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Perico, C., & Sparkes, I. (2018). Plant organelle dynamics: cytoskeletalcontrol and membrane contact sites. New Phytologist, 220(2), 381-394.https://doi.org/10.1111/nph.15365
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Link to published version (if available):10.1111/nph.15365
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For Peer Review
Plant organelle dynamics: cytoskeletal control and
membrane contact sites
Journal: New Phytologist
Manuscript ID NPH-TR-2018-26701.R1
Manuscript Type: TR - Commissioned Material - Tansley Review
Date Submitted by the Author: n/a
Complete List of Authors: Perico, Chiara; University of Bristol, School of Biological Sciences Sparkes, Imogen; University of Bristol, School of Biological Sciences
Key Words: actin, dynamics, membrane contact site, myosin, organelle, tether, cytoskeleton
Manuscript submitted to New Phytologist for review
For Peer Review
1
Plant organelle dynamics: cytoskeletal control and membrane contact sites 1
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Chiara Perico1, Imogen Sparkes
1* 4
1 School of Biological Sciences, University of Bristol, 24 Tyndall Avenue, Bristol, BS8 1TQ, UK 5
6
*Corresponding author 7
i.sparkes@bristol.ac.uk 8
+44 (0)117 39 41211 9
10
Word count: 7599 (excluding summary, reference list and figure legends) 11
4 figures: Fig1, 2 and 4 in colour, Fig 3 in black and white 12
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Summary 14
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Organelle movement and positioning are correlated with plant growth and development. Movement 16
characteristics are seemingly erratic yet respond to external stimuli including pathogens and light. 17
Given these clear correlations, we still do not understand the specific roles that movement plays in 18
these processes. There are few exceptions including organelle inheritance during cell division, and 19
photorelocation of chloroplasts to prevent photodamage. The molecular and biophysical 20
components that drive movement can be broken down into cytoskeletal components, motor 21
proteins and tethers which allow organelles to physically interact with one another. Our 22
understanding of these components and concepts has exploded over the past decade, with recent 23
technological advances allowing an even more in depth profiling. Here, we provide an overview of 24
the cytoskeletal and tethering components and discuss the mechanisms behind organelle movement 25
in higher plants. 26
27
Keywords: Actin, dynamics, cytoskeleton, membrane contact sites, myosin, organelle, tether 28
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Introduction 32
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A defining characteristic of eukaryotic life is subcellular compartmentalisation in the form of 34
membrane bounded organelles. Organelles allow spatial separation of components required to carry 35
out key biochemical reactions in distinct microenvironments. However, biochemical pathways can 36
span multiple organelles requiring exchange of key metabolites. How can exchange occur if 37
organelles are spatially separated within the cytoplasmic milieu? How can an organelle deliver 38
components to targeted sites? Ultimately, how do organelles ‘communicate’ with one another, and 39
what controls the choreography of such events? 40
Since the early observations of cytoplasmic streaming events, it is clear that organelles whilst being 41
discrete entities are not static, they are highly motile. A basic analogy can be drawn between a cell 42
and an island, where organelles are cars with cytoskeletal components acting as roads. For the island 43
(cell) to function there must be clear communication allowing delivery and transfer of goods in a 44
coordinated manner. The emerging picture is that in eukaryotic systems organelle movement is 45
dynamic with physical interactions occurring between organelles allowing ‘communication’ and 46
exchange to occur. These exchanges occur at membrane contact sites (MCS). Exchange between 47
organelles also occurs via vesicle mediated pathways (Bonifacino & Glick, 2004). Ultimately, 48
organelle movement is therefore driven by an interplay between physical interaction between 49
organelles and motor driven activity to allow separation and independent motion. 50
The combination of technological advances in imaging, genetic manipulation of model systems and 51
introduction of a multitude of fluorophores has enabled a relatively fast description of basic 52
parameters relating to organelle dynamics and interactions. Here, we introduce the elements that 53
control organelle dynamics in higher plant cells, and why it is important that we unpick the 54
molecular mechanisms controlling movement and positioning of organelles. 55
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Basic movement characteristics 57
58
Cytoplasmic streaming is an actin dependent event resulting in the bulk flow of cytoplasm in a 59
directed manner (Shimmen, 2007; Tominaga & Ito, 2015). In cell types undergoing polarised tip 60
growth (root hairs and pollen tubes) streaming events are ordered with flows along the outer wall 61
towards the tip changing direction and flowing down the central axis (Hepler et al., 2001). This 62
process is called reverse fountain streaming. Other cell types however display less ordered ‘patterns’ 63
of movement with streams rotating and bifurcating in a seemingly random manner. Modelling the 64
process of cytoplasmic streaming events indicates that actin dynamics themselves (sliding of actin 65
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filaments, polymerisation / depolymerisation) cannot generate the required shear forces in the 66
cytoplasm to generate bulk flow. However, the movement of a net like structure, which effectively 67
has wider ‘filaments’ than actin, being dragged through the cytoplasm could produce bulk flow of 68
the cytoplasm (Nothnagel & Webb., 1982). These studies infer that it is the movement of the ER 69
which generates the observed flows. Based on correlations between ER and actin rearrangements, 70
Ueda et al. postulated that movement of the ER may act to rearrange the actin filaments (Ueda et 71
al., 2010). More recently, data indicated that changes in ER network movement were correlated 72
with changes in streaming rates based on movement of discrete organelles such as Golgi and 73
mitochondria (Stefano et al., 2014). In summary, cytoplasmic streaming is complex and the 74
molecular mechanism is likely driven through an interplay between actin and organelle movement. 75
Note, while studies in Chara have provided advances in our understanding of cytoplasmic streaming 76
(Nothnagel & Webb, 1982; Kachar & Reese, 1988; Woodhouse & Goldstein, 2013; Goldstein et al. 77
2008), these may not directly map on to angiosperms. For example, differences in myosin 78
mechanochemical properties, organelle and actin filament positioning may all impact streaming, and 79
the modelling of such events. 80
81
Initially it was proposed that organelle movement was a passive process with organelles moving 82
along in cytoplasmic streams. With the development of tracking algorithms and implementation of 83
fluorescent probes targeted to organelles, it quickly became clear that individual organelles can 84
display a multitude of movement characteristics within a short time frame, and that movement 85
outside of the streams is directed and not passive. For example, smaller seemingly ‘spheroid’ 86
organelles observed with a light microscope, such as Golgi, mitochondria and peroxisomes undergo 87
‘stop-go’ movement; fast motion with acceleration, deceleration, pausing and changing direction 88
with speeds reported up to 8-10µm/s (Boevink et al., 1998; Nebenführ et al., 1999; Logan & Leaver, 89
2000; Jedd & Chua, 2002; Mano et al., 2002; Mathur et al., 2002; Van Gestel et al., 2002). It has been 90
suggested that sites of organelle pausing occurs at cross-over points between actin and microtubule 91
filaments, which can occur around ER-PM anchor points (see later). The ER is a network of 92
interconnected tubules and cisternae which readily interconvert. Movement here is through tubule 93
extension and retraction, tubule fusion and lateral sliding to form new polygons, with possible 94
polygonal filling to generate new cisternae (Sparkes, I et al., 2009). In addition to movement 95
(remodelling) of the ER network structure, movement of membrane proteins (surface flow) also 96
occurs (Sparkes, I et al., 2009; Runions , J et al., 2006). An example of ER (green) and Golgi 97
(magenta) movement over a 30 second period in Arabidopsis leaf epidermal cells is depicted in 98
Figure 1. Comparing 3 individual frames (0, 15 and 30 seconds) indicates the dramatic level of 99
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movement of both the ER network and individual Golgi bodies; merge image (h and l) highlights 100
Golgi and regions of the ER which have remained relatively stationary (white) compared with other 101
regions within the network / discrete Golgi. It is important to note that the vast majority of discrete 102
organelles appear to be in close association with the ER (Fig1a-c). Whether this reflects a true 103
physical connection between organelles, or is an indirect effect due to the large central vacuole 104
effectively forcing cytoplasmic contents together or organelles occupying the same cytoskeletal 105
track is unclear (see later). 106
107
Actin and associated motors, myosins, play a primary role in plant organelle movement and 108
positioning. 109
Organelle re-positioning within the plant cell cytoplasm largely depends on the actin cytoskeleton 110
and on myosins, the associated motor proteins (Boevink et al., 1998; Nebenführ et al., 1999; Mathur 111
et al., 2002; Van Gestel et al., 2002; Sparkes et al., 2008; Avisar, Dror et al., 2009). Myosin motors 112
generally consist of an N-terminal motor domain which binds actin in an ATP dependent manner, a 113
neck domain with calmodulin-binding IQ-repeats and a region of dimerization, and a C-terminal 114
globular tail domain (GTD) involved in cargo binding. 115
Actin monomers (G-actin) polymerise under physiological conditions to form filaments (F-actin) 116
which constitute a dynamic, rail-like structure. It is generally thought that myosin dimers are 117
recruited to organelle membranes and proceed on these tracks in a step-wise ATP-dependent 118
manner towards the faster growing plus end, dragging the organelle along the actin filament. 119
Disruption of actin with depolymerising drugs such as Latrunculin B (LatB) or Cytochalasin D (CytD) 120
halts organelle movement and, more generally, cytoplasmic streaming (Nebenführ et al., 1999; 121
Mathur et al., 2002) . While these studies have indeed highlighted the central role of the actin 122
network in organelle dynamics, it does not indicate regulatory mechanisms to allow specific control 123
of the movement and positioning of one organelle over another. This is provided through myosin 124
interaction, and organelle interactions at membrane contact sites. 125
Based on the structure of the conserved motor domain, two sub-classes of myosin have been 126
identified in the model plant Arabidopsis thaliana: class VIII and class XI, which include 4 and 13 127
members respectively (Foth et al., 2006; Peremyslov et al., 2011). Myosins from class VIII have been 128
observed to localise at the cell plate (Damme et al., 2004), plasmodesmata (Golomb et al., 2008), 129
nucleolus (Avisar, Dror et al., 2009) and in association with phragmoplast microtubules (Wu & 130
Bezanilla, 2014). In higher plants they do not appear to control organelle movement, and, as 131
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suggested by Ryan and Nebenführ, may act as tensors interacting with actin to control tension 132
within the network (Ryan & Nebenführ, 2018). Class XI myosins contain a conserved Dilute domain 133
and are similar to class V myosins from mammals and fungi (Foth et al., 2006). Class V and XI 134
myosins are involved in organelle movement. In higher plants, the role of class XI myosins has been 135
determined through knock out, RNAi and expression of dominant negative myosin fusions. The latter 136
is proposed to act in a dominant negative manner as fusions lack the motor domain which could 137
prevent movement, yet retain the cargo binding domain. Transient over-expression of dominant 138
negative tail and IQ-tail of all the 17 A.thaliana myosins in N.tabacum and N.benthamiana leaf 139
epidermal cells highlighted the role of multiple myosins in controlling Golgi, mitochondria and 140
peroxisome dynamics (Sparkes et al., 2008; Avisar, Dror et al., 2009). Tail and IQ-tail of myosins XI-1 141
(MYA1), XI-2 ( MYA2), XI-C, XI-E, XI-I, and XI-K significantly decrease Golgi displacement rate in both 142
Nicotiana species, without affecting actin organisation or colocalising to the Golgi membrane (Avisar, 143
Dror et al., 2009). Interestingly, IQ-tail of the same myosins also inhibit mitochondria movement 144
(Avisar, Dror et al., 2009), whilst myosin XI-E and XI-K tail fusions were observed to stop peroxisome 145
movement (Sparkes et al., 2008), thus suggesting that a single myosin can control the dynamics of 146
multiple organelles. It was reported that Arabidopsis knock-outs (KO) in single myosin XI genes didn’t 147
display a noticeable phenotype, apart from xi-k and xi-2 plants, which display a reduced root hair 148
growth (Peremyslov, Valera V. et al., 2008). Much like what happens upon the expression of XI-K and 149
XI-2 dominant negative mutants, Golgi, peroxisomes and mitochondria movement drastically 150
decrease in leaves of xi-k and xi-2 Arabidopsis lines (Peremyslov, Valera V. et al., 2008). Considering 151
that multiple class XI myosins appear to control global organelle movement, with XI-K and XI-2 152
having the most marked effect, implies a certain degree of functional redundancy within the class XI 153
myosin gene family. 154
Considering that several myosins control global organelle movement, one would hypothesis that 155
they would be located to the surface of the organelle in question. Myosin localisation data through 156
expression of fluorescent fusions to full length and truncated myosins, and immunohistochemistry 157
data have not provided a clear answer to this issue. The vast majority of data for class XI myosins 158
indicate diffuse cytoplasmic localisation, or to unknown puncta / vesicles; IQ-Tail and tail fusions are 159
diffuse or on unknown puncta (Avisar, D. et al., 2009), ‘DIL’ and ‘PAL’ domains are frequently on 160
unknown puncta (Sattarzadeh et al., 2011; Sattarzadeh et al., 2013), XI-K full length fusion is on 161
unknown highly motile vesicles (Peremyslov et al., 2012). There are few exceptions including 162
fluorescent fusions of XI-I residing on the nuclear envelope (Avisar, D. et al., 2009; Tamura et al., 163
2013), expression of portions of the GTD of XI-1 located to either peroxisomes and / or Golgi (Li & 164
Nebenführ, 2007), and immunohistochemistry indicating XI-2 on peroxisomes and additional puncta 165
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(Hashimoto et al., 2005). How can we reconcile this disconnect between myosin localisation and 166
activity on organelle movement? There are several hypothesis; (1) the number of functional myosin 167
molecules required to generate movement is small and so expression of truncations may bind and 168
saturate receptors on the organelle surface, which are ‘masked’ and indistinguishable from the 169
higher levels present in the cytoplasm, (2) cytoplasmic levels titrate out components required to 170
recruit endogenous myosins to the surface of the target organelle, in effect inhibiting its action. 171
Considering that results from expression of truncated myosin fusions are corroborated by knockout 172
(T-DNA) and knock down (RNAi) studies highlights all three experimental approaches are bona fide 173
and results are not artefactual. Given that seemingly opposing methods of study, which either 174
downregulate (T-DNA, RNAi) or ‘upregulate’ myosin levels (albeit a non-functional motorless 175
variant), lead to the same experimental outcome may be due to truncations heterodimerising with 176
endogenous myosins and effectively preventing formation of functional myosin dimers. In addition, 177
expression of XI-K tail fusions containing point mutations indicates that two arginine residues control 178
the dominant negative phenotype (Avisar et al., 2012). Here, it was suggested that perhaps the tail 179
fusions bind to the head domains of endogenous myosins inhibiting action. This is based on the 180
arginine residues in XI-K corresponding with Lysine residues in myoVa which are important for 181
affecting tail to head interactions which render myoVa inactive (Li et al., 2008). Similar head-tail 182
interactions in plants myosins have not been confirmed. The exact mode of action of complete 183
myosin molecules and the dominant negative results displayed through the expression of myosin 184
truncations, will remain a mystery until a greater understanding of intramolecular (such as possible 185
head to tail) and intermolecular (myosin dimerization) interactions, and components which regulate 186
myosin recruitment and activity are determined. 187
Mechanisms of myosin recruitment: a tightly regulated system? 188
Andreas Nebenführ’s group have begun to decipher intra- and intermolecular interactions and 189
effects on organelle localisation (Li & Nebenführ, 2007; Li & Nebenführ, 2008). Here, the suggestion 190
is that regions within the GTD specify the target organelle, and that dimerization stabilises 191
association with the organelle surface; portions of the XI-1 GTD locate to ether Golgi or peroxisomes, 192
expression of full length tail (including GTD) dimerise and stabilise binding to organelles. While these 193
results provide interesting insight into the mechanism behind myosin recruitment, we are still some 194
way off from understanding how this occurs within a complete, rather than truncated, myosin 195
molecule, and whether interacting proteins mediate conformational changes within the myosin 196
hetero/homodimers to enable targeting to specific organelles. 197
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The large number of myosins in plant cells, and their seemingly overlapping functions, suggests that 198
identification of additional components at the myosin-organelle interface is required to provide 199
specificity in recruitment and regulation of myosin activity. Studies from mammals and yeast suggest 200
a myosin recruitment process mediated by receptors and adaptor proteins, either through a single 201
membrane protein (Fagarasanu et al., 2006) or larger complexes including Rab GTPases, receptors 202
and adaptors (Tang et al., 2003; Yan et al., 2005; Hume et al., 2007; Pylypenko et al., 2013). For 203
instance, myosin Myo2p of Saccharomyces cerevisiae is required for both peroxisome and vacuole 204
inheritance. Coordination of these organelles towards the bud is mediated by different receptors 205
and adaptors, which are independently regulated, at the interface between Myo2p and the 206
peroxisomal (Fagarasanu et al., 2006) or vacuolar (Tang et al., 2003) membranes. Interestingly, 207
Arabidopsis XI-2 was shown to interact in vitro with the small GTPases AtRabD1 and AtRabC2a via 208
distinct regions in its tail domain (Hashimoto et al., 2008). 209
There are few examples showing interactor binding to a myosin affects the movement of the 210
organelle to which it is bound. A.thaliana myosin XI-I specifically interacts with the proteins WIT1 211
and WIT2 located to the outer nuclear membrane (Fig.2a). Lack of either myosin XI-I or one of the 212
two WIT proteins diminishes dark-induced nuclear positioning (Tamura et al.). Notably, expression of 213
XI-I tail fusion also influences the dynamics of other organelles (Avisar, Dror et al., 2009), whereas 214
lack of WIT1/WIT2 only influences nuclear movement. Therefore, interaction between XI-I and 215
WIT1/2 controls nuclear migration. Other examples of myosin interactors include DECAPPING 216
PROTEIN 1 on P-bodies and RISAP on TGN (Steffens et al., 2014; Stephan, O. et al., 2014). 217
Recent advances in myosin recruitment studies in A.thaliana led to the identification of two new 218
classes of myosin-binding proteins: the MyoB family of potential myosin receptors (Peremyslov et 219
al., 2013; Peremyslov et al., 2015) and the MadA/MadB adaptor proteins (Kurth et al., 2017). A 220
yeast-2-hybrid (Y2H) screen on a cDNA library using the globular tail domain of myosin XI-K from 221
A.thaliana as the bait led to the identification of the MyoB family of potential myosin receptors 222
(Peremyslov et al., 2013). Whilst the lengths and architectures of the 16 identified potential 223
receptors are variable, they all share a plant specific myosin-binding DUF593 domain (Domain of 224
Unknown Function 593), originally associated with the product of the Floury1 gene in Z.mays and 225
hence named “Zein-binding domain”(Holding et al., 2007). DUF593 proteins are also present in rice 226
(Oryza sativa) (Holding et al., 2007) and in Nicotiana species (Stephan, Octavian et al., 2014; 227
Peremyslov et al., 2015). The interpretation of the role of MyoB potential receptors in myosin 228
recruitment has so far been quite challenging. Unlike other interactors including WIT1/WIT2, 229
fluorescent fusions of MyoB do not localise to known organelles. Instead, they are on highly motile 230
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unidentified membrane bounded punctae identified as unknown vesicles (Peremyslov et al., 2013; 231
Peremyslov et al., 2015; Kurth et al., 2017). The MyoB vesicles move at higher average speeds than 232
organelles such as Golgi, mitochondria and peroxisomes (Sparkes et al., 2008; Avisar, Dror et al., 233
2009; Peremyslov et al., 2015). Transformation of MyoB1-YFP into Columbia, xi-k, and xi-k/xi-1/xi-2 234
backgrounds showed that their movement and localisation is myosin-dependent (Peremyslov et al., 235
2013); MyoB1-YFP punctae proceed at high speeds in linear tracks in Columbia wild type 236
background, whereas in knockout backgrounds, and especially in the triple knockout, MyoB1-YFP 237
shows a more diffuse localisation with a few immotile punctae. Moreover, the DUF593 domain 238
seems to directly mediate the binding of MyoB potential receptors with myosins; DUF593 of 239
AtMyoB2 binds to myosin XI-K GTD in vitro (Peremyslov et al., 2013), although it is not known where 240
this interaction takes place. Similarly, the TGN-localised potential myosin receptor NtRISAP, interacts 241
in vitro with the GTD of a tobacco pollen tube myosin XI through DUF593 (Stephan, Octavian et al., 242
2014). Transient over-expression of the DUF593 domain from NbMyoB1 and NbMyoB2 in 243
N.benthamiana leads to a strong decrease in Golgi, peroxisomes and mitochondria mean velocities 244
(Peremyslov et al., 2015), perhaps suggesting a similar effect to that obtained through over-245
expression of dominant negative myosin tails (Avisar, Dror et al., 2009). Single gene knock-out (KO) 246
of myob1-4 genes does not lead to any noticeable phenotype, whereas triple 247
(myob1/myob2/myob3) and quadruple (myob1/myob2/myob3/myob4) KOs present delayed 248
flowering and reduced height (Peremyslov et al., 2013; Peremyslov et al., 2015). Interestingly, xi-249
k/xi-2/myob1/myob2/myob3 (5KO) plants show exacerbated developmental defects when 250
compared to either xi-k/xi-2 (2KO) or myob1/myob2/myob3 (3KO) plants (Peremyslov et al., 2015), 251
suggesting that MyoB proteins and myosins might be effectively involved in the same processes. 252
Peremyslov et al (Peremyslov et al., 2015) suggest that the MyoB compartment could play a major 253
role in driving cytoplasmic streaming, actively moving on the actin filament at high velocities in a 254
myosin-dependent way and thus creating the motive force for other organelles to move into the 255
cytoplasm. Whether the MyoB vesicles actually define a novel membrane compartment or a 256
specialised subdomain of a known organelle remains unclear. 257
Two novel classes of myosin-binding proteins, recently uncovered by Kurth et al. (Kurth et al., 2017), 258
are the MadA and MadB proteins. These two independent families, formed of four members each, 259
are structurally different from one another and from the MyoB family (Peremyslov et al., 2013; 260
Kurth et al., 2017) but, just as the MyoB and myosin families they appear to be conserved in all land 261
plants (Kurth et al., 2017). Aside from MadA1, which localises to the nucleoplasm, Mad proteins 262
locate to motile punctae and their localisation is myosin-dependent, similar to certain MyoBs 263
(Peremyslov et al., 2013; Kurth et al., 2017). Interestingly, single KO plants for the MadB1 gene show 264
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reduced root hair length (Kurth et al., 2017), much like what had been described for xi-2 and xi-k 265
mutants (Peremyslov, Valera V. et al., 2008). To determine whether Mad and MyoB proteins interact 266
with myosins a yeast two hybrid screen using XI-K, XI-1, XI-I, XI-F and XI-C as bait was performed. It 267
was found that numerous permutations of Mad and MyoB interactions with myosins appeared to 268
occur. Assuming that these interactions are representative of those in planta could infer that 269
control of organelle movement occurs through multiple rounds of myosin interaction with different 270
interactors to fine tune control within a short time frame. Alternatively, these combinations could 271
reflect tissue specific or developmental regulated differences or activation under certain stimuli. 272
How can we reconcile observations of organelle movement with the effects of myosins and myosin 273
interactors to generate a mechanistic model accounting for movement? The more traditional view of 274
myosins being recruited to the surface of the organelle whose movement it controls appears to hold 275
for XI-I’s control of nuclei movement (Fig 2a, b). A model for passive motion of organelles through 276
movement in cytoplasmic streams has been suggested based on MyoB / MadA-B work (Fig 2c). 277
AtMyoB2 interacts with the GTD of XI-K, both collocate to highly motile structures which move faster 278
than other spheroid organelles. XI-K is also implicated in cytoplasmic streaming. These observations 279
have led to the proposal that MyoB/MadA-B vesicles bound by myosin drive streaming, which in turn 280
affects organelle movement in a passive manner (Fig 2c). Alternatively, perhaps MyoB/MadA-B 281
vesicles are in fact decorating subdomains of organelles, such as the ER, which in turn drive 282
movement in a directed manner (Fig 2b,d). 283
284
Microtubules, associated motors, and interplay with actin 285
Two types of motor proteins are associated with microtubules: kinesins, plus-end directed, and 286
dyneins, minus-end directed. So far, sixty-one potential kinesin encoding genes have been identified 287
in the A.thaliana genome, 21 of which are suggested to be minus-end directed kinesins based on the 288
presence of a particular peptide within the neck region (Reddy & Day, 2001), which could suffice for 289
the apparent absence of dynein genes. Notably, not all plants lack genes potentially encoding for 290
dyneins: for example, four predicted dynein heavy chains have been identified in the O.sativa 291
genome (King, 2002). Similarly to myosins, kinesins general architecture is composed of a motor 292
domain, a stalk and a tail and they are classified into multiple subfamilies, named kinesin-1 to 293
kinesin-14, based on similarities in their motor domain (Lawrence et al., 2004). Unlike in mammals 294
and some fungi, microtubules and possibly kinesins appear to play a minor, but significant, role in 295
organelle movement in plants through short range rather than long distance trafficking associated 296
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with actomyosin system (reviewed in Brandizzi & Wasteneys, 2013; Cai et al., 2015). Treatments 297
with microtubule depolymerizing drugs doesn’t seem to affect long range movement of organelles 298
(Nebenführ et al., 1999; Lovy-Wheeler et al., 2007), but might affect their distribution (Van Gestel et 299
al., 2002; Idilli et al., 2013). It has been suggested that here, microtubule dependent motion allows 300
organelles to pause. Golgi bodies are known to move in the cell in an acto-myosin dependent way 301
(Boevink et al., 1998; Nebenführ et al., 1999; Sparkes et al., 2008), and to pause at specific sites in 302
correspondence of cortical microtubules (Crowell et al., 2009). Similar observations have been made 303
for mitochondria and peroxisomes pausing at actin microtubule junctions. However, pausing may 304
not be microtubule dependent events as it occurred upon microtubule depolymerisation but was 305
noted to occur at ER junctions (Hamada et al., 2012). Given that stable ER junctions are possible sites 306
of tethering to the PM (see later), perhaps organelles pause either due to a physical blockage from 307
the ER, actin, microtubule complex, or perhaps pausing serves a functional role through 308
communication at ER-PM sites (Bayer et al., 2017). In addition to actin, microtubules have been 309
shown to contribute to ER tubule elongation and anchoring in A.thaliana (Hamada et al., 2014), and 310
exo- and endocytosis and endosomal movement in tobacco pollen tube tip (Idilli et al., 2013). 311
Moreover, kinesins have been detected on organelle membranes; Kinesin-13A on Golgi stacks (Lu, 312
Ling et al., 2005) and Golgi-associated vesicles (Wei et al., 2009), KAC1 and KAC2 on chloroplast 313
(Suetsugu et al., 2010), which are interestingly able to bind actin. In At kinesin-13 mutant plants, 314
Golgi stacks aggregate in trichomes and in other epidermal cells, with trichomes often exhibiting 4 315
branches instead of 3 (Lu, L. et al., 2005). Although some Golgi stacks were found to be associated 316
with microtubules, it is currently unknown whether Kinesin-13 has an active role in moving Golgi 317
along microtubules. 318
Interestingly, there may be functional cooperation and coordination between actin and 319
microtubules which could affect organelle dynamics. Studies in A.thaliana seedlings treated with 320
cytoskeleton stabilizing and de-stabilizing drugs suggest a coordinated re-organisation of actin and 321
microtubules (Sampathkumar et al., 2011). It has been observed that some plant formins, proteins 322
traditionally associated with regulating actin dynamics, can also associate and bind to microtubules 323
(Deeks et al., 2010; Wang et al., 2013; Sun et al., 2017). The rice formin OsFH15, for example, was 324
shown to bind to and induce bundling of both actin and microtubules, being also able to cross-link 325
the two cytoskeletal networks (Sun et al., 2017). Therefore, actin and microtubules are intimately 326
linked and coordinated networks. Myosins, classically thought of as ‘binding’ organelles and 327
controlling their movement directly, may in fact control organelle movement indirectly through 328
controlling actin dynamics (Peremyslov et al., 2010; Cai et al., 2014; Madison et al., 2015). Therefore, 329
could myosins affect both actin and microtubule networks directly or indirectly respectively? 330
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331
In conclusion, both actin and microtubules are important regulators of organelle dynamics. Future 332
research into coordinated activities, either through myosin / kinesin or dynamics of their 333
corresponding cytoskeletal tracks, will help decipher the molecular cues resulting in the observed 334
seemingly chaotic movement displayed in cells undergoing diffuse growth. 335
336
Role of organelle interactions: tales of tethers 337
338
Organelle interactions occur at membrane contact sites (MCS), where membranes from apposing 339
organelles are within 10-30nm of one another. In mammals and yeast numerous interactions at 340
MCS’ between various organelle pairs, and the molecular components required for physical 341
association, have been elucidated. These interactions play important roles in exchange of 342
components such as lipids and calcium (Prinz, 2014). Therefore, physical interaction between 343
organelles plays a functional role in communication between organelles, and therefore affects 344
organelle positioning. 345
In plants, the role of MCS is currently limited to the identification of a physical association of a 346
handful of organelle pairings (such as ER-Golgi, peroxisome-chloroplast), with scant identification of 347
the molecular components involved at the tether site (eg. CASP for ER-Golgi, retromer components 348
for oil body-peroxisome). Here, we will focus on each organelle pairing in turn, provide a brief 349
rational for the functional role for association, and the evidence that interaction occurs. Note, 350
correlation of movement between two closely opposed organelles has been assumed to reflect 351
physical tethering between compartments. Isolation and characterisation of bona fide molecular 352
tethering components will provide more conclusive evidence on interaction between organelles. 353
The potential role for membrane extensions seen emanating from, and in between, certain 354
organelles will also be discussed. It is important to note that whilst our understanding of 355
interactions in plants appears rudimentary at this stage compared with other model systems, the 356
development and application of novel biophysical tools (such as optical tweezers and pressure wave 357
technology) in plants will allow for a greater understanding of the regulation of interactions as the 358
molecular components are isolated. 359
360
ER-Golgi 361
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The secretory pathway begins with protein production in the ER and subsequent processing in the 362
Golgi. The functional relationship and observed close associations between these two organelles 363
lead to the hypothesis that physical association, be it through membrane continuity or large 364
proteineous tethering complex, must occur. Whilst trafficking of cargo between the two organelles is 365
cytoskeleton independent, movement of the two organelles are correlated and are cytoskeleton 366
dependent processes (see earlier, Brandizzi et al., 2002; Runions et al., 2006; Stefano et al., 2014). 367
To probe physical association between the two, optical tweezers attached to either a confocal 368
microscope or TIRF imaging system have been utilised. The principle behind optical tweezers is that 369
it uses a focused infrared beam which allows the user to physically trap and move an object which 370
has a different refractive index to the surrounding media (Sparkes et al., 2018; Sparkes, 2016). 371
Unlike the ER, Golgi bodies are amenable to being trapped in intact Arabidopsis leaf epidermal cells. 372
Physical association between the ER and Golgi would result in a trapped Golgi body dragging the ER. 373
If Golgi bodies were not physically associated with the ER, then the subsequent movement of a 374
trapped Golgi would separate it from the underlying ER. Qualitative assessment indicated that the 375
ER did, in fact, move with the Golgi and, on occasion, Golgi could be separated from the ER (Sparkes, 376
IA et al., 2009). More recently, semi-quantitative work has shown that CASP, a Golgi associated 377
protein, affects association between the two organelles (Osterrieder et al., 2017). These results, and 378
the application of optical tweezers, clearly highlight how we can determine rates of association 379
between organelles, and the relative strength of such through affecting the molecular components 380
involved in the tethering process. Future studies are required to elucidate components, in addition 381
to CASP, that are involved in the tethering of Golgi to the ER. Furthermore, application of a fully, 382
rather than partially, quantifiable optical trapping platform (Gao et al., 2016) will address the 383
relative importance of CASP and interactors in the tethering process (Sparkes et al., 2018). 384
385
Nucleus-chloroplast 386
Movement of nuclei and chloroplasts are controlled through short actin filament assembly at the 387
organelle surface. Tethering between nuclei and chloroplasts has been inferred through photo 388
induced migration of chloroplasts affecting the movement of associated nuclei (Higa et al., 2014). 389
Physical interaction between these two organelles is likely required for retrograde signalling. Elegant 390
signalling studies have shown that hydrogen peroxide levels in chloroplasts juxtaposed to the 391
nucleus alter in response to light, which ultimately changes nuclear expression of genes required for 392
photosynthesis (Exposito-Rodriguez et al., 2017). Identification and characterisation of molecular 393
tethers required for interaction between the nucleus and chloroplast will provide conclusive 394
evidence of physical association between the two compartments. 395
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396
ER-peroxisomes / oil bodies-peroxisomes 397
Peroxisome biogenesis is a contentious topic with two overarching proposed mechanisms; de novo 398
synthesis from the ER or fission from pre-existing peroxisomes. In plants there is very little evidence 399
for the former (Kao et al., 2018). Live cell imaging indicates that peroxisomes, and the resulting 400
membrane emanation peroxules, are closely aligned with the ER (Sinclair et al., 2009; Barton et al., 401
2013). It is unclear if this represents a direct physical association, or simply occupying the same actin 402
filament, or due to spatial constrictions imposed by the large central vacuole of cytoplasmic content, 403
in effect, pushing content together. 404
405
The ER is the site of production for oil bodies. Energy production through β-oxidation of the fatty 406
acids derived from oil bodies occurs in plant peroxisomes. Recent evidence has emerged indicating 407
close association between oil bodies and peroxisomes, which may be mediated by the retromer 408
complex (Thazar-Poulot et al., 2015). This retromer mediated interaction appeared to occur at 409
peroxisome membrane extensions (peroxules, see later). Changes in metabolic requirements 410
through provision of sucrose alters the frequency of oil body-peroxisome juxtaposition (Cui et al., 411
2016). In combination these results suggest that peroxisomes are associated with oil bodies 412
during β-oxidation, but it remains to be seen whether this interaction is direct or indirectly mediated 413
by an ER bridge. 414
415
Peroxisome-chloroplast 416
Peroxisomes and chloroplasts are functionally linked through the photorespiratory pathway, which 417
also spans mitochondria. Early micrographs again highlight close association between these 418
organelles. By altering the molecular components which drive chloroplast motion (chup1), 419
neighbouring peroxisomes appeared to move with chloroplasts as though physically attached 420
(Oikawa et al., 2003). Clear evidence for physical association came through optical tweezer (Fig 3) 421
and pressure wave studies (Oikawa et al., 2015; Gao et al., 2016). The latter generates a wave which 422
can affect, and effectively ‘push’ all subcellular components within the pressure wave radius, 423
whereas optical tweezers targets individual organelles with submicron accuracy. Both studies 424
indicated that peroxisomes are attached to chloroplasts, with pressure wave studies inferring that 425
interaction maybe affected by photosynthetic rate. Given that photosynthesis and photorespiration 426
are linked processes, it is unlikely that photosynthesis per se drives the organelle association and is 427
perhaps more likely a correlation. Isolating the molecular components which drive these interactions 428
will allow a deeper understanding of regulation and subsequent role of peroxisome-chloroplast 429
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interaction. Considering the functional repertoire of plant peroxisomes has expanded beyond earlier 430
concepts of lipid catabolism (β-oxidation), reactive oxygen species (ROS) production and scavenging , 431
it is not impossible to propose that novel functional roles may still exist requiring shuttling between 432
chloroplasts and peroxisomes (Kao et al., 2018). For example, β-oxidation is not only required for 433
energy mobilisation in germinating oilseeds, but also plays roles in production of hormones including 434
Jasmonic acid (JA). JA synthesis spans both peroxisomes and chloroplasts. Considering that 435
metabolism is a complex process, by controlling physical location of organelles we may be able to 436
alter metabolic balance. 437
438
ER-plasma membrane 439
Early work by Ridge et al. indicated static regions within the ER network (Ridge et al., 1999). More 440
recently, through the development of a quantitative platform to measure static elements within the 441
highly dynamic ER network (termed persistency mapping), static tubules, nodes and cisternal regions 442
were isolated which comprised ~5% of the ER network in tobacco leaf epidermal cell (Sparkes, I et 443
al., 2009). By observing the movement around these nodes, it appears as the though the ER network 444
‘wrapped’ itself and anchored in place to generate new geometric configurations of the network. 445
The premise of anchoring was again shown using optical tweezers; moving trapped Golgi bodies 446
around ER nodes to wrap and anchor the ER in place (Sparkes, IA et al., 2009). These anchor points 447
were in fact regions of the ER pinned to the plasma membrane. Plasmolysis studies result in the 448
plasma membrane being ‘pulled’ away from the cell wall to generate Hechtian strands (Wang et al., 449
2016). These strands highlight that the PM is fixed to the cell wall at junctions. The Hechtian strands 450
are filled with ER which reach up to the tip of the strand where the PM meets the cell wall (Wang et 451
al., 2016; Cheng et al. 2017). This arrangement is indicative of the ER being anchored to the PM, 452
which is fixed to the cell wall at discrete regions (Wang & Hussey, 2015). This arrangement could 453
enable fast transmission of signals between the extracellular and intracellular environments, and 454
also act to constrain and maintain an even distribution of the ER within the cell cortex (Wang & 455
Hussey, 2015; Bayer et al., 2017). Interestingly, attempts to model ER network formation based on 456
parameters including static and mobile ER node distribution shows a high correlation with formation 457
in vivo (Lin et al., 2014; Lin et al., 2017). As previously mentioned anchoring is through actin and 458
microtubules, and requires a complex of proteins including Net3, VAP27, synaptotagmin 71 and 459
Syp73 (Bayer et al., 2017; Wang et al., 2014, 2016, 2017; Wang & Hussey, 2017; Perez-Sancho et al., 460
2015; Cao et al., 2016). This complex could possibly affect the movement of other organelles, or act 461
as a functional nexus in the cell allowing organelles to ‘communicate’. Future studies to determine 462
how node formation is regulated, turned over and how the ER appears to randomly ‘hook’ and 463
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‘unhook’ itself from the PM will promote our understanding of the functional role of ER-PM 464
interactions. 465
466
467
Stromules, peroxules and matrixules: Tubular extensions as connections to other organelles? 468
Stromules, peroxules and matrixules are membrane protrusions containing luminal content 469
emanating from chloroplasts, peroxisomes and mitochondria respectively. Several hypothesis have 470
been proposed for the functional role of such extensions; to increase surface area, to allow 471
organelles to communicate and exchange components at the tip of the extension, or / and to anchor 472
organelles to one another. These hypothesis are not mutually exclusive. For example communication 473
could occur at MCS’ between two organelles, if the organelles start to move at differing rates then a 474
physical manifestation of maintaining contact would be formation of extensions (Figure 4). 475
Interestingly, peroxule formation occurs upon trapping and pulling peroxisomes away from 476
chloroplasts (Gao et al., 2016) (Fig 3). Quantification of this event indicated that peroxules at the 477
chloroplast-peroxisome contact site are anchored with a degree of tension being induced. Modelling 478
of the forces required to ‘pull’ peroxisomes back to the chloroplast gives an indication of the force 479
required by myosins to pull peroxisomes away from chloroplasts in vivo. 480
481
Evidence for transfer occurring at the tips of extensions is contentious, and is based on studies of 482
exchange between stromules from neighbouring plastids (Hanson & Sattarzadeh, 2013; Mathur et 483
al., 2013). Membrane extensions also form in response to other stimuli including ROS and is 484
reviewed in (Mathur et al., 2012). More recently stromule formation has been linked with pathogen 485
infection (Caplan et al., 2015). 486
487
Stromules and peroxules appear to co-align with the ER network, leading to the proposal that they 488
are physically attached to the ER network (Schattat et al., 2011; Barton et al., 2013). Peroxules are 489
single extensions unlike stromules which can form branched networks. The ER is pervasive 490
throughout the cortex, organelles in highly vacuolated cells are constrained and can appear to 491
neighbour one another at random, and movement of both the ER and these organelles are largely 492
driven by acto-myosin dependent processes. These observations make it difficult to determine 493
whether there is a true physical association between the ER and these other organelles, or whether 494
they are ‘connected’ through association to the same actin filament. Future studies are required to 495
reconcile which of these scenarios is correct. 496
497
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Interestingly, a biochemical complementation approach highlighted that the ER and chloroplasts can 498
exchange nonpolar metabolites (Mehrshahi et al., 2013). This exchange was suggested to likely occur 499
at closely opposed membrane sites. Photostimulation studies highlighted a potential interaction site 500
between the two organelles which may affect movement of the ER network and protein within 501
(Griffing L, 2011). Optical tweezer studies in laser ablated cells indicated a physical association 502
between the ER and chloroplasts (Andersson et al., 2007). Future studies to pinpoint molecular 503
tethering components will determine if there is a true physical association between these two 504
organelles. 505
506
Summary model to describe organelle movement in higher plants 507
To take into account the level of evidence for physical interaction between organelles and the role of 508
actomyosin system on movement, Figure 4 proposes how movement of tethered organelles is 509
coordinated and regulated. Here, movement of discrete untethered organelles are depicted as being 510
controlled independently of one another (Fig 4a). This could be through specific myosin-organelle 511
interaction mediated by myosin recruitment factors (receptors, adaptors previously mentioned) and 512
relates to Fig 2b. For organelles tethered to one another (Fig 4b-d), movement could be regulated by 513
one of the two organelles in turn pulling the other (Fig 4b, organelle A pulls tethered organelle B), or 514
movement of the two tethered organelles could be co-regulated by coordinated action of myosins 515
on each organelle (Fig 4c). Here, co-regulation could move the two organelles in the same direction 516
through interaction of organelle specific myosins binding and traversing different F-actin strands. In 517
addition, it is interesting to point out that if one organelle moves faster than the other, or changes 518
direction of motion, here it may result in changes in morphology of the tethered organelles resulting 519
in membrane extensions such as peroxules, matrixules and stromules. Evidence supporting this 520
could be drawn from increased peroxisome speeds under ROS stress which also induces peroxule 521
formation; are these organelles simply trying to maintain contact with other organelles? Given that 522
the ER may be involved in multiple organelle interactions then here movement of organelles could 523
be mediated through a three way organelle interaction complex; organelle A moves which causes ER 524
movement which in turn affects organelle B’s movement (Fig 4d). In addition, organelle movement 525
could possibly be driven through direct interaction with actin, with actin polymerisation itself driving 526
organelle movement. For example, proteins such as Net3 and Syp73 connect the ER to actin. These 527
models do not include cytoplasmic streaming, an additional complication which needs to be taken 528
into account to aid our understanding of the local and global regulation of organelle movement and 529
interaction. Could streaming increase the chance of organelle collisions which then ‘stick’ to one 530
another if they contain the correct tethering components? Could the scenario in Fig 4d explain 531
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cytoplasmic streaming; movement of organelle A and B controls ER movement which in turn 532
generates viscous drag to entrain the cytoplasmic flow? To begin to unpick which, if any, of these 533
suggested models occurs will require further understanding of the tethers involved in allowing 534
interaction of organelles and how this impacts on organelle movement and positioning. 535
536
537
Why is organelle movement important? 538
539
Recent advances suggest that organelle movement plays a fundamental role in plant growth and 540
development and in response to biotic and abiotic stresses/stimuli. Readers are directed towards 541
recent reviews covering this topic in more depth (Duan & Tominaga, 2018; Ryan & Nebenführ, 542
2018). In brief, here, myosin mutants with reduced organelle movement can display a reduction in 543
aerial tissue growth and therefore biomass, altered flowering timing and reduced seed set. These 544
gross morphological phenotypes are due to changes in cell size rather than cell number, and as 545
shown in root hairs, reduced polar tip growth (Peremyslov, V. V. et al., 2008; Prokhnevsky et al., 546
2008; Peremyslov et al., 2010). This is also seen in pollen tubes and so may reflect reduced 547
fertilisation affecting seed set (Madison et al., 2015). In addition, there are also alterations in 548
trichome growth (Ojangu et al., 2007; Ojangu et al., 2012). 549
Key players in organelle movement and actin dynamics include myosins XI-K, XI-1 and XI-2. 550
Considering that these myosins have also been implicated in cytoplasmic streaming, could these 551
phenotypes be a direct result of alterations in streaming rates rather than directed organelle 552
movement per se? Here, the hypothesis would be that streaming mixes the cytoplasm and that 553
organelle movement is passive in the process. Work presented by Tominaga et al. tested this 554
hypothesis by generating transgenic Arabidopsis expressing XI-2 chimeras which either had a higher 555
or slower ‘speed’ (Tominaga et al., 2013). Here, they found that higher ‘speed’ motor resulted in 556
faster growth compared with plants expressing the ‘lower’ speed variant, and that growth rates 557
related to cell size. They concluded that cytoplasmic streaming is therefore a determinant of cell 558
size. While these results show a clear correlation between streaming and cell size, it does not negate 559
that size may be due to effecting the movement of the organelles rather than mixing of the 560
cytoplasm. It is difficult to reconcile the relative contributions of these two processes (cytoplasmic 561
mixing versus organelle movement) on cell growth given our current molecular tools. Specifically 562
targeting and affecting the movement of individual classes of organelles will begin to unpick these 563
processes and relative impacts on cell growth. Clear evidence, however, for the role of chloroplast 564
and nuclear movement has been elucidated. 565
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566
Both chloroplasts and nuclei positioning changes in response to light, a process called 567
photorelocation (Tsuboi et al., 2007; Kodama et al., 2008; Wada, 2013; Wada, 2016). Chloroplasts 568
locate to the periclinal walls of mesophyll cells of A.thaliana under low blue light intensities 569
(accumulation response) in order to maximise light capture and, therefore, photosynthesis. They 570
then migrate to anticlinal walls (avoidance response) in conditions of high blue light intensities in 571
order to avoid photodamage (Kasahara et al., 2002). Phototropin1 (phot1) and Phototropin2 (phot2) 572
are the blue light receptors involved in perceiving the intensity of blue light: both phot1 and phot2 573
are involved in the accumulation response (Sakai et al., 2001), whereas only phot2 is involved in the 574
avoidance response (Kagawa et al., 2001). Interestingly, phot2 is also involved in nuclei blue light 575
avoidance response (Iwabuchi et al., 2010). 576
The position that organelles occupy in the cell also appears to be finely regulated. During root hair 577
growth, for instance, the nucleus moves forward maintaining the same distance from the tip 578
(Chytilova et al., 2000; Ketelaar et al., 2002), until the growth process is complete. Drug treatments 579
with actin depolymerising drugs and immobilisation of the nucleus by optical trapping (Ketelaar et 580
al., 2002), suggest that correct positioning of the nucleus from the growing tip is essential. Under 581
certain growth conditions, wild type plants can produce and extend two root hairs from the same 582
single trichoblast cell. These root hairs grow and elongate in a coordinated manner albeit with a 583
slight lag in growth of the nuclear ‘free’ root hair (Jones & Smirnoff, 2006). These events are rare, 584
and question whether nuclear positioning is completely essential for root hair elongation. 585
586
587
Conclusions and future perspectives 588
589
We have provided an overview of the molecular mechanisms underpinning organelle movement and 590
positioning in plants; largely actomyosin dependent and interaction between organelles. Whilst 591
there appears to be a degree of functional redundancy between the myosin class XI family, it should 592
be noted that there appears to be a certain degree of specific control over ER dynamics (Griffing et 593
al., 2014). Could a similar level of control occur within subpopulations of discrete organelles such as 594
Golgi, peroxisomes and mitochondria? When observing movement of these organelles it is usual to 595
monitor a fluorophore targeted to the organelle through a targeting sequence which doesn’t 596
necessarily represent a fully functional native protein. The population of a certain class of organelle 597
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19
therefore appears homogenous. In reality, could organelle groups be more heterogenous, and could 598
this heterogeneity reflect the movement characteristics of the organelle itself? For example, a cargo 599
laden organelle may need to move faster to exchange and release cargo unlike another which has a 600
lower cargo load, or a different cargo load. Future studies into organelle heterogeneity and how this 601
regulates movement will unveil this exciting possibility that organelles regulate their own movement 602
patterns through signalling cascades. This has already begun to occur through studies of the effects 603
of ROS on organelle dynamics. To be able to pull out patterns of movement, and relate them to a 604
functional response will therefore require development and application of novel sensors (such as 605
ratiometric ATP probe (De Col et al., 2017), novel applications of imaging techniques (such as optical 606
tweezers for testing and quantifying organelle interactions), and mathematical and biophysical 607
models. The latter will provide predictive power to determine which parts of the system provide the 608
ultimate control over organelle dynamics. For example, models of ER network formation have 609
provided a first principle approximation of the biophysical properties required to form a dynamic 610
model of network formation which fits in vivo ER network dynamics (Lemarchand et al., 2014; Lin et 611
al., 2015; Griffing et al., 2017; Lin et al., 2017). 612
613
614
Acknowledgements 615
Owing to space limitations we apologise to those whose work we could not include herein. IS and CP 616
are supported by the Gatsby foundation and the Leverhulme Trust. Some of the work reported 617
herein was supported by the BBSRC and STFC. 618
619
Authors contributions 620
621
Both authors contributed equally to the work 622
623
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958
Figure legends 959
960
Figure 1. Time lapse images showing ER and Golgi dynamics in A. thaliana leaf epidermal cell. 961
The position of Golgi (magenta) and ER (green) at timepoints 0s (a), 15s (b) and 30s (c) are indicated. 962
Blue (e, i), yellow (f, j) and magenta (g,k) panels indicate images of ER (i, j, k) and Golgi (e,f,g) taken 963
at timepoints of 0, 15 and 30 seconds respectively. Overlays of all the timepoints for ER (l) and Golgi 964
(h) indicate regions of slow movement (white) versus highly dynamic movement. White arrow (h) 965
indicates a static Golgi body. White box in panel h is magnified in panel d. Magenta outline of Golgi 966
bodies (d) indicates 0s position and subsequent movement is shown by coloured line tracks . Scale 967
bar 5 µm except for panel (d) which is 1µm Image capture rate was 0.4 frames per second. 968
969
Figure 2. Schematic highlighting myosin recruitment and organelle movement 970
Diagram depicting XI-I recruitment and interaction with components at the outer and inner nuclear 971
envelope (a). Scheme adapted from Tamura et al. (2013). 972
Possible roles for the MyoB/MadA-B decorated vesicles in myosin recruitment and organelle 973
dynamics (b,c,d). MyoB/MadA-B proteins could be located on organelle membranes and mediate 974
the recruitment of specific myosins to the organelle membrane. Said organelle then moves actively 975
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along the actin cytoskeleton (b). MyoB vesicles have been shown to move faster than any other 976
organelle. Therefore, as suggested by Peremyslov et al. (2015), MyoB/MadA-B decorated vesicles 977
could be the only structures actively moving on the actin through myosin motors, and able to 978
generate cytoplasmic streaming. Other organelles possibly follow the flow passively (c). MyoB 979
decorated vesicles might be located in proximity, and interact, with certain sub-domains of known 980
organelles, dragging the organelle along the actin filaments in a myosin dependent way (d). Given 981
the size of MyoB and myosin gene families, models are not necessarily mutually exclusive and so a 982
more complex picture of coexistence of several of these options could occur (b, c, d). 983
984
985
Figure 3. Peroxisome-chloroplast interaction in plants 986
Peroxisome (p) juxtaposed to a chloroplast (cp) are depicted prior to optical trapping (a). Peroxisome 987
is trapped and pulled away from the chloroplast (b) inducing a peroxule (white arrowhead) which 988
appears to maintain contact between the two organelles. Upon releasing the peroxisome from the 989
trap (c) it ‘springs’ back towards its original position (a) along with a resultant retraction in the 990
peroxule. Peroxules therefore appear to be membrane manifestations of membrane contact sites 991
between the two organelles. Scale bar 5µm. 992
993
Figure 4. Models proposed for the coordinated action of myosin dependent processes and 994
organelle tethering on subsequent organelle movement and positioning. 995
Organelles may not be tethered and will therefore recruit myosins and move independently from 996
one another in the cell cytoplasm (a). Note, this links to Figure 2. In the case where organelle A is 997
tethered to organelle B (red and black hooks) movement could be controlled via the mechanisms 998
depicted in panels b-d. Organelle A’s movement is myosin dependent which in turn drags tethered 999
organelle B; organelle B is therefore ‘piggybacking’ on organelle A (b). In (c) and (d) both tethered 1000
organelles recruit specific myosins on their membrane and the direction in which the organelles will 1001
move depends on the polarity of actin, and on the resulting forces derived by myosins moving along 1002
the actin filaments. Here, coordinated regulation of myosins on organelle A and B is required to 1003
allow simultaneous movement of both organelles to maintain tethering (c). Tethering between 1004
organelles may involve multiple organelles rather than just two organelles (d). Here, similar to that 1005
depicted in (c), movement is coordinated between three organelles rather than just two. There is 1006
speculation that the ER may act as a bridge between certain organelles. If the rate of movement of 1007
tethered organelles is not coordinated (whereby one moves faster than the other or they move in 1008
opposite directions, as indicated by red arrows in panels c and d), then contacts maybe maintained 1009
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through membrane extensions such as peroxules, stromules and matrixules. Note, upper actin 1010
filament in c and d reflects potential for either orientation of filament polarity, with arrows for 1011
organelle movement referring to plus end directed motion. 1012
1013
1014
1015
1016
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Figure 1. Time lapse images showing ER and Golgi dynamics in A. thaliana leaf epidermal cell. The position of Golgi (magenta) and ER (green) at timepoints 0s (a), 15s (b) and 30s (c) are indicated. Blue
(e, i), yellow (f, j) and magenta (g,k) panels indicate images of ER (i, j, k) and Golgi (e,f,g) taken at timepoints of 0, 15 and 30 seconds respectively. Overlays of all the timepoints for ER (l) and Golgi (h)
indicate regions of slow movement (white) versus highly dynamic movement. White arrow (h) indicates a static Golgi body. White box in panel h is magnified in panel d. Magenta outline of Golgi bodies (d) indicates 0s position and subsequent movement is shown by coloured line tracks . Scale bar 5 µm except for panel (d)
which is 1µm Image capture rate was 0.4 frames per second.
199x155mm (300 x 300 DPI)
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Figure 2. Schematic highlighting myosin recruitment and organelle movement Diagram depicting XI-I recruitment and interaction with components at the outer and inner nuclear envelope
(a). Scheme adapted from Tamura et al. (2013).
Possible roles for the MyoB/MadA-B decorated vesicles in myosin recruitment and organelle dynamics (b,c,d). MyoB/MadA-B proteins could be located on organelle membranes and mediate the recruitment of
specific myosins to the organelle membrane. Said organelle then moves actively along the actin cytoskeleton (b). MyoB vesicles have been shown to move faster than any other organelle. Therefore, as suggested by Peremyslov et al. (2015), MyoB/MadA-B decorated vesicles could be the only structures
actively moving on the actin through myosin motors, and able to generate cytoplasmic streaming. Other organelles possibly follow the flow passively (c). MyoB decorated vesicles might be located in proximity, and interact, with certain sub-domains of known organelles, dragging the organelle along the actin filaments in a myosin dependent way (d). Given the size of MyoB and myosin gene families, models are not necessarily
mutually exclusive and so a more complex picture of coexistence of several of these options could occur (b, c, d).
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Figure 3. Peroxisome-chloroplast interaction in plants Peroxisome (p) juxtaposed to a chloroplast (cp) are depicted prior to optical trapping (a). Peroxisome is
trapped and pulled away from the chloroplast (b) inducing a peroxule (white arrowhead) which appears to
maintain contact between the two organelles. Upon releasing the peroxisome from the trap (c) it ‘springs’ back towards its original position (a) along with a resultant retraction in the peroxule. Peroxules therefore appear to be membrane manifestations of membrane contact sites between the two organelles. Scale bar
5microns.
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Figure 4. Models proposed for the coordinated action of myosin dependent processes and organelle tethering on subsequent organelle movement and positioning.
Organelles may not be tethered and will therefore recruit myosins and move independently from one
another in the cell cytoplasm (a). Note, this links to Figure 2. In the case where organelle A is tethered to organelle B (red and black hooks) movement could be controlled via the mechanisms depicted in panels b-d.
Organelle A’s movement is myosin dependent which in turn drags tethered organelle B; organelle B is therefore ‘piggybacking’ on organelle A (b). In (c) and (d) both tethered organelles recruit specific myosins on their membrane and the direction in which the organelles will move depends on the polarity of actin, and on the resulting forces derived by myosins moving along the actin filaments. Here, coordinated regulation of
myosins on organelle A and B is required to allow simultaneous movement of both organelles to maintain tethering (c). Tethering between organelles may involve multiple organelles rather than just two organelles (d). Here, similar to that depicted in (c), movement is coordinated between three organelles rather than just
two. There is speculation that the ER may act as a bridge between certain organelles. If the rate of movement of tethered organelles is not coordinated (whereby one moves faster than the other or they move
in opposite directions, as indicated by red arrows in panels c and d), then contacts maybe maintained through membrane extensions such as peroxules, stromules and matrixules. Note, upper actin filament in c
and d reflects potential for either orientation of filament polarity, with arrows for organelle movement referring to plus end directed motion.
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