avisar et al., 2009
Plasma membrane
FP-ATM1IQtail/tail (838–1167 aa, 943–1167 aa); FP-ATM2 IQtail/tail (688–1031 aa, 774–1031 aa); FP-VIIIA IQtail/tail (820–1154 aa, 925–1154 aa); FP-VIIIB IQtail/tail (813–1127 aa, 905–1127 aa)a
Avisar et al., 2009
Location data are based on transient expression of fluorescent myosin fusions either in tobacco (a) or Arabidopsis (b) leaf epidermal cells. Bimolecular fluorescence complementation assays whereby two regions of the same myosin highlighted in the table were transiently co-expressed in Arabidopsis leaf epidermal cells along with a peroxisome marker highlighted location (c). * 20% of cells showed Golgi rather than peroxisome location. The location of native XI-2 to peroxisomes has been confirmed by immunofluorescence using an anti-XI-2 serum (Hashimoto et al., 2005). Abbreviations denote the following: FP, fluorescent protein; CC, coiled coil domain; GT1/GT2, globular tail region 1/2; aa, amino acid.
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Association of specific plant myosins with actin have been documented; myosin VIII-1 in tobacco BY-2 cells (Yokota et al., 2011) and Arabidopsis XI-2 (also known as MYA2) (Hashimoto et al., 2005) co-sedimented with actin in an ATP-dependent manner, and a XI-2 (MYA2) head fusion construct localized to actin filaments (Walter and Holweg, 2008). In vitro motility assays have shown that XI-1 (also known as MYA1) can
translocate along actin at up to 1.8 lm s 1 (Hachikubo et al., 2007), and a 175-kDa tobacco myosin (75% sequence similarity to XI-2) was shown to process along actin with 35-nm step size at a rate of 7 lm s 1 (Tominaga et al., 2003). Whilst association between myosin and actin implies a role relating to myosincargo motility, it does not preclude roles in regulating actin organization and/or dynamics. Through RNA silencing of two class XI myosins in Moss (XIa, XIb), it was shown that actin organization but not dynamics were altered, resulting in disorganized actin arrays rather than longitudinal filaments (Vidali et al., 2010). In higher plants, Ueda et al. (2010) reported that Arabidopsis xi-k xi-2 double and xik xi-1 xi-2 triple myosin mutants displayed altered actin organization. Actin bundles were randomly orientated and there was a reduction in the number of parallel actin bundles in cotyledonary petiole epidermal cells. A similar distribution of actin bundles was rarely seen in control and single mutants (xi-k, xi-1, xi-2), thus indicating that all three myosins are required collectively to maintain actin bundle organization in the cell types under study. In a further more comprehensive study of triple (xi-k xi-1 xi-2) and quadruple (xi-k xi-1 xi-2 xi-i) myosin mutants on plant development (see below) and actin organization, Peremyslov et al. (2010) documented a similar affect in the leaf midvein cells of both mutants, with actin bundles being orientated up to 90 to the cell axis. Additionally, defects in root hair actin organization were also evident in the triple and quadruple mutants where actin bundles were no longer excluded from the root hair tip. However, observations of leaf pavement and root epidermal cells did not indicate a discernable difference in actin microfilament architecture (Peremyslov et al.,
Sparkes
PLANT MYOSINS AND ORGANELLE MOVEMENT: CHARACTERIZING ER NETWORK REMODELING The majority of organelle movement in plants is actin-dependent and recent advances have begun to assign specific myosin activity to this process. Previously, a role in organelle dynamics was inferred through chemical inhibition studies using drugs that affect actin dynamics (latrunculin b or cytochalasin) or myosin activity (BDM). Thus, myosins have been attributed to the movement of the ER (Quader et al., 1987; Liebe and Menzel, 1995; Sparkes et al., 2009a), Golgi (Boevink et al., 1998; Nebenfu¨hr et al., 1999), mitochondria (Van Gestel et al., 2002; Zheng et al., 2009), peroxisomes (Jedd and Chua, 2002; Mano et al., 2002; Collings et al., 2003), plastids (Wada et al., 2003; Paves and Truve, 2007), and the vacuole (Higaki et al., 2006). A more targeted approach through expression of dominant negative myosin mutants, RNAi, and T-DNA knockouts has revealed several members of the class XI myosins are involved in organelle movement: XI-1, XI-2, XI-C, XI-E, XI-I, and XI-K (see Sparkes, 2010, review and references therein). Expression of dominant negative fusions that lack the motor domain are predicted to either bind to cargo or titrate out accessory factors required for endogenous myosin recruitment, therefore perturbing myosin function. This approach has produced variable results. Truncations can bind to organelles but do not stop their movement (see Table 1), or they are diffuse in the cytoplasm or label motile puncta and stop organelle movement (XI-1, XI-2, XI-C, XI-E, XI-I, XI-K). Studies of myosin tail domain binding to cargo indicate that cargo binding and dimerization are cooperative processes (see Li and Nebenfu¨hr, 2008a, 2008b, and references therein). Inferences from all of these experiments indicate that myosin folding and alterations in conformation are complex and could explain why expression of small myosin fragments have variable affects on cargo binding and movement (see Table 1). Table 1 is included for reference to highlight the specific regions of myosins that have been expressed as fluorescent protein fusions and been assigned to a specific subcellular location. Note, however, that none of the fusions were reported to affect the movement of
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the organelle to which they were bound except for ATM1 IQ tail, which drastically reduced the movement of Ara6-labeled endosomes, yet the movement of FYVE-containing endosomes was only perturbed when a large excess of ATM1IQtail surrounded these structures (Golomb et al., 2008). A detailed comprehensive study of serial deletions and point mutations of conserved residues within myosin tail domains is required to not only address location, but also quantify effects on organelle movement. Future studies with full-length myosin fusions in combination with a genetic approach should yield a more directed experimental approach to understand the functional role and mechanistics of individual myosins. Using the experimental approaches described above, specific myosins have been implicated in having a global affect on the movement of several classes of organelle including mitochondria, peroxisomes, and Golgi (Avisar et al., 2008b; Peremyslov et al., 2008; Prokhnevsky et al., 2008; Sparkes et al., 2008; Avisar et al., 2009; Peremyslov et al., 2010). Roles in stromule (Natesan et al., 2009; Sattarzadeh et al., 2009) and ER dynamics (Sparkes et al., 2009a; Peremyslov et al., 2010; Ueda et al., 2010; Yokota et al., 2011) have recently been added to the list. The ER is a complex network of polygonal tubules interconnected by three-way junctions and anchor points that are proposed to tether it to the plasma membrane. Whilst the majority of the network remodels through tubule growth and shrinkage, tubule sliding to form polygonal rings, and a seemingly ready conversion between tubular and cisternal forms, there appears to be a fast streaming component beneath the more static peripheral network (Figure 2 and Supplemental Movie 1). Cells with a high secretory load display a more cisternal form of ER (Stephenson and Hawes, 1986; Ridge et al., 1999), and those undergoing pathogen attack develop cisternal ER around the site of invasion that can be mimicked by mechanical stimulus (Takemoto et al., 2003; Hardham et al., 2008). Therefore, there appears to be a functional role for ER remodeling. Also, the shear forces generated during ER streaming have been suggested to play a role in cytoplasmic streaming (Kachar and Reese, 1988; Ueda et al., 2010). Immunoblotting of a sucrose density centrifuged microsomal fraction indicated that myosin XI-K co-sedimented with the ER fraction (Ueda et al., 2010). An antibody against a 175kDa tobacco myosin with 75% sequence similarity to XI-2 appeared to label the ER in BY-2 suspension cell culture cells and also co-sedimented with the ER fraction (Yokota et al., 2009). None of the fluorescent fusions to Arabidopsis myosin truncations to date has specifically labeled the ER. However, there did appear to be a correlation between the plasma membrane puncta labeled by ATM1-IQtail fusion and the underlying ER network (Golomb et al., 2008). It remains to be seen whether or not ATM1 plays a role in ER network formation. Two recent studies used different approaches to quantify global ER movement in vivo (Sparkes et al., 2009a; Ueda et al., 2010). Persistency mapping, an analytical approach to characterizing static components (nodes, tubules, cisternae)
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2010). Similarly, expression of tail and IQ tail fusions of the entire Arabidopsis myosin gene family did not exhibit gross rearrangement of actin architecture in tobacco leaf epidermal cells (Sparkes et al., 2008; Avisar et al., 2009). Therefore, it appears that, whilst certain myosins (XI-K, XI-1, XI-2) affect actin organization, this appears to be a tissue-specific phenomenon and it is unclear how the myosin could mediate such an affect. Through the use of BDM, a myosin ATPase inhibitor, Staiger et al. (2009) have reported that myosins have a direct or indirect affect on actin filament buckling and straightening. One proposal could be that myosins provide the tensile force to effectively pull an actin filament straight. Future studies are therefore required to assess whether myosins could effectively tether and provide tensile force on actin filaments, with XI-K, XI-1, and XI-2 being likely candidates for initial studies.
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PLANT MYOSINS AND DEVELOPMENT
Figure 2. Persistency Mapping of Cortical ER in Tobacco Leaf Epidermal Cells. Persistency maps of the cortical ER were generated where the first frame (A) and last frame (B) of the movie are shown. Persistent nodes/cisternae (C) and tubules (D) are highlighted. Arrow indicates the direction of streaming that is observed in Supplemental Movie 1, panels a and b. Scale bar = 10 lm. Images taken from Sparkes et al. (2009a). Copyright American Society of Plant Biologists, www.plantcell.org.
in the ER network, highlighted persistent static nodes of ER in tobacco leaf epidermal cells proposed to act as growth or anchor points. A potential role in anchoring the ER perhaps through connectivity to the plasma membrane has also been inferred from optical tweezer studies (Sparkes et al., 2009c). Based on latrunculin b treatment or co-expression of the tail domain of Arabidopsis XI-K, it was evident that both actin dynamics and XI-K are required to maintain ER remodeling (Sparkes et al., 2009a). A separate study characterizing ER dynamics in the cotyledonary petioles of Arabidopsis single, double, and triple T-DNA insertional myosin mutants also indicated that XI-K plays a role, with XI-1/2 exerting minor affects on ER streaming (Ueda et al., 2010). An in vitro reconstitution assay has been used to determine the role of actin and myosin in generating the tubular ER network (Yokota et al., 2011). Through density centrifugation, a microsome fraction containing cytosol was separated and the effects on tubular-like network regeneration in the presence and absence of actin, energy (ATP/GTP), a 175-kDa class XI myosin, and a class VIII myosin was observed. Results indicated that formation of a network-like structure reminiscent of the ER required all of the latter components except the class VIII myosin. GTPase activity could be attributed to a requirement for Rabs or perhaps a role of RHD3 in the experimental system as the rhd3-1 mutant has unbranched ER tubules (Zheng et al., 2004).
Plant growth and development are complex coordinated processes affected by extracellular signals such as light and temperature. The majority of plant cells undergo anisotrophic growth (i.e. expansion along a preferred axis). At the cellular level, cytoskeletal elements are important in controlling expansion, elongation, and regulated shape changes through deposition of microtubules and actin microfilament patches (Yang, 2008). This coordinated process allows the targeting and trafficking of components required for growth such as the cellulose synthase complex and mediates endo/exocytotic events important for tip growth. Therefore, disruption in this finely tuned balance, and the molecular motors that traverse the cytoskeletal networks, can have drastic changes in the final body plan of the plant. There is a distinct correlation between plant growth defects and the role of myosins. The exact biological role(s) of, for example, secretion and endocytosis, resulting in these developmental defects, is unclear at this time, but a role in actin remodeling and organelle movement has been shown (see earlier). Root hairs undergo polarized tip growth that is due to the addition of membrane and wall material at the tip of the cell. A small group of Arabidopsis single myosin mutants (xik and xi2) display shorter root hairs (Ojangu et al., 2007; Peremyslov et al., 2008). Double mutants of paralogous myosin pairs indicated an additive affect for XI-K, XI-2, and XI-B on root hair length (Prokhnevsky et al., 2008). Defects in moss protonemal tip growth are also evident in a double myosin mutant xia xib (Vidali et al., 2010). In quadruple Arabidopsis mutants, defects in cells undergoing expansion through diffuse growth were observed and appeared to correlate with cell size; moderate reduction in mesophyll cell size compared to an increased reduction in the larger midvein cells (Peremyslov et al., 2010). Stunted growth (reduced rosette diameter, plant height, root length) and reduced flowering time were observed in triple and quadruple mutants, with xi-k xi-1 xi-2 xi-i displaying the severest phenotype (Peremyslov et al., 2010). Golgi and peroxisome movement in this mutant was reduced to ‘residual wobbling’
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Based on the experimental approaches discussed above, it has been suggested that the acto-myosin system is required for ER tubule growth and shrinkage through actin polymerization and myosin processivity. Movement effectively pulls out ‘excess’ membrane from cisternal regions to form the tubular network and sliding of tubules forms closed polygonal rings (Sparkes et al., 2009a). In the absence of cytoskeletal elements, the ER network is stabilized presumably through anchor point connections to the plasma membrane and scaffolding/stabilizing elements such as reticulons (see Sparkes et al., 2009b, and references therein). It will be interesting to see whether and how several myosins coordinate this process and whether the coordination is through specific myosin–ER interactions, or regulation of myosin interaction through homo and heterodimerization of myosin complexes.
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Figure 3. Expression Patterns of Arabidopsis Myosin Genes. Public affymetrix DNA microarray data were assessed using Genevestigator (www.genevesigator.com; Hruz et al., 2008) to generate a heat map of Arabidopsis myosin gene expression in various tissues. Predicted expression levels are indicated in the top left-hand corner inset, where dark blue refers to the highest predicted level of expression.
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attributed to Brownian movement or a ‘tug-of-war’ between remaining myosins failing to move either class of organelle in a processive manner (Peremyslov et al., 2010). The double myosin mutant xi-k xi-1 also displayed reduced fecundity and was stunted (Prokhnevsky et al., 2008). Therefore, it appears that reduced plant stature is in part due to reduced cell size, but, as was documented for leaf pavement and mesophyll cells, also a reduction in cell number (Peremyslov et al., 2010). Given the large number of myosin genes in plants, and the apparent lack of phenotypic defects in the majority of single mutants, it has been proposed that there is functional redundancy within the family. However, whilst the single myosin mutant studies monitored and quantified affects in certain tissues (root hair length, for example), a comprehensive study of all tissues and stages of development has not been provided. Therefore, potential defects in tissues shown by microarray analysis to express these genes (Figure 3) in wild-type plants should also be monitored. An additional hypothesis is that certain myosins have specific roles under perturbed noncontrolled growth conditions such as altered light and temperature. Peremyslov et al. (2011) analyzed microarray data and have characterized the expression profile of Arabidopsis and rice myosin genes. Interestingly, the general trend is that expression follows photocycles and thermal cycles, and three appear to display a pattern indicative of circadian regulation (rhythmic expression irrespective of light/dark cycles; XI-B, XI-H, VIIIB). Photoperiod regulation was previously shown for myosin XI-B in rice (Jiang et al., 2007). Based on these expression profiles, it will be interesting to see whether single and multiple myosin mutants display aberrant morphologies under altered growth conditions, and also in response to abiotic and biotic factors. Another interesting level of regulation could be through dimerization with headless domain variants that, in effect, could mimic the experimental response seen through overexpression of myosin tail fusions. Such a transcript for XI-K was revealed by Peremyslov et al. (2011) and expression was shown through a GUS reporter trap. Further bioinformatic mining of the genome combined with experimentation will reveal whether similar headless variants are present and represent true transcripts rather than genetic relics, and whether they play a role in myosin regulation.
actin and microtubule-dependent in elongating internodal cells (Foissner et al., 2009), and a role for kinesin 13-a in Golgi positioning has been suggested (Lu et al., 2005).
FUTURE DIRECTIONS
Foissner, I., Menzel, D., and Wasteneys, G.O. (2009). Microtubuledependent motility and orientation of the cortical endoplasmic reticulum in elongating characean internodal cells. Cell Motility Cytoskeleton. 66, 142–155.
Supplementary Data are available at Molecular Plant Online.
FUNDING Some of the work reported here was supported by Oxford Brookes University, the BBSRC, and the Leverhulme Trust.
ACKNOWLEDGMENTS I would like to thank Chris Hawes for critical reading of the manuscript and Larry Griffing for kindly agreeing to use his movie published in Sparkes et al. (2009a). No conflict of interest declared.
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Kong, S.-G., and Wada, M. (2011). New insights into dynamic actinbased chloroplast photorelocation movement. Mol Plant. 4, doi: 10.1093/mp/SSR061.
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Li, J., and Nebenfu¨hr, A. (2007). Organelle targeting of myosin XI is mediated by two globular tail domains with separate cargo binding sites. J. Biol. Chem. 282, 20593–20602.
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Li, J.F., and Nebenfu¨hr, A. (2008a). Inter-dependence of dimerisation and organelle binding in myosin XI. Plant J. 55, 478–490.
Radford, J.E., and White, R.G. (1998). Localisation of a myosin-like protein to plasmodesmata. Plant J. 14, 743–750.
Li, J.F., and Nebenfu¨hr, A. (2008b). The tail that wags the dog: the globular tail domain defines the function of myosin V/XI. Traffic. 9, 290–298.
Radford, J.E., and White, R.G. (2011). Inhibitors of myosin, but not actin, alter transport through Tradescantia plasmodesmata. Protoplasma. 248, 205–216.
Liebe, S., and Menzel, D. (1995). Actomyosin-based motility of endoplasmic reticulum and chloroplasts in Vallisneria mesophyll cells. Biol. Cell. 85, 207–222.
Reddy, A.S.N., and Day, I.S. (2001). Analysis of the myosins encoded in the recently completed Arabidopsis thaliana genome sequence. Genome Biol. 2, 1–18.
Lipka, V., et al. (2005). Pre- and postinvasion defences both contribute to non host resistance in Arabidopsis. Science. 310, 1180–1183.
Reisen, D., and Hanson, M.R. (2007). Association of six YFP-myosin XI-tail fusions with mobile plant cell organelles. BMC Plant Biol. 7, doi: 10.1186/1471-2229-1187-1186.
Lu, L., Lee, Y.R.J., Pan, R., Maloof, J.N., and Liu, B. (2005). An internal motor kinesin is associated with the Golgi apparatus and plays a role in trichome morphogenesis in Arabidopsis. Mol. Biol. Cell. 16, 811–823. Mano, S., Nakamori, C., Hayashi, M., Kato, A., Kondo, M., and Nishimura, M. (2002). Distribution and characterization of peroxisomes in Arabidopsis by visualization with GFP: Dynamic mor-
Ridge, R.W., Uozumi, Y., Plazinski, J., Hurley, U.A., and Williamson, R.E. (1999). Developmental transitions and dynamics of the cortical ER of Arabidopsis cells seen with green fluorescent protein. Plant Cell Physiol. 40, 1253–1261. Rodriguez-Serrano, M., Romero-Puertas, M.C., Sparkes, I.A., Hawes, C., del Rio, L.A., and Sandalio, L.M. (2009). Peroxisome
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Hruz, T., Laule, O., Szabo, G., Wessendorp, F., Bleuler, S., Oertle, L., Widmayer, P., Gruissem, W., and Zimmerman, P. (2008). Genevestigator V3: a reference expression database for the meta-analysis of transcriptomes. Adv. Bioinformat. doi: 10.1155/2008/420747.
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dynamics in Arabidopsis plants under oxidative stress induced by cadmium. Free Radical Biol. Med. 47, 1632–1639. Sattarzadeh, A., Franzen, R., and Schmelzer, E. (2008). The Arabidopsis class VIII myosin ATM2 is involved in endocytosis. Cell Motility Cytoskeleton. 65, 457–468. Sattarzadeh, A., Krahmer, J., Germain, A.D., and Hanson, M. (2009). A myosin XI tail domain homologous to the yeast myosin vacuolebinding domain interacts with plastids and stromules in Nicotiana benthamiana. Mol. Plant. 2, 1351–1358. Shimmen, T., and Yokota, E. (2004). Cytoplasmic streaming in plants. Curr. Opin. Cell Biol. 16, 68–72. Sparkes, I. (2010). Motoring around the plant cell: insights from plant myosins. Biochemical Society Transactions. 38, 833–838.
vander Honing, H.S., de Ruijter, N.C.A., Emons, A.M.C., and Ketelaar, T. (2010). Actin and myosin regulate cytoplasm stiffness in plant cells: a study using optical tweezers. New Phytol. 185, 90–102. VanGestel, K., Kohler, R.H., and Verbelen, J.P. (2002). Plant mitochondria move on F-actin, but their positioning in the cortical cytoplasm depends on both F-actin and microtubules. J. Exp. Bot. 53, 659–667. Vidali, L., Burkart, G.M., Augustine, R.C., Kerdavid, E., Tuzel, E., and Bezanilla, M. (2010). Myosin XI is essential for tip growth in Physcomitrella patens. Plant Cell. 22, 1868–1882.
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Wada, M., Kagawa, T., and Sato, Y. (2003). Chloroplast movement. Ann. Rev. Plant Biol. 54, 455–468.
Sparkes, I.A., Ketelaar, T., Ruijter, N.C., and Hawes, C. (2009c). Grab a Golgi: laser trapping of Golgi bodies reveals in vivo interactions with the endoplasmic reticulum. Traffic. 10, 567–571.
Walter, N., and Holweg, C. (2008). Head-neck domain of Arabidopsis myosin XI, MYA2, fused with GFP produces F-actin patterns that coincide with fast organelle streaming in different plant cells. BMC Plant Biol. 8, doi: 10.1186/1471-2229-1188-1174.
Sparkes, I.A., Teanby, N.A., and Hawes, C. (2008). Truncated myosin XI tail fusions inhibit peroxisome, Golgi and mitochondrial movement in tobacco leaf epidermal cells: a genetic tool for the next generation. J. Exp. Bot. 59, 2499–2512. Staiger, C.J., Sheahan, M.B., Khurana, P., Wang, X., McCurdy, D.W., and Blanchoin, L. (2009). Actin filament dynamics are dominated by rapid growth and severing activity in the Arabidopsis cortical array. J. Cell Biol. 184, 269–280. Stephenson, J.L.M., and Hawes, C.R. (1986). Stereology and stereometry of the endoplasmic reticulum during differentiation in the maize root cap. Protoplasma. 131, 32–46. Takemoto, D., Jones, D.A., and Hardham, A.R. (2003). GFP-tagging of cellcomponentsrevealsthedynamicsofsubcellularre-organisation in response to infection of Arabidopsis by oomycete pathogens. Plant J. 33, 775–792. Tominaga, M., Kojima, H., Yokota, E., Orii, H., Nakamori, R., Katayama, E., Anson, M., Shimmen, T., and Oiwa, K. (2003). Higher plant myosin XI moves processively on actin with 35 nm steps at high velocity. EMBO J. 22, 1263–1272.
Yang, Z. (2008). Cell polarity signaling in Arabidopsis. Ann. Rev. Cell Dev. Biol. 24, 551–575. Yokota, E., Ueda, H., Hashimoto, K., Orii, H., Shimada, T., Hara-Nishimura, I., and Shimmen, T. (2011). Myosin XI dependent formation of tubular structures from endoplasmic reticulum isolated from tobacco cultured cells, BY-2. Plant Physiol. doi: 10.1104/pp.1111.175018. Yokota, E., Ueda, S., Tamura, K., Orii, H., Uchi, S., Sonobe, S., HaraNishimura, I., and Shimmen, T. (2009). An isoform of myosin XI is responsible for the translocation of endoplasmic reticulum in tobacco cultured BY-2 cells. J. Exp. Bot. 60, 197–212. Zheng, H., Kunst, L., Hawes, C., and Moore, I. (2004). A GFP-based assay reveals a role for RHD3 in transport between the endoplasmic reticulum and Golgi apparatus. Plant J. 37, 398–414. Zheng, M., et al. (2009). Actin turnover is required for myosin-dependent mitochondrial movements in Arabidopsis root hairs. PLos One. 4, doi: 10.1371/journal.pone.0005961.
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Sparkes, I., Runions, J., Hawes, C., and Griffing, L. (2009a). Movement and remodeling of the endoplasmic reticulum in non dividing cells of tobacco leaves. Plant Cell. 21, 3937–3949.
Ueda, H., Yokota, E., Kutsuna, N., Shimada, T., Tamura, K., Shimmen, T., Hasezawa, S., Dolja, V.V., and Hara-Nishimura, I. (2010). Myosin-dependent endoplasmic reticulum motility and F-actin organization in plant cells. Proc. Natl Acad. Sci. U S A. 107, 6894–6899.
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Volume 4
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Number 5
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Pages 805–812
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September 2011
REVIEW ARTICLE
Recent Advances in Understanding Plant Myosin Function: Life in the Fast Lane Imogen Sparkes1 School of Life Sciences, Oxford Brookes University, Gipsy Lane, Oxford, OX3 0BP, UK
Key words:
Myosin; actin; endoplasmic reticulum; organelle movement; plant development.
INTRODUCTION The advent of live cell imaging and fluorescent probes has confirmed the highly dynamic intracellular nature of plant cells. Whilst plants maybe sessile, intracellularly they certainly live life in the fast lane, with cytoplasmic streaming reaching rates of 100 lm s 1 in Chara (Shimmen and Yokota, 2004). The general phenomenon of streaming was attributed to bulk flow of organelles resulting in a hydrodynamic flow of the cytoplasm (Esseling-Ozdoba et al., 2008). However, quantification of these events has identified what appear to be directed types of movement displayed by several classes of organelle. Organelles do not simply move at the same speed in the same direction; they stop and go, move bi-directionally, and can reach speeds of up to 7–8 lm s 1. It is still a mystery as to why and how plant organelles display such seemingly random movements; however, insights gleaned from organelle positioning and movement in response to biotic and abiotic factors indicate that they do have a functional role. For example, repositioning of the nucleus and chloroplasts in response to light (Wada et al., 2003; Iwabuchi et al., 2007) increased peroxisome movement in response to cadmium (Rodriguez-Serrano et al., 2009), and drastic intracellular rearrangements are seen in response to pathogen attack and mechanical stimuli (Takemoto et al., 2003; Lipka et al., 2005; Hardham et al., 2008). Therefore, intracellular dynamics is analogous to a subcellular roadmap with routes of fast movement on intracellular ‘highways’ and more random meandering movements on the ‘byways’, all of which are implicated in delivery, collection, and shuttling of cargo, metabolites, and signaling intermediates for cellular homeostasis.
Plant myosins are classified into two groups: class XI and class VIII. Class XI is similar to mammalian class V owing to comparable protein domains including the dilute domain in the carboxy terminus, and they also share a common function in organelle movement. Plant myosins contain an amino terminal motor head domain responsible for actin binding and ATPase activity, a regulatory neck region containing IQ motifs, and a carboxy terminal implicated in cargo binding (Figure 1). Genome sequencing has shown that there are 9, 3, 13, and 12 class XI myosins and 2, 5, 4, and 2 class VIII myosins in brachypodium, moss, Arabidopsis, and rice, respectively (Reddy and Day, 2001; Jiang and Ramachandran, 2004; Vidali et al., 2010; Peremyslov et al., 2011). Algae also appear to encode for both class XI and class VIII myosins (Odronitz and Kollmar, 2007; Peremyslov et al., 2011). Recently, there has been an explosion in data relating to plant myosin location (see Table 1), function, and mechanistics (see reviews Shimmen and Yokota, 2004; Li and Nebenfu¨hr, 2008b; Sparkes, 2010, and references therein). Myosins have been implicated in endocytosis (Baluska et al., 2004; Golomb et al., 2008; Sattarzadeh et al., 2008), organelle movement (see below), cytoplasmic streaming and stiffness (Shimmen and Yokota, 2004; Esseling-Ozdoba et al., 2008; van der Honing et al., 2010),
1 To whom correspondence should be addressed. E-mail isparkes@brookes. ac.uk, tel. +44 (0)1865 483639, fax +44 (0)1865 483955.
ª The Author 2011. Published by the Molecular Plant Shanghai Editorial Office in association with Oxford University Press on behalf of CSPB and IPPE, SIBS, CAS. doi: 10.1093/mp/ssr063, Advance Access publication 19 July 2011 Received 4 May 2011; accepted 20 June 2011
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ABSTRACT Plant myosins are required for organelle movement, and a role in actin organization has recently come to light. Myosin mutants display several gross morphological phenotypes, the most severe being dwarfism and reduced fecundity, and there is a correlation between reduced organelle movement and morphological defects. This review aims to discuss recent findings in plants relating to the role of myosins in actin dynamics, development, and organelle movement, more specifically the endoplasmic reticulum (ER). One overarching theme is that there still appear to be more questions than answers relating to plant myosin function and regulation.
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cytokinesis (Collings et al., 2003; Nebenfu¨hr, 2007), plasmodesmata function (Radford and White, 1998, 2011), and viral movement (Avisar et al., 2008a; Harries et al., 2009). Here, this short review will focus on recent discoveries relating to the potential role of myosin in actin organization, organelle movement, more specifically ER remodeling, and plant development. Stromule movement and the role of myosin XI-F are covered in the accompanying review by Kong and Wada (2011) in this special issue.
PLANT MYOSINS AND ACTIN ORGANIZATION
Figure 1. Schematic Representation of Myosin Structure. Schematic representation of a class XI myosin highlighting the amino terminal head domain that binds ATP and actin (blue), the neck region containing six calmodulin binding IQ motifs (green) and coiled coil regions (red), and a carboxy globular tail containing a Dilute domain (black). The globular tail is composed of alpha helices proposed to form two interacting globular tail subdomains (GT1, GT2, see Table 1; Li and Nebenfu¨hr, 2007). Note, this schematic is based on myosin XI-K.
Table 1. Subcellular Location of Arabidopsis Myosin Fusions. Organelle
Myosin
Reference
Peroxisome
FP-XI-2 (1053–1505 aa)a
Reisen and Hanson, 2007
FP-GT1 OR GT2 XI-1 (1099–1301 aa, 1283–1520 aa)b
Li and Nebenfu¨hr, 2007
FP-GT2 XI-2 (1279–1505 aa)b
Li and Nebenfu¨hr, 2007
GT1+GT2; XI-1 (1099–1301 aa, 1283–1520 aa); XI-2 (1099–1297 aa, 1279–1505 aa); XI-I (1115–1290 aa, 1280–1522 aa); XI-K (1124–1325 aa, 1307–1545 aa)c
Li and Nebenfu¨hr, 2007
CCGT+CCGT XI-1 (867–1520 aa)c
Li and Nebenfu¨hr, 2008a
Golgi
FP-GT2 XI-1 (1283–1520 aa)*b
Li and Nebenfu¨hr, 2007
Chloroplast
FP-XI-F (1272–1313 aa)a
Sattarzadeh et al., 2009
Endosomes
FP-ATM1IQtail (838–1167 aa)a
Golomb et al., 2008
a
Golomb et al., 2008
Plasmodesmata
FP-ATM1IQtail (838–1167 aa)
Nuclear envelope
FP-XI-I IQtail/tail (752–1523 aa, 881–1523 aa)a
Avisar et al., 2009
Nucleolus
FP-VIIIB IQtail/tail (813–1127 aa, 905–1127 aa); FP-ATM2 tail (774–1031 aa)a
Avisar et al., 2009
Plasma membrane
FP-ATM1IQtail/tail (838–1167 aa, 943–1167 aa); FP-ATM2 IQtail/tail (688–1031 aa, 774–1031 aa); FP-VIIIA IQtail/tail (820–1154 aa, 925–1154 aa); FP-VIIIB IQtail/tail (813–1127 aa, 905–1127 aa)a
Avisar et al., 2009
Location data are based on transient expression of fluorescent myosin fusions either in tobacco (a) or Arabidopsis (b) leaf epidermal cells. Bimolecular fluorescence complementation assays whereby two regions of the same myosin highlighted in the table were transiently co-expressed in Arabidopsis leaf epidermal cells along with a peroxisome marker highlighted location (c). * 20% of cells showed Golgi rather than peroxisome location. The location of native XI-2 to peroxisomes has been confirmed by immunofluorescence using an anti-XI-2 serum (Hashimoto et al., 2005). Abbreviations denote the following: FP, fluorescent protein; CC, coiled coil domain; GT1/GT2, globular tail region 1/2; aa, amino acid.
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Association of specific plant myosins with actin have been documented; myosin VIII-1 in tobacco BY-2 cells (Yokota et al., 2011) and Arabidopsis XI-2 (also known as MYA2) (Hashimoto et al., 2005) co-sedimented with actin in an ATP-dependent manner, and a XI-2 (MYA2) head fusion construct localized to actin filaments (Walter and Holweg, 2008). In vitro motility assays have shown that XI-1 (also known as MYA1) can
translocate along actin at up to 1.8 lm s 1 (Hachikubo et al., 2007), and a 175-kDa tobacco myosin (75% sequence similarity to XI-2) was shown to process along actin with 35-nm step size at a rate of 7 lm s 1 (Tominaga et al., 2003). Whilst association between myosin and actin implies a role relating to myosincargo motility, it does not preclude roles in regulating actin organization and/or dynamics. Through RNA silencing of two class XI myosins in Moss (XIa, XIb), it was shown that actin organization but not dynamics were altered, resulting in disorganized actin arrays rather than longitudinal filaments (Vidali et al., 2010). In higher plants, Ueda et al. (2010) reported that Arabidopsis xi-k xi-2 double and xik xi-1 xi-2 triple myosin mutants displayed altered actin organization. Actin bundles were randomly orientated and there was a reduction in the number of parallel actin bundles in cotyledonary petiole epidermal cells. A similar distribution of actin bundles was rarely seen in control and single mutants (xi-k, xi-1, xi-2), thus indicating that all three myosins are required collectively to maintain actin bundle organization in the cell types under study. In a further more comprehensive study of triple (xi-k xi-1 xi-2) and quadruple (xi-k xi-1 xi-2 xi-i) myosin mutants on plant development (see below) and actin organization, Peremyslov et al. (2010) documented a similar affect in the leaf midvein cells of both mutants, with actin bundles being orientated up to 90 to the cell axis. Additionally, defects in root hair actin organization were also evident in the triple and quadruple mutants where actin bundles were no longer excluded from the root hair tip. However, observations of leaf pavement and root epidermal cells did not indicate a discernable difference in actin microfilament architecture (Peremyslov et al.,
Sparkes
PLANT MYOSINS AND ORGANELLE MOVEMENT: CHARACTERIZING ER NETWORK REMODELING The majority of organelle movement in plants is actin-dependent and recent advances have begun to assign specific myosin activity to this process. Previously, a role in organelle dynamics was inferred through chemical inhibition studies using drugs that affect actin dynamics (latrunculin b or cytochalasin) or myosin activity (BDM). Thus, myosins have been attributed to the movement of the ER (Quader et al., 1987; Liebe and Menzel, 1995; Sparkes et al., 2009a), Golgi (Boevink et al., 1998; Nebenfu¨hr et al., 1999), mitochondria (Van Gestel et al., 2002; Zheng et al., 2009), peroxisomes (Jedd and Chua, 2002; Mano et al., 2002; Collings et al., 2003), plastids (Wada et al., 2003; Paves and Truve, 2007), and the vacuole (Higaki et al., 2006). A more targeted approach through expression of dominant negative myosin mutants, RNAi, and T-DNA knockouts has revealed several members of the class XI myosins are involved in organelle movement: XI-1, XI-2, XI-C, XI-E, XI-I, and XI-K (see Sparkes, 2010, review and references therein). Expression of dominant negative fusions that lack the motor domain are predicted to either bind to cargo or titrate out accessory factors required for endogenous myosin recruitment, therefore perturbing myosin function. This approach has produced variable results. Truncations can bind to organelles but do not stop their movement (see Table 1), or they are diffuse in the cytoplasm or label motile puncta and stop organelle movement (XI-1, XI-2, XI-C, XI-E, XI-I, XI-K). Studies of myosin tail domain binding to cargo indicate that cargo binding and dimerization are cooperative processes (see Li and Nebenfu¨hr, 2008a, 2008b, and references therein). Inferences from all of these experiments indicate that myosin folding and alterations in conformation are complex and could explain why expression of small myosin fragments have variable affects on cargo binding and movement (see Table 1). Table 1 is included for reference to highlight the specific regions of myosins that have been expressed as fluorescent protein fusions and been assigned to a specific subcellular location. Note, however, that none of the fusions were reported to affect the movement of
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the organelle to which they were bound except for ATM1 IQ tail, which drastically reduced the movement of Ara6-labeled endosomes, yet the movement of FYVE-containing endosomes was only perturbed when a large excess of ATM1IQtail surrounded these structures (Golomb et al., 2008). A detailed comprehensive study of serial deletions and point mutations of conserved residues within myosin tail domains is required to not only address location, but also quantify effects on organelle movement. Future studies with full-length myosin fusions in combination with a genetic approach should yield a more directed experimental approach to understand the functional role and mechanistics of individual myosins. Using the experimental approaches described above, specific myosins have been implicated in having a global affect on the movement of several classes of organelle including mitochondria, peroxisomes, and Golgi (Avisar et al., 2008b; Peremyslov et al., 2008; Prokhnevsky et al., 2008; Sparkes et al., 2008; Avisar et al., 2009; Peremyslov et al., 2010). Roles in stromule (Natesan et al., 2009; Sattarzadeh et al., 2009) and ER dynamics (Sparkes et al., 2009a; Peremyslov et al., 2010; Ueda et al., 2010; Yokota et al., 2011) have recently been added to the list. The ER is a complex network of polygonal tubules interconnected by three-way junctions and anchor points that are proposed to tether it to the plasma membrane. Whilst the majority of the network remodels through tubule growth and shrinkage, tubule sliding to form polygonal rings, and a seemingly ready conversion between tubular and cisternal forms, there appears to be a fast streaming component beneath the more static peripheral network (Figure 2 and Supplemental Movie 1). Cells with a high secretory load display a more cisternal form of ER (Stephenson and Hawes, 1986; Ridge et al., 1999), and those undergoing pathogen attack develop cisternal ER around the site of invasion that can be mimicked by mechanical stimulus (Takemoto et al., 2003; Hardham et al., 2008). Therefore, there appears to be a functional role for ER remodeling. Also, the shear forces generated during ER streaming have been suggested to play a role in cytoplasmic streaming (Kachar and Reese, 1988; Ueda et al., 2010). Immunoblotting of a sucrose density centrifuged microsomal fraction indicated that myosin XI-K co-sedimented with the ER fraction (Ueda et al., 2010). An antibody against a 175kDa tobacco myosin with 75% sequence similarity to XI-2 appeared to label the ER in BY-2 suspension cell culture cells and also co-sedimented with the ER fraction (Yokota et al., 2009). None of the fluorescent fusions to Arabidopsis myosin truncations to date has specifically labeled the ER. However, there did appear to be a correlation between the plasma membrane puncta labeled by ATM1-IQtail fusion and the underlying ER network (Golomb et al., 2008). It remains to be seen whether or not ATM1 plays a role in ER network formation. Two recent studies used different approaches to quantify global ER movement in vivo (Sparkes et al., 2009a; Ueda et al., 2010). Persistency mapping, an analytical approach to characterizing static components (nodes, tubules, cisternae)
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2010). Similarly, expression of tail and IQ tail fusions of the entire Arabidopsis myosin gene family did not exhibit gross rearrangement of actin architecture in tobacco leaf epidermal cells (Sparkes et al., 2008; Avisar et al., 2009). Therefore, it appears that, whilst certain myosins (XI-K, XI-1, XI-2) affect actin organization, this appears to be a tissue-specific phenomenon and it is unclear how the myosin could mediate such an affect. Through the use of BDM, a myosin ATPase inhibitor, Staiger et al. (2009) have reported that myosins have a direct or indirect affect on actin filament buckling and straightening. One proposal could be that myosins provide the tensile force to effectively pull an actin filament straight. Future studies are therefore required to assess whether myosins could effectively tether and provide tensile force on actin filaments, with XI-K, XI-1, and XI-2 being likely candidates for initial studies.
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Figure 2. Persistency Mapping of Cortical ER in Tobacco Leaf Epidermal Cells. Persistency maps of the cortical ER were generated where the first frame (A) and last frame (B) of the movie are shown. Persistent nodes/cisternae (C) and tubules (D) are highlighted. Arrow indicates the direction of streaming that is observed in Supplemental Movie 1, panels a and b. Scale bar = 10 lm. Images taken from Sparkes et al. (2009a). Copyright American Society of Plant Biologists, www.plantcell.org.
in the ER network, highlighted persistent static nodes of ER in tobacco leaf epidermal cells proposed to act as growth or anchor points. A potential role in anchoring the ER perhaps through connectivity to the plasma membrane has also been inferred from optical tweezer studies (Sparkes et al., 2009c). Based on latrunculin b treatment or co-expression of the tail domain of Arabidopsis XI-K, it was evident that both actin dynamics and XI-K are required to maintain ER remodeling (Sparkes et al., 2009a). A separate study characterizing ER dynamics in the cotyledonary petioles of Arabidopsis single, double, and triple T-DNA insertional myosin mutants also indicated that XI-K plays a role, with XI-1/2 exerting minor affects on ER streaming (Ueda et al., 2010). An in vitro reconstitution assay has been used to determine the role of actin and myosin in generating the tubular ER network (Yokota et al., 2011). Through density centrifugation, a microsome fraction containing cytosol was separated and the effects on tubular-like network regeneration in the presence and absence of actin, energy (ATP/GTP), a 175-kDa class XI myosin, and a class VIII myosin was observed. Results indicated that formation of a network-like structure reminiscent of the ER required all of the latter components except the class VIII myosin. GTPase activity could be attributed to a requirement for Rabs or perhaps a role of RHD3 in the experimental system as the rhd3-1 mutant has unbranched ER tubules (Zheng et al., 2004).
Plant growth and development are complex coordinated processes affected by extracellular signals such as light and temperature. The majority of plant cells undergo anisotrophic growth (i.e. expansion along a preferred axis). At the cellular level, cytoskeletal elements are important in controlling expansion, elongation, and regulated shape changes through deposition of microtubules and actin microfilament patches (Yang, 2008). This coordinated process allows the targeting and trafficking of components required for growth such as the cellulose synthase complex and mediates endo/exocytotic events important for tip growth. Therefore, disruption in this finely tuned balance, and the molecular motors that traverse the cytoskeletal networks, can have drastic changes in the final body plan of the plant. There is a distinct correlation between plant growth defects and the role of myosins. The exact biological role(s) of, for example, secretion and endocytosis, resulting in these developmental defects, is unclear at this time, but a role in actin remodeling and organelle movement has been shown (see earlier). Root hairs undergo polarized tip growth that is due to the addition of membrane and wall material at the tip of the cell. A small group of Arabidopsis single myosin mutants (xik and xi2) display shorter root hairs (Ojangu et al., 2007; Peremyslov et al., 2008). Double mutants of paralogous myosin pairs indicated an additive affect for XI-K, XI-2, and XI-B on root hair length (Prokhnevsky et al., 2008). Defects in moss protonemal tip growth are also evident in a double myosin mutant xia xib (Vidali et al., 2010). In quadruple Arabidopsis mutants, defects in cells undergoing expansion through diffuse growth were observed and appeared to correlate with cell size; moderate reduction in mesophyll cell size compared to an increased reduction in the larger midvein cells (Peremyslov et al., 2010). Stunted growth (reduced rosette diameter, plant height, root length) and reduced flowering time were observed in triple and quadruple mutants, with xi-k xi-1 xi-2 xi-i displaying the severest phenotype (Peremyslov et al., 2010). Golgi and peroxisome movement in this mutant was reduced to ‘residual wobbling’
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Based on the experimental approaches discussed above, it has been suggested that the acto-myosin system is required for ER tubule growth and shrinkage through actin polymerization and myosin processivity. Movement effectively pulls out ‘excess’ membrane from cisternal regions to form the tubular network and sliding of tubules forms closed polygonal rings (Sparkes et al., 2009a). In the absence of cytoskeletal elements, the ER network is stabilized presumably through anchor point connections to the plasma membrane and scaffolding/stabilizing elements such as reticulons (see Sparkes et al., 2009b, and references therein). It will be interesting to see whether and how several myosins coordinate this process and whether the coordination is through specific myosin–ER interactions, or regulation of myosin interaction through homo and heterodimerization of myosin complexes.
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Figure 3. Expression Patterns of Arabidopsis Myosin Genes. Public affymetrix DNA microarray data were assessed using Genevestigator (www.genevesigator.com; Hruz et al., 2008) to generate a heat map of Arabidopsis myosin gene expression in various tissues. Predicted expression levels are indicated in the top left-hand corner inset, where dark blue refers to the highest predicted level of expression.
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attributed to Brownian movement or a ‘tug-of-war’ between remaining myosins failing to move either class of organelle in a processive manner (Peremyslov et al., 2010). The double myosin mutant xi-k xi-1 also displayed reduced fecundity and was stunted (Prokhnevsky et al., 2008). Therefore, it appears that reduced plant stature is in part due to reduced cell size, but, as was documented for leaf pavement and mesophyll cells, also a reduction in cell number (Peremyslov et al., 2010). Given the large number of myosin genes in plants, and the apparent lack of phenotypic defects in the majority of single mutants, it has been proposed that there is functional redundancy within the family. However, whilst the single myosin mutant studies monitored and quantified affects in certain tissues (root hair length, for example), a comprehensive study of all tissues and stages of development has not been provided. Therefore, potential defects in tissues shown by microarray analysis to express these genes (Figure 3) in wild-type plants should also be monitored. An additional hypothesis is that certain myosins have specific roles under perturbed noncontrolled growth conditions such as altered light and temperature. Peremyslov et al. (2011) analyzed microarray data and have characterized the expression profile of Arabidopsis and rice myosin genes. Interestingly, the general trend is that expression follows photocycles and thermal cycles, and three appear to display a pattern indicative of circadian regulation (rhythmic expression irrespective of light/dark cycles; XI-B, XI-H, VIIIB). Photoperiod regulation was previously shown for myosin XI-B in rice (Jiang et al., 2007). Based on these expression profiles, it will be interesting to see whether single and multiple myosin mutants display aberrant morphologies under altered growth conditions, and also in response to abiotic and biotic factors. Another interesting level of regulation could be through dimerization with headless domain variants that, in effect, could mimic the experimental response seen through overexpression of myosin tail fusions. Such a transcript for XI-K was revealed by Peremyslov et al. (2011) and expression was shown through a GUS reporter trap. Further bioinformatic mining of the genome combined with experimentation will reveal whether similar headless variants are present and represent true transcripts rather than genetic relics, and whether they play a role in myosin regulation.
actin and microtubule-dependent in elongating internodal cells (Foissner et al., 2009), and a role for kinesin 13-a in Golgi positioning has been suggested (Lu et al., 2005).
FUTURE DIRECTIONS
Foissner, I., Menzel, D., and Wasteneys, G.O. (2009). Microtubuledependent motility and orientation of the cortical endoplasmic reticulum in elongating characean internodal cells. Cell Motility Cytoskeleton. 66, 142–155.
Supplementary Data are available at Molecular Plant Online.
FUNDING Some of the work reported here was supported by Oxford Brookes University, the BBSRC, and the Leverhulme Trust.
ACKNOWLEDGMENTS I would like to thank Chris Hawes for critical reading of the manuscript and Larry Griffing for kindly agreeing to use his movie published in Sparkes et al. (2009a). No conflict of interest declared.
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