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The Plant Cell, Vol. 21: 3937–3949, December 2009, www.plantcell.org ã 2009 American Society of Plant Biologists
Movement and Remodeling of the Endoplasmic Reticulum in
Nondividing Cells of Tobacco Leaves
W
I. Sparkes,a J. Runions,a C. Hawes,a and L. Griffingb,1
a School
b Biology
of Life Sciences, Oxford Brookes University, Oxford OX3 0BP, United Kingdom
Department, Texas A&M University, College Station, Texas 77843-3258
Using a novel analytical tool, this study investigates the relative roles of actin, microtubules, myosin, and Golgi bodies on
form and movement of the endoplasmic reticulum (ER) in tobacco (Nicotiana tabacum) leaf epidermal cells. Expression of a
subset of truncated class XI myosins, which interfere with the activity of native class XI myosins, and drug-induced actin
depolymerization produce a more persistent network of ER tubules and larger persistent cisternae. The treatments
differentially affect two persistent size classes of cortical ER cisternae, those >0.3 mm2 and those smaller, called punctae.
The punctae are not Golgi, and ER remodeling occurs in the absence of Golgi bodies. The treatments diminish the mobile
fraction of ER membrane proteins but not the diffusive flow of mobile membrane proteins. The results support a model
whereby ER network remodeling is coupled to the directionality but not the magnitude of membrane surface flow, and the
punctae are network nodes that act as foci of actin polymerization, regulating network remodeling through exploratory
tubule growth and myosin-mediated shrinkage.
INTRODUCTION
The endoplasmic reticulum (ER) is the port of entry into the
secretory pathway, yet very little is understood about the dynamic nature of this pleomorphic structure (Sparkes et al.,
2009a). Actomyosin-based directional movement and cytoplasmic streaming drives the movement and dispersion of endomembrane compartments, such as the Golgi apparatus in plants
(Avisar et al., 2008, 2009; Peremyslov et al., 2008; Prokhnevsky
et al., 2008; Sparkes et al., 2008). Plant myosins, especially class
XI myosins, including XIK, motorize Golgi bodies (and mitochondria and peroxisomes) just as myosin-V motorizes endomembrane organelles in yeast (Estrada et al., 2003) and nerve cells
(Brown et al., 2004). While expression of the tail domain of certain
Arabidopsis thaliana class XI myosins perturb organelle movement, these fluorescent fusions do not collocate to the organelles
whose movement they effect, thus indicating titration of accessory factors or low levels of organelle binding. Here, analytical
software has been developed to address questions of whether
the increased movement and dispersion that characterize plant
organelles, relative to organelles of animal cells, also pertain to
the ER and how this movement and dispersion is driven by the
actomyosin system.
As in cultured animal cells and yeast, the ER of higher plants is
made of two primary domains, a polygonal tubule network, and
cisternae (flattened membrane saccules) that are coextensive
with the network. The movement of the network is not an entire
1 Address
correspondence to [email protected].
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy described
in the Instructions for Authors (www.plantcell.org) is: L. Griffing
([email protected]).
W
Online version contains Web-only data.
www.plantcell.org/cgi/doi/10.1105/tpc.109.072249
shifting of a stationary set of polygons relative to, for instance, the
plasma membrane. Indeed, the network appears more anchored
to the plasma membrane than are other organelles (Quader et al.,
1987). In the absence of global shifting of the network, movement
within the network is either the movement of persistent network
structures or the remodeling of the network through the formation or movement of network nodes and branches (Wolfram,
2002). Therefore, movement of the ER can be assessed by
determining the persistency of network structures, such as
nodes and branches. Here, this is done with persistency maps,
images that show the level of persistency of different network
structures in the ER, including tubules (branches and polygons),
nodes (punctae or cisternae with an area between 0.1 and 0.3
mm2), and cisternae (membrane sheets with an area more than
0.3 mm2).
ER movement includes movement of proteins within the
ER membrane and network remodeling. Directional, actindependent movement of membrane protein in the ER has recently been demonstrated in plants (Runions et al., 2006). The
remodeling of the cisternal domain into tubules through the
recruitment of reticulons, proteins that drive tubulation, may lead
to the dynamic transition between tubular and cisternal domains
(Tolley et al., 2008). In yeast (De Craene et al., 2006) and
mammalian cells (Voeltz et al., 2006), the nuclear envelope is
the most reliably visible cisternal domain of the ER. In plants,
tubular-cisternal transitions are developmental (Ridge et al.,
1999) and may generally represent regions of differing ER function (Staehelin, 1997).
The plant ER network overlies the actin cytoskeleton (Boevink
et al., 1998), but does the actomyosin system regulate the forms
of movement described above? Here, changes in the persistency of the cortical ER of tobacco (Nicotiana tabacum) leaf
epidermal cells are determined after treatments that depolymerize actin filaments (latrunculin b) or microtubules (oryzalin),
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perturb organelle motility (coexpression of a truncated class XI
myosin) or remove Golgi bodies (Brefeldin A). Although the
results indicate that some form of linkage between the actin
cytoskeleton and the ER membrane might drive ER movement
and change the mobile fraction of ER surface proteins, they do
not support the notion that the Golgi bodies, which are intimately
associated with the ER (Boevink et al., 1998; Runions et al., 2006;
Sparkes et al., 2009b) and which are also driven by the actomyosin system (Boevink et al., 1998; Nebenführ et al., 1999),
mediate ER movement.
RESULTS
ER Network Persistence
Image maps showing the persistence of tubules and cisternae
are called persistency maps. Two different control tobacco
epidermal cells, representative of 30 such cells, expressing the
fluorescent marker green fluorescent protein (GFP)-HDEL (see
Supplemental Movie 1 online) are shown in Figure 1, which
includes the first and last frames of a 50-frame movie (Figures 1A,
1A’, 1B, and 1B’), the total imaged membrane over the time of
imaging (image sum) used for normalization (Figures 1C and 1C’),
and tubule (Figures 1D, 1D’, 1F, and 1F’) and cisternal persistency maps (Figures 1G, 1G’, 1I, and 1I’). Persistence is displayed as a gray level gradient (Figures 1D, 1D’, 1G, and 1G’) or
discrete gray levels (Figures 1E, 1E’, 1H, and 1H’). The darkest,
most persistent tubules above a certain size (Figures 1F and 1F’)
are quantified (see below). Likewise, the most persistent cisternal
structures, punctae that are smaller than 0.3 mm2 and cisternae
that are larger than 0.3 mm2, are quantified (see below). Tubule
and cisternal persistence are lower in some cells (Figures 1A’ to
1I’) than in others (Figures 1A to 1I). Both contain fast lanes of ER
movement with low persistence gray regions adjacent to the
arrow (Figures 1E and 1E’), where tubules change rapidly and
active streaming occurs in the direction indicated. Neither example has persistent polygons within the network, nor do less
persistent polygonal structures move (no gray tubules remain as
polygons in Figures 1E and 1E’). There are few persistent
cisternae, but less persistent ones (dark gray, particularly punctae, Figures 1H and 1H’) move. Therefore, as expected, and thus
validating the modeling system, the movement of the network is
not movement of polygonal subdomains of the network but of
small nodal punctae and tubule branches that form, grow, and
shrink or generate cisternae.
Actomyosin Requirement for ER Network Remodeling
To ascertain the contribution of the cytoskeleton to ER remodeling, tobacco epidermal cells were treated with drugs to depolymerize either actin filaments or microtubules or subjected to
overexpression of class XI myosin tail domains, XIK or XIJ, fused
to enhanced yellow fluorescent protein (eYFP). To confirm that
latrunculin b and oryzalin depolymerize actin filaments and
microtubules, respectively, pieces of tobacco leaf epidermal
cells expressing fluorescent markers for both were treated and
are shown in Supplemental Figure 1 online. Previous studies
have shown that XIK tail fusions perturb Golgi, peroxisome, and
mitochondria movement (Sparkes et al., 2008). Proposed explanations for these effects include competition for myosin effectors
and cargo binding sites; however, since little is known about the
latter, the formal proof for the mode of action of plant myosin tail
domains remains elusive (Sparkes et al., 2008; Avisar et al.,
2009). This fusion, along with other class XI tail domain fusions,
does not collocate to these organelles, is present in motile
puncta, and is diffuse throughout the cytosol. This diffuse cytoplasmic distribution could reflect a low level of binding to their
target organelles. Similarly, the tail domain of XIJ fused to eYFP is
diffuse throughout the cytosol but has a negligible affect on Golgi
movement (Avisar et al., 2009). Therefore, as a control for XIK
specificity on ER modeling, the effects of the tail domain of XIJ
fused to eYFP were tested in parallel.
ER network movement is quite different when cells coexpress
the myosin XIK tail domain (Figure 2; see Supplemental Movie 2
online, representative of 27 cells) or are treated with latrunculin b
(Figure 3; see Supplemental Movie 3 online, representative of 14
cells). The pattern of tubules and cisternae changes less (Figures
2, 3A, and 3B; see Supplemental Movies 2 and 3 online) than in
controls (Figures 1A, 1B, 1A’, and 1B’; see Supplemental Movie
1 online). The tubule persistency maps of the treatments (Figures
2D to 2Fand 3D to 3F) show a connected network of persistent
tubules that include polygonal rings not found in control cells
(Figures 1D to 1F and 1D’ to 1F’). The most persistent tubules
(Figures 2E, 2F, 3E, and 3F) are longer and more abundant
throughout the network than in control cells (Figures 1E, 1F, 1E’,
and 1F’). The two treatments produce more and longer persistent
tubules than control (Figure 4A). However, there is little difference
in persistent tubules between the latrunculin b treatment and
myosin XIK tail coexpression (Figure 4A). Both treatments produce more persistent cisternae (Figures 2G to 2I and 3G to 3I)
than found in the controls (Figures 1G to 1I and 1G’ to 1I’).
Although there is little difference between the number of persistent punctae per 100 mm2 of imaged membrane in the control
and cells coexpressing truncated myosin XIK, there is a significant increase in punctae in the latrunculin b treatment compared
with the control (Figure 4B).
After myosin XIK tail coexpression, high persistency punctae
are near the periphery of the larger persistent and less persistent
cisternae (arrows, Figure 2I), whereas the lower persistency
punctae subtend tubular networks (arrows, Figure 2H). Many
punctae seen in the latrunculin b treatment are not so much nodal
at the branching or bending points of tubules, as they are on the
periphery of the large cisternae (arrows, Figures 3A, 3E, and 3I).
Less persistent punctae sometimes appear at the blind end of
tubules (arrowheads, Figures 3E and 3H). Persistent tubules in
control cells terminate into a network of lower persistency
tubules (Figures 1E and 1E’), whereas short persistent tubules
in myosin XIK tail coexpression and latrunculin b treatments do
not (Figures 2E and 3E).
Cisternae increase in both area and number after truncated
myosin XIK coexpression and latrunculin b treatment (Figure 4C;
compare Figures 1G and 1G’ with 2G and 3G). Although both
treatments show a similar increase in the average area of the
cisternae over control, there is a larger increase in the number of
Cytoskeletal Control of Plant Endoplasmic Reticulum
3939
persistent cisternae after latrunculin b treatment than after truncated myosin XIK coexpression (Figure 4C).
To ascertain whether microtubules and another class XI myosin (XIJ) affect ER remodeling, movies were captured of cells
coexpressing eYFP-XIJ tail domain and GFP-HDEL and after
oryzalin treatment of cells expressing GFP-HDEL (see Supplemental Figures 2 and 3 and Supplemental Movies 4 and 5 online).
Oryzalin has no significant effect on tubule length and persistency (Figure 4A) but does slightly increase persistent cisternae
size compared with the control (Figure 4C; see Supplemental
Figure 2 online for persistency maps). Prior to quantification, it
was apparent that coexpression of the XIJ tail domain did not
perturb ER remodeling. A small sample size was analyzed and
indicated that the XIJ tail domain did not affect ER cisternal size
or number, but there were no persistent tubules above the
threshold level of 0.3mm2 (see Supplemental Figure 3 online for
persistency maps).
Actomyosin Requirement for ER Membrane Protein Mobility
Figure 1. Persistency of Tubules, Punctae, and Cisternae of Cortical ER
in Two Different Control Tobacco Epidermal Cells Expressing GFPHDEL.
The first ([A] and [A’]) and last frame ([B] and [B’]) of a 50-frame 80-s
movie. Summed frames of the entire movie ([C] and [C’]), showing total
imaged membrane (gray plus black) and the brighter punctae in the
summed image (black). The tubule persistency map ([D] and [D’]) shows
more persistent tubules as darker values. The posterized gray-level
tubule persistency map ([E] and [E’]) shows high persistency (black),
mid-range persistency (dark gray), and low persistency (light-gray gradient). The black arrow is positioned adjacent to an area of active
streaming, showing the direction of streaming. High persistency tubules
([F] and [F’]) from (E) and (E’) that were >0.3 mm2 in area are the tubules
counted in Figure 4A. In (F), the arrow indicates a persistent tubule that is
subtended by persistent punctae ([I], below). The persistency map ([G]
Fluorescence recovery after photobleaching (FRAP) and photoactivation studies reveal the dynamics of ER membrane proteins
after disruption of the actomyosin system. Overexpression of
GFP-CXN generates large ER cisternae over time (Figure 5A).
Cisternalization provides large two-dimensional sheets desirable
for FRAP photobleaching. FRAP analysis of cisternal GFP-CXN
yielded rates (t1/2 values) of protein movement and a measure of
the mobile fraction of membrane protein. Control cells (Figures
5A to 5C), latrunculin b–treated cells (Figures 5D to 5F), and cells
coexpressing GFP-CXN and the eYFP-XIK tail domain (Figures
5G to 5I) had similar recovery rates (t1/2 = ;2.4 s; Table 1, Figure
5L). However, coexpression of GFP-CXN and the eYFP-XIK tail
domain resulted in a significantly lower mobile fraction of fluorescent protein, for example, exchange of bleached molecules
with fluorescent molecules outside of the spot bleach region
(79.09% 6 13.20%, Tukey’s HSD P < 0.001) compared with
control and latrunculin b treatments, which were the same
(87.72% 6 10.18% and 89.14% 6 7.39%, respectively; Table
1, Figure 5L).
Photoactivation studies using CXN-paGFP (Runions et al.,
2006) measured protein mobility and mobile fraction independently of FRAP. Protein mobility is measured as fluorescence
intensity decline as photoactivated CXN-paGFP disperses from
the activation spot (Figure 6). The t1/2 values of ;2 s for
movement away from the activation spot were the same in all
cases (Figures 6A to 6K; see also Table 2). Mobile fractions,
and [G’]) shows more persistent cisternae and punctae as darker values.
The posterized gray-level cisternal persistency map ([H] and [H’]) shows
high persistency (black), mid-range persistency (dark gray), and low
persistency (light gray gradient) cisternae and punctae (arrowheads [H’]
show midrange persistency punctae). The black arrow is positioned
adjacent to an area of active streaming, showing the direction of
streaming. The high persistency map ([I] and [I’]) from (G) and (G’)
shows both the punctae (0.1 to 0.3 mm2, circularity > 0.5) and cisternae
(0.3 mm2). Arrowheads show persistent punctae connected by a persistent tubule. Bar = 10 mm.
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coexpressing CXN-paGFP and ST-mRFP after treatment with
BFA for 3 h reveals t1/2 values for movement away from the
activation spot (1.79 6 0.98 s) similar to control (2.01 6 1.15 s).
The mobile fraction (91.88% 6 8.08%) was significantly lower
after BFA than in control samples (97.63% 6 4.26%, Tukey’s
HSD P = 0.001; Table 2, Figures 6I to 6K).
DISCUSSION
Visualization of Cortical ER Network Movement and
Cytoplasmic Streaming
Figure 2. Persistency of Tubules, Punctae, and Cisternae of Cortical ER
in a Tobacco Epidermal Cell Coexpressing GFP-HDEL and the eYFP-XIK
Tail Domain.
(A) to (I) Same as in Figure 1. Because there are fewer or no fast lanes in
cells coexpressing myosin-XIK tails and treated with latrunculin b, the
gray regions in (E) indicate regions of remodeling cisternae that are
differentially skeletonized. Arrows in (H) indicate less persistent punctae
associated with tubules. Arrows in (I) indicate more persistent punctae
associated with cisternae. Bar = 10 mm.
The nature of cortical ER network movement in plant cells is not
widely understood. In higher plants, the ER cisternal/tubular
network does not move as a whole or as cisternal/tubular
network fragments, as is often illustrated for the involvement of
ER in charophyte cytoplasmic streaming (Kachar and Reese,
1988; Lodish et al., 2008). Instead, about 5% of the total imaged
ER is very persistent (Figure 4A) and forms the framework upon
which movement occurs. This framework is revealed with persistency maps, visual maps of the ER that show its recent degree
of persistency. The maps show ER features that remain relatively
static for periods of longer than 8 s (five time-lapse frames) in an
80-s data set (Figures 1D, 1D’, 1I, and 1I’; see Supplemental
Movie 1 online). These persistency maps do not have persistently
connected networks of polygonal rings (Figures 1D, 1D’, 1E, and
1E’); polygonal subunits do not persist. Polygon ring formation
however, of cells coexpressing the mRFP-XIK tail domain
(91.27% 6 6.12%) and those treated with latrunculin b
(89.42% 6 7.04%) were significantly lower than in control cells
(97.63% 6 4.26%, Tukey’s HSD, P < 0.001; Figure 6L).
Are Golgi Bodies Involved in ER Remodeling and ER
Membrane Mobility?
In various plant tissues, Golgi bodies appear to be associated
with the ER and move either over, or with the ER membrane, but
as seen here do not collocate with persistent cortical ER puncta
(Figures 7A to 7C).
To test the putative role of Golgi bodies in ER remodeling and
membrane flow, cells were treated with Brefeldin A (BFA), which
deconstructs Golgi and results in varying degrees of cisternalization of the ER (Boevink et al., 1999). This cisternalization
precludes comparison of tubule and cisternal persistency with
control cells because drug effects could be confused with the
direct effect of Golgi body absence.
Cells coexpressing GFP-HDEL and the Golgi marker STmRFP were treated with BFA for 3 h, resulting in disappearance
of Golgi bodies and redistribution of ST-mRFP to the ER (Figures
7D to 7F). Stills from a representative movie (Figures 7G to 7I; see
Supplemental Movie 6 online) show that the ER remodels in the
absence of Golgi bodies. Photoactivation analysis of cells
Figure 3. Persistency of Tubules, Punctae, and Cisternae of Cortical ER
in a Tobacco Epidermal Cell Expressing GFP-HDEL and Treated with
Latrunculin b.
(A) to (I) Same as in Figure 1. Arrows in (A), (E), and (I) indicate similar
location near the periphery of cisternae where a persistent puncta
occurs. Bar = 10 mm.
Cytoskeletal Control of Plant Endoplasmic Reticulum
3941
and closure are two of the more transient events in the network.
Until now, most analysis of the tubular network has focused on
these transient events that generate and remove the tubule
polygons that make up the network (Lee and Chen, 1988; Knebel
et al., 1990; Lichtscheidl and Url, 1990), but some have noted
that punctate structural features persist (Knebel et al., 1990;
Lichtscheidl and Url, 1990). Observations using persistency
maps confirm that there is a population of distributed, persistent
punctae (cisternae between 0.1 and 0.3 mm2; Figures 1G to 1I
and 1G’ to 1I’) that could function as the nodal framework upon
which network movement takes place. Although these punctae
may serve to anchor persistent features in the otherwise dynamic
network, they are less persistent and may move (gray punctae
near arrowheads, Figure 1H’) in regions of rapid streaming.
Therefore, instead of referring to them as anchor sites, here we
prefer to describe them as anchor/growth sites, and we predict
that they are the same anchor sites around which the ER tubules
can be remodeled manually using laser-trapped Golgi bodies
(Sparkes et al., 2009b).
Figure 4. Quantification of Persistent ER Tubules and Cisternae under
Various Treatment Conditions.
(A) Persistent tubules in control GFP-HDEL–expressing cells (black,
Control), GFP-HDEL, and eYFP-XIK tail coexpressing cells (white, XIK),
latrunculin b–treated GFP-HDEL–expressing cells (light gray, LATB), and
oryzalin-treated GFP-HDEL–expressing cells (gray). Columns show average value, and error bars show SD in total number of movies evaluated,
n (Control, n = 30; XIK, n = 27; LATB, n = 14; oryzalin, n = 16). Percentage
of tubule area is the percentage of the total imaged membrane (from
summed images, refer to Figures 1 to 3C, black and gray regions) that is
persistent tubules (high persistency tubules, refer to Figures 1 to 3F).
Number/100 mm2 is the number of persistent tubules per 100 mm2 of total
imaged membrane. Length average is the average length in micrometers
of the persistent tubules. The differences between all except the controloryzalin pair (all categories) and the XIK-LATB pair (tubule length cate-
gory) are significant (P < 0.05, Tukey’s HSD multiple comparisons of
means, 95% family-wise confidence level).
(B) Persistent punctae in control GFP-HDEL–expressing cells (black,
Control), GFP-HDEL, and eYFP-XIK tail coexpressing cells (white),
latrunculin b (light gray), and oryzalin (medium gray) treated GFPHDEL–expressing cells, and GFP-HDEL and eYFP-XIJ tail coexpressing
cells (dark gray). Columns show average value, and error bars show SD in
total number of movies evaluated, n (Control, n = 30; XIK, n = 27; LATB,
n = 14; oryzalin, n = 16; XIJ, n = 4). Percentage of cisternal area is the
percentage of the total imaged membrane (from summed images, refer
to Figures 1 to 3C, black and gray regions) that is persistent punctae
(high persistency cisternae with areas between 0.1 and 0.3 mm2 and
circularity > 0.5; refer to Figures 1 to 3I; see Supplemental Figures 2 and
3F online). Number/100 mm2 is the number of persistent punctae per 100
mm2 of total imaged membrane. The LATB treatment is significantly
different (P < 0.05) from all of the other treatments, if the XIJ data are not
included in the set because of low sample size. The other treatments do
not significantly differ from each other (Tukey’s HSD multiple comparison
of means, 95% family-wise confidence level).
(C) Persistent cisternae in control GFP-HDEL–expressing cells (black),
GFP-HDEL and eYFP-XIK tail coexpressing cells (white), latrunculin
b–treated (light gray) and oryzalin-treated (medium gray) GFP-HDEL–
expressing cells, and GFP-HDEL and eYFP-XIJ tail coexpressing cells
(dark gray). Columns show average value, and error bars show SD in total
number of movies evaluated, n (same as [B]). Percentage of cisternal
area is the percentage of the total imaged membrane (from summed
images, refer to Figures 1 to 3C, black and gray regions) that is persistent
cisternae (high persistency cisternae >0.3 mm2, refer to Figures 1 to 3I;
see Supplemental Figures 2 and 3F online). The Control-LATB and -XIK
pairs are significantly different, as are the Oryzalin-LATB and -XIK pairs
and the XIJ-LATB pair (P < 0.05). Number/100 mm2 is the number of
persistent cisternae per 100 mm2 of total imaged membrane. All pair
comparisons except the Control-XIJ, Oryzalin-XIJ, XIK-XIJ, and OryzalinXIK pairs are significant (P < 0.05). Area average is the average area in
mm2 of the persistent cisternae. The only pair comparisons that are
significant (P < 0.05) are Control-LATB, Control-XIK, and Oryzalin-XIK.
Tukey’s HSD multiple comparisons of means were done at 95% familywise confidence level.
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associated motors might mediate the actomyosin dependence
of ER movement (Brandizzi et al., 2002; Hawes et al., 2008).
However, the results presented here indicate that Golgi bodies
do not affect ER remodeling (Figure 7) because when the Golgi
have been deconstructed with BFA, new ER tubules still actively
form and retract (see Supplemental Movie 6 online). The rate of
movement of ER membrane protein (t1/2 values) after BFA
treatment, as estimated by photoactivation (Figures 6I to 6L),
did not differ from other treatments, but the mobile fraction was
lower, matching myosin XIK tail coexpression values most
closely (Figure 6L). Although Golgi bodies do not appear to drive
ER, an as yet unanswered question is, does ER drive Golgi
bodies? However, the ER does appear to exert an effect on Golgi
stack location in terms of constraining them to the tubular ER or
to the rims of cisternal regions (Figure 7) (Boevink et al., 1998;
Runions et al., 2006; Hawes et al., 2008). While a comprehensive
study of all the Arabidopsis myosins, through transient expression studies in tobacco epidermal cells, has been implemented,
it has been shown that the XIK tail domain, but not the XIJ tail,
significantly affects movement of Golgi and mitochondria (Avisar
et al., 2009). Similarly, we show a comparable differential effect
on ER movement with the same myosin tail domains, perhaps
indicating an interdependent motion of the two organelles driven
by the same myosin.
The Effect of the Actomyosin System and Microtubules on
ER Remodeling and Movement
Figure 5. The Effect of the eYFP-XIK Tail Domain and Latrunculin b on
FRAP of Tobacco Leaf Epidermal ER Marker GFP-CXN.
FRAP of a bleached spot area in control cells ([A] to [C]), cells treated
with latrunculin b ([D] to [F]), and those coexpressing eYFP-XIK tail
domain ([G] to [I]) were quantified where prebleach ([A], [D], and [G]),
bleach ([B], [E], and [H]) and 1.21 s postbleach ([C], [F], and [I]) are
shown. Coexpression of GFP-CXN and the eYFP-XIK tail domain ([J];
merged GFP-CXN image in [K]) prior to the bleach experiment shown in
(G) to (I) is indicated. The bleached area of GFP-CXN (circle) was
monitored over time, and the fluorescence recovery was plotted ([L];
where the green line is control, black line is latrunculin b treatment, and
red line is coexpression of GFP-CXN and the eYFP-XIK tail domain).
Bars = 2 mm.
The Role of the ER-Golgi Interface in ER Remodeling
There is an intimate association of Golgi bodies with moving ER
exit sites (daSilva et al., 2004), ER tubule growth appears to
follow Golgi body movement (Brandizzi et al., 2002), and the flow
of proteins within the ER membrane corresponds in direction and
magnitude to that of streaming nearby Golgi (Runions et al.,
2006). These observations prompted the notion that Golgi-
Both eYFP-XIK myosin tail coexpression and latrunculin b treatment affect the appearance and dynamics of the ER (Figures 2
and 3; see Supplemental Movies 2 and 3 online). The eYFP-XIK
myosin tail domain perturbs streaming of Golgi, mitochondria,
and peroxisomes (Peremyslov et al., 2008; Sparkes et al., 2008),
presumably by interfering with the activity of endogenous myosin
XIK through titration of accessory factors required for binding
cargo or through titration of active myosin dimers required to
bind cargo. Latrunculin b treatment reduces the pool of polymerized F-actin by complexing G-actin monomers so that actin
filaments disappear (Coue et al., 1987; see Supplemental Figures
1C and 1D online). Not only do both these treatments result in an
increase in the presence of persistent ER tubules (Figure 4A;
number/100 mm2 and length average), but they also produce a
pattern of more persistent polygonal ring structures (cf. Figures
2, 3D, and 3E with Figures 1D, 1D’, 1E, and 1E’), resulting in a
persistency map of polygonal ring structures connected by
persistent tubules. Furthermore, although both treatments give
rise to increased persistent cisternal area (Figure 4C, area
average), latrunculin b treatment appears to generate more of
Table 1. ER Membrane Dynamics Determined with FRAP of GFP-CXN
under Various Treatment Conditions
Treatment
t1/2 (s)
Mobile Fraction
(Plateau) (%)
Control (n = 57)
Latrunculin B (n = 16)
eYFP-XIK tail (n = 48)
2.37 6 0.49
2.45 6 0.44
2.33 6 0.58
87.72 6 10.18
89.14 6 7.39
79.09 6 13.20
Cytoskeletal Control of Plant Endoplasmic Reticulum
3943
an increase in the number of persistent punctae (Figure 4B,
number/100 mm2) and persistent cisternae (Figure 4C; number/
100 mm2) than truncated myosin XIK coexpression.
The increase in the size of persistent ER cisternae (Figure 4C,
area average) may arise through a common mechanism, (i.e.,
inhibition of dynamic or exploratory tubular outgrowth of membranes from anchor sites on cisternae that would otherwise
diminish cisternal area). The observation that persistent punctae
are redistributed to cisternae and their periphery after XIK tail
coexpression (arrows, Figure 2I) and latrunculin b treatment
(arrowhead, Figure 3A, 3E, and 3I) indicates that the nodal
punctae may not only be anchor sites but may also act as growth
sites, stopped at some point in the tubule growth cycle.
We observed no significant effect of oryzalin on ER dynamics
other than an increase in cisternae. Oryzalin at the concentration
and time used is effective at depolymerizing microtubules (see
Supplemental Figures 1A and 1B online). Recent studies have
highlighted that oryzalin can have a secondary effect leading to
ER nodulation (Langhans et al., 2009), which could explain the
apparent increase in small persistent cisternae observed here.
Unlike Knebel et al. (1990), who also noted some ER nodulation,
Langhans et al. (2009) also reported an effect on ER dynamics
but attributed this to the drug and not to the absence of
microtubules. Our evidence supports that of Knebel et al.
(1990); microtubule depolymerization has no effect on ER dynamics. In addition, the small persistent cisternae may actually
be swollen anchor/growth sites, as suggested by Knebel et al.
(1990). We should caution, however, that this by no means
excludes a role for microtubules exerting a regulating influence
on the dynamics of the ER at other stages of the cell cycle.
Indeed, it is quite likely that during the transition to metaphase,
while the nuclear envelope resorbs into the ER, which then forms
aggregates at the spindle poles, there is a switch over from an
actin-based movement to microtubule-based movement on
spindle microtubules (Gupton et al., 2006). The association of
ER membrane with spindle microtubules in plants is well established (Hepler and Wolniak, 1984).
Figure 6. The Effect of mRFP-XIK Tail Domain, Latrunculin b, and BFA
on the Mobility of CXN-Photoactivated GFP in Tobacco Leaf Epidermal ER.
A spot activated area (circle) of CXN-photoactivatable GFP was generated, and its mobility out of the activation spot was monitored over time
([A] to [K]) and quantified ([L]; where the green line is control, black line is
latrunculin b treatment, red line is coexpression of myosin XIK tail
domain, and the blue line is BFA treatment). Mobility of CXN-paGFP in
control cells ([A] to [C]), cells treated with latrunculin b ([D] and [E]),
those coexpressing CXN-paGFP and the mRFP-XIK tail domain ([F] to
[H]) or treated with BFA ([I] to [K]) were quantified where activation ([B],
[D], [G], and [J]) and 1.93 s postactivation ([C], [E], [H], and [K]) are
shown. An example of preactivation (A) highlights the lack of fluorescence prior to photoactivation in control cells. Expression of the mRFPXIK tail domain (F) and the effect of relocation of ST-mRFP back to
the ER (I) in the activation experiments shown ([F] to [K]) are indicated.
Bars = 2 mm.
A Model for the Involvement of Actin and Myosin in ER Form
and Remodeling
Observations of ER form reveal a dynamic structure that undergoes rapid remodeling in an actin-dependent manner. Here,
we present the development of analytical software in order to
quantify these observations, with some surprising revelations
relating to persistent growth/anchor sites and the movement of
the ER surface in relation to the active remodeling structure.
Table 2. ER Membrane Dynamics Determined with Photoactivation of
CXN-paGFP under Various Treatment Conditions
Treatment
t1/2 (s)
Control (n = 45)
Latrunculin B (n = 20)
mRFP-XIK tail (n = 31)
BFA (n = 28)
2.01
2.31
1.85
1.79
6
6
6
6
1.15
1.22
1.11
0.98
Plateau (%)
Mobile
Fraction (%)
2.37 6 4.26
10.58 6 7.04
8.73 6 6.12
8.12 6 8.08
97.63
89.42
91.27
91.88
6
6
6
6
4.26
7.04
6.12
8.08
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The Plant Cell
Figure 7. Golgi Bodies Do Not Collocate with Persistent ER Puncta in
Tobacco Epidermal Cells and Do Not Affect ER Membrane Remodeling.
Cells coexpressing GFP-HDEL ([A]; green) and ST-mRFP ([B]; magenta)
treated with latrunculin b highlight the persistent puncta on the cisternal
ER and show that the Golgi bodies do not collocate to these regions (C).
Cells coexpressing GFP-HDEL and ST-mRFP were treated with BFA (3 h,
100 mg/mL) ([D] to [I]), and remodeling of the ER network was monitored
over time. The Golgi marker ST-mRFP has relocated to the ER after BFA
treatment, indicating Golgi body absence ([E] and merged image in [F]).
Consecutive images ([G] to [I]) show that the ER network is able to
remodel and tubule outgrowth still occurs (arrowhead) after BFA treatment. Bars = 2 mm.
Based on these results, we propose a model for how the
actomyosin system regulates ER remodeling by interacting
with key structural elements, including growing, shrinking, and
stabilized tubules, three-way junctions, anchor/growth sites, and
tubule/cisternal transitions (Figure 8). In plants, extremely little is
understood regarding any of these key elements, and only
recently has it emerged that reticulons are involved in constricting the ER, and we have proposed a role for them in modulating
cisternalization versus tubulation (reviewed in Tolley et al., 2008;
Sparkes et al., 2009a).
ER Tubule Growth/Shrinkage
Since it is not clear how ER tubules and microfilaments mutually
orient themselves during growing/shrinking cycles, the relative
roles of myosin and actin polymerization in ER tubule remodeling
remain speculative. However, generating a model that helps
explain the results in the context of what is currently known about
cytoskeletal dynamics is desirable. The central features of the
model (Figure 8) are that the ER is centered on anchor/growth
sites and moves along microfilaments, mainly through its attachment to a myosin such as XIK, but also via actin polymer-
ization. This generates tubule nonpersistency through cycles of
tubule elongation and shrinkage (Figure 8, processes 1 to 3) as
well as mutual sliding of tubules generating polygonal ring
closure (Figure 8, process 5). Both myosin XIK tail coexpression
with GFP-HDEL and latrunculin b treatments diminish dynamic,
exploratory tubules by blocking either tubule shrinkage or growth
(Figure 8, processes 1 to 3). This is supported by Figure 4A,
where both latrunculin b and myosin XI-K expression increase
the length and number of persistent tubules. Inhibition of growing/shrinking cycles of tubules after F-actin has depolymerized is
supported by the large number of blind ended tubules in Figures
2 and 3, which may have started a cycle but not completed it. We
further postulate that there is a polarity for ER tubule remodeling
and the actomyosin system (Figure 8, processes 1 to 3 and 5);
growth is mainly driven by actin polymerization, as supported by
the observation that the rate of in vivo actin polymerization (1.7
mm/s; Staiger et al., 2009) is very close to the rate of tubule
growth previously reported (1.34 6 0.54 mm/s; Sparkes et al.,
2009b), while shrinkage is mediated by (+) end-directed myosin
processivity. Myosins are classically defined as (+) end-directed
motors. Although this is true for a tobacco myosin (Tominaga
et al., 2003), the processivity and directionality of Arabidopsis
myosins is not known at this time.
Based on the BFA studies presented here, ER tubule remodeling still occurs in the absence of Golgi bodies. Therefore, the
potential role of ER tubule growth in Golgi body movement is also
depicted whereby a physical link, be it direct membrane continuity or interacting protein tethers, between the two organelles
results in interdependent movement (Figure 8, process 4).
Three-Way Junction and Anchor Sites Are Stabilization
Factors to Maintain Tubules
Three-way junctions and anchor/growth sites are evident under
all treatment conditions. Through micromanipulating Golgi bodies, we showed that the underlying ER did indeed remodel and
resulted in the identification of ER anchor sites (Sparkes et al.,
2009b). However, it is unclear at this time whether all anchor sites
are indeed growth sites and vice versa. The data presented here
indicate two populations of anchor sites/growth sites; a stable
persistent group and a transitory nonpersistent group. Upon
latrunculin b treatment, the latter become persistent, resulting in
a significant increase in number over the other treatment conditions (Figure 4B). Therefore, perhaps an actin-dependent process is required to determine the extent of movement of anchor
sites (Figure 8, process 3). Myosins may also be involved in the
movement of tubules toward moving anchor points but do not
affect the movement of anchor sites as much as latrunculin b
because the number of persistent punctae do not increase after
truncated myosin XIK expression (Figure 4B).
Proteinaceous factors regulating tubule stability can also be
tested using this system. For example, in mammals, CLIMP63
(cytoskeletal linking membrane protein p63) is required to interact with microtubules to stabilize the cortical ER network (see
review in Vedrenne and Hauri [2006] and references therein;
Klopfenstein et al., 1998). Growth of tubules in animal systems is
also dependent on integral ER membrane proteins (e.g., stromal
interaction molecule 1) that bind the plus end of growing
Cytoskeletal Control of Plant Endoplasmic Reticulum
3945
Figure 8. Model of the Different Activities of F-Actin and Myosin at Anchor/Growth Sites in Tubule Dynamics at Nonanchored Sites and in Determining
the Mobile Fraction of ER Membrane Proteins.
Process 1: Tubules grow outwards, driven by actin polymerization at an anchor/growth site. The anchor site is proposed to be a region where the
plasma membrane and ER are closely associated. These sites may contain actin nucleation and growth proteins. The mechanism of actin attachment to
the growing tubule is represented diagrammatically as a hypothetical protein associated with the growing tubule tip at the pointed () end of the actin
filament. Process 2: Tubules shrink or retract in the (+) direction in a myosin-dependent fashion. This is supported by Figure 4A, where both latrunculin b
and myosin XI-K expression increase the length and number of persistent tubules. We interpret this as arising from an absence of shrinkage and
stabilization of untethered tubules. Process 3: Less persistent anchor/growth sites can move, perhaps recruited at branch points in the actin network,
indicated here by the presence of an ARP2/3 complex. There are fewer driven junctions after latrunculin b and myosin XI-K expression (Figures 2 and 3),
as indicated by persistent kinks. Blind-end tubules in Figures 2 and 3 may also be anchor sites that cannot move far from the point of origin but have
tubules attached. Note that three-way junctions do not require tethering to an anchor site. Process 4: Golgi do not regulate all of the growth or shrinkage
of ER tubules. After BFA treatment, which deconstructs the Golgi, new ER tubules still grow and shrink (Figure 7). However, since the moving ER and
moving Golgi remain closely associated, an interdependence of ER membrane movement and Golgi movement seems likely. Process 5: Myosin
mediates ring closure of polygons. The association of myosin with an unanchored three-way junction is postulated, and the special nature of the
junction is inferred. The observation that intact rings of the polygonal network are persistent after myosin XIK tail expression (Figures 2D to 2F) and
latrunculin b treatment (Figures 3D to 3F) is evidence to support this part of the model. Process 6: Actomyosin dependence of polygon filling. Removal of
myosin may result in the collapse of the corner with high angle of curvature and polygon filling, leading to cisternae and giving rise to the larger cisternal
area and larger number of cisternae after myosin XIK tail expression (Figure 4C).
microtubules (Grigoriev et al., 2008). By isolating analogous
actin-dependent factors in plants, such as the hypothetical
minus-end tip associated complex in Figure 8, persistency
maps can determine which proteins are required for tubule,
anchor point, and three-way junction stability.
The model presented for cortical ER dynamics in higher plants
relies heavily on myosin and actin dynamics. Therefore, one
assumption would be that upon actin depolymerization, the ER
network would disintegrate; however, it remained in place over
the time frame of the experiment albeit with altered tubule length/
dynamics/cisternalization (see Boevink et al., 1998). We hypothesize that the anchor points, which might be connected to the
plasma membrane, are required to hold the ER network in place
and that the tubules are stabilized by accessory scaffolding or
coat proteins in the absence of actin. This is shown in the model
in Figure 8, where the tubules forming the central polygon are
tethered between anchor/growth sites in the absence of directly
underlying microfilaments.
Tubule/Cisternal Transitions
The actomyosin system also appears to be involved to some
extent in tubulation versus cisternalization as latunculin b treatment and overexpression of the myosin XIK tail domain resulted
in increased persistent cisternae (Figure 4C) compared with
control and other treatment conditions. Based on the large
number of observed persistent polygonal structures (Figures 2
and 3) compared with the control (Figure 1), we propose that the
actomyosin system is also required for polygon remodeling. This
is achieved by the relative lateral sliding of newly formed
interconnected tubules (Figure 8, process 5). Actomyosin involvement in polygon ring closure (Figure 8, process 5) may have
a flip side. In the absence of functional actomyosin, branch sites
making up the polygon ring may become unstable and the inner
side of the tubule corner, where there are high degrees of
curvature, would fill with membrane, causing more cisternalization (Figure 4C) through polygon filling (Figure 8, process 6).
3946
The Plant Cell
The Effect of the Actomyosin System on ER Membrane
Protein Mobility
The data presented here, based on the mobility of the transmembrane domain of calnexin (CXN) fused to GFP, indicate that
ER remodeling and the direction of movement of the surface
membrane could be linked but are not linked to the magnitude of
the diffusive movement. Initial membrane diffusion rates (t1/2) are
similar in control to a latrunculin b–treated immotile ER network
system. Latrunculin treatment and overexpression of myosin XIK
tail domain remove the directional restriction on diffusion, while
restricting the available amount of molecules available for diffusion (mobile fraction, Tables 1 and 2).
We hypothesize that native myosin XIK might naturally maintain a high mobile fraction of the CXN-GFP construct in the ER
membrane by destabilizing actin-membrane linkages during
shrinkage (Figure 8, process 2) or outgrowth of the membrane
(Figure 8, process 3). The eYFP-XIK myosin tail domain is present
throughout the cytosol and could therefore directly interact with
the cortical ER, producing a lower mobile fraction of ER membrane proteins by interfering with the activity of native myosin
XIK. Alternatively, instead of interfering directly with native myosin, the tail domain could remove an active component required
for membrane flow (through titration of accessory factors) or
introduce a rigid locked cytoskeletal scaffold on the ER surface
causing a barrier to free diffusive movement of proteins.
Latrunculin b treatment decreased the mobile fraction of ER
membrane CXN fused to paGFP/GFP in the photoactivation
experiments but not in the FRAP experiments. Photoactivation
may target either tubules or cisternae (both are invisible prior to
photoactivation), while FRAP targets preselected cisternae.
Latrunculin b treatment produces more persistent tubules (Figures 2 to 4), stabilized perhaps in a way that would diminish the
mobile fraction of protein in the membrane. Latrunculin b also
produces cisternae with many peripheral punctae (Figures 3H
and 3I, arrows) that may be stabilized by myosins, while the
center of the cisternae remains relatively unaffected, with no
apparent decrease in mobile fraction, as detected by FRAP.
Comparison of the results of this study with an earlier one using
a similar technique to evaluate ER membrane dynamics (Runions
et al., 2006) shows an apparent disparity in the diffusion rate of
CXN-paGFP. Half-times of recovery in the previous study were
reported to be 5 to 10 times longer than in this study, while mobile
fraction percentages are comparable. Reexamination of the
older data shows that in the previous study, entire small cisternal
regions of ER were photoactivated so that diffusion of activated
paGFP was only possible through connecting tubules of small
radius. This resulted in a bottleneck effect and slower movement
of GFP away from the activation region than measured in this
study, where FRAP and photoactivation experiments were
performed on the tubular network of ER or larger cisternal sheets
that allowed a steeper diffusion gradient of GFP.
This study builds on the previous work by establishing the
diffusive, nonmotorized nature of movement of ER membrane
protein even during rapid ER remodeling. Previous work
(Runions et al., 2006) demonstrated a directionality to this
movement that can be seen in Figures 6B and 6C (control) and
Figures 6J and 6K (BFA treatment). The directionality of this
diffusion is changed when the actomyosin system is perturbed
(Runions et al., 2006; Figures 6D, 6E, 6G, and 6H). Here, we show
that the average rate of movement itself is not affected by the
actomyosin network, indicating nonmotorized and diffusive
movement. However, occasional fast, apparently nondiffusive
movements were monitored but were infrequent in comparison
to diffusional movements. A better quantification is required
to determine vectorial, directional diffusive versus apparent
active movements. The fact that directional diffusion occurs
after BFA treatment helps establish that directional diffusion is
not a property of tubules alone. This process of directional
diffusion within tubules (near one-dimensional) and cisternae
(near two-dimensional) may provide a mechanism whereby
proteins and metabolites can be more efficiently transferred
through the cell with minimal expenditure of energy at rates that
far exceed those of nondirectional diffusion in three dimensions
(Adam and Delbruck, 1968; Loverdo et al., 2008; Mirny, 2008;
Pickard, 2003).
In conclusion, we have found that the cortical ER network in
tobacco epidermal cells undergoes rapid remodeling that is
coupled to the directionality but not the magnitude of ER surface
flow. ER remodeling is an actomyosin-dependent process and
is independent of microtubules. Quantification of ER tubule
growth, cisternalization, and recognition of anchor/growth sites
has led to a model defining the basic principles behind remodeling of the plant cortical ER in leaf epidermal cells. Additionally,
ER network remodeling occurs in the absence of Golgi bodies,
thereby addressing the long-held postulate that Golgi body
movement might drive ER tubule growth. Future work using
this model and persistency mapping will help identify factors
required for macro- and microscopic changes in ER network
remodeling.
METHODS
Plant Material and Constructs
Nicotiana tabacum plants were grown according to Sparkes et al. (2005).
Fluorescent protein fusion constructs including the ER marker GFP-HDEL
(Batoko et al., 2000), GFP-FABD2 (Sheahan, et al., 2004), GFP-TUA6
(Ueda et al., 1999), eYFP- and mRFP-myosin XIK tail domain (eYFP-XIK
and mRFP-XIK; Sparkes et al., 2008), eYFP-XIJ tail domain (eYFP-XIJ;
Avisar et al., 2009), transmembrane domain of calnexin (CXN-paGFP)
(Runions et al., 2006), GFP transmembrane domain of CXN (GFP-CXN;
Irons et al., 2003), and sialyl transferase signal anchor sequence-mRFP
(ST-mRFP; Saint-Jore et al., 2002) were all infiltrated according to
Sparkes et al. (2006) with an optical density of 0.1 except for GFPFABD2 at 0.02 and GFP-HDEL, CXN fusions, and ST-mRFP, which
required 0.04 optical density. The mRFP-XIK tail domain-pB7WGR2
construct was generated by Gateway cloning using the entry vector XIK
tail-pDonor 207 (Sparkes et al., 2008).
Sample Preparation and Image Acquisition
The ER in the outermost cortical region of adaxial leaf epidermal pavement cells was imaged. Where stated, leaf segments of ;25 mm2 were
incubated either in BFA (100 mg/mL) for 3 h or in latrunculin b (25 mm) or
oryzalin (14 mm) for 30 min prior to imaging.
Dual imaging of YFP and GFP was done using multitracking in line
switching mode on a Zeiss LSM510 Meta confocal microscope. GFP was
Cytoskeletal Control of Plant Endoplasmic Reticulum
excited with the 458-nm line, and eYFP with the 514-nm line of an argon
ion laser using a 458/514 dichroic mirror, and the subsequent emission
was detected using 470- to 500-nm and 560- to 615-nm band-pass
filters. Similarly, GFP and mRFP emission was captured using multitracking in line switching mode. GFP was excited with a 488-nm argon
laser and mRFP with a 543-nm laser and their emissions detected using a
488/543 dichroic mirror and 505- to 530-nm and 560- to 615-nm bandpass filters. All imaging was performed using a 363 1.4–numerical
aperture oil immersion objective.
For persistency mapping, time-lapse images of GFP-HDEL were
captured using a 2- to 3-mm pinhole, 512 3 512 pixel resolution, and
32.3 digital zoom. To reduce noise, 43 line averaging was used. The
scan rate was increased by imaging a 955- to 960-mm2 region of interest
so that 50 frames per 80 s were captured (0.63 frames/s). For samples
where cells were coexpressing a fluorescent myosin tail domain and
GFP-HDEL, coexpression was verified before time-lapse imaging of the
GFP-HDEL alone was performed.
FRAP and Photoactivation
ð1Þ
The exponentially decreasing curve used to fit photoactivation data
took the form:
IðtÞ ¼ Aeð 2 BtÞ þ ðC=ðt þ 1ÞÞ;
(plateau) and the half-time of fluorescence recovery (t1/2). The plateau
value was the highest-intensity value postbleach for FRAP and the lowest
intensity value for photoactivation in each data set. To determine the
plateau, the calculated parameters A, B, and C, which gave the best fit in
Equations 1 and 2 were used to fit a theoretical curve to the duration of
each data set. Half time of fluorescence recovery/decay is the time at
which half of the fluorescence has recovered/decayed after bleaching/
photoactivation. Half-fluorescence intensity (I1/2) was half the difference
between minimum and plateau fluorescence values. Half-time of fluorescence recovery was interpolated at I1/2 using Prism 5 software
(GraphPad Software).
Multivariate analysis of variance (MANOVA) was used to compare the
families of curves derived in each of the experiments based on their
plateau and t1/2 values using SPSS software. Wilks’ lambda was used as a
test statistic for differences between experiments, and post-hoc tests to
compare differences between individual pairs of experiments were done
by computing Tukey’s HSD statistic.
Modeling the Cortical ER in Tobacco Epidermal Cells
For FRAP experiments, five prebleach scans were captured using settings as for GFP-HDEL (above) with the 488-nm laser transmission set to
1% transmission prior to bleaching of a 10- to 11-mm2 spot using 50
iterations of a 405-nm diode laser set to 100% transmission. Fast intensity
recovery required a high scan rate (4.57 frames/s); therefore, time-lapse
imaging was performed using 256 3 256 pixel resolution and 2.83 digital
zoom.
For photoactivation of CXN-paGFP, 30 prephotoactivation scans were
captured using settings as for GFP-HDEL (above) with the 488-nm laser
transmission set to 1% transmission prior to activation of the 8- to 10-mm2
spot using five iterations of the 405-nm diode laser set at 100% transmission. Inherent high mobility of the ER required a high scan rate.
Postactivation time-lapse imaging was therefore performed using 128 3
128 resolution (6.8 frames/s) and 2.83 digital zoom with the 488-nm laser
set at 1% transmission.
Mean fluorescence intensity was calculated (Zeiss LSM software)
within the bleach (FRAP) or activation (photoactivation) region during a
30- to 40-s postbleach or postactivation period. For FRAP, recovery of
fluorescence occurred on a scale normalized from 0% for the first
postbleach scan to 100% for the prebleach mean fluorescence level
within the region of interest. For photoactivation, normalized intensity
occurred on a scale between 0%, which represented mean preactivation
intensity, and 100%, the immediate postactivation intensity (Graumann
et al., 2007).
Normalized data from the time of bleaching/photoactivation were
plotted against time using Prism 5 software (GraphPad Software). Nonlinear regression was used to fit curves.
The exponentially increasing curve used to fit the FRAP data took the
form:
IðtÞ ¼ AeðB=ðtþ1ÞÞ
3947
Persistency maps were generated in Image J (version 1.37v; Wayne
Rasband, National Institutes of Health, Bethesda, MD). Beginning at
frame number 5 of 50, every fifth frame was compared with frames 8 s
(five frames) earlier with a Boolean “AND” operation and by subtraction.
The subtraction result was subsequently subtracted from the AND result
(with minor improvement), resulting in a set of 45 frames with objects that
persisted, unmoving for at least 8 s. Binary conversion of objects in the 13
to 255 (the brightest pixels) gray level range was followed either by a
closing operation (default in ImageJ, one iteration, one pixel count) that
connected tubules that may have become discontinuous during the
second subtraction operation or by an opening operation that removed
tubules between cisternae. The closed binary images were skeletonized
and summed, producing the tubule persistency map. The opened binary
images were summed, producing the cisternal persistency map. To count
and measure persistent tubules and cisternae, eight-bit persistency
maps were segmented between 0 and 180 gray levels of 256 (the darkest
tubules and cisternae). For tubule maps, the number and length (half the
perimeter) of resultant persistent tubules were measured. For cisternae
maps, the number and area of the resultant cisternae were measured.
To generate percentage area values and number per 100 mm2, the total
imaged membrane for each movie was determined by summing the
frames and segmenting between 15 and 255 gray levels (gray and black
areas in Figures 1 to 3C). The brightest regions, between 50 and 255 gray
levels (black areas in Figures 1 to 3C) look somewhat like persistent
cisternae and punctae (Figures 1 to 3H) but do not show good spatial
correlation. These regions may either be persistent or have high local
fluorophore concentration.
Quantitation of the most persistent tubules, punctae, and cisternae was
done, normalizing each movie to the total membrane area imaged (panel
C in Figures 1 to 3) in the percentage area and number/100 mm2 values.
Statistical comparison of the data was done using R version 2.7.1 (http://
www.r-project.org/) using analysis of variance and Tukey’s HSD two-way
comparison at 95% confidence interval.
ð2Þ
Supplemental Data
where A, B, and C are parameters of the curve and e is the base of the
natural logarithms.
Individual data sets within each experiment were fit independently to
assess variation within groups, and data points from all sets within a
group were fitted by sharing curve parameters in order to derive a master
fit for graphical presentation. Comparison across experimental groups
was done by deriving the mobile fraction of bleached/activated molecules
The following materials are available in the online version of this article.
Supplemental Figure 1. Oryzalin and Latrunculin b Depolymerize
Microtubules and Actin, Respectively.
Supplemental Figure 2. Persistency of Tubules, Punctae, and Cisternae of Cortical ER in a Tobacco Epidermal Cell Expressing GFPHDEL Treated with Oryzalin.
3948
The Plant Cell
Supplemental Figure 3. Persistency of Tubules, Punctae, and Cisternae of Cortical ER in a Tobacco Epidermal Cell Coexpressing GFPHDEL and eYFP-XIJ Myosin Tail Construct.
Supplemental Movie 1. Cortical ER Network Remodeling in Two
Control Tobacco Epidermal Cells.
Supplemental Movie 2. Cortical ER Network Remodeling in Tobacco
Epidermal Cells Coexpressing GFP-HDEL and the eYFP-XIK Tail
Domain.
Supplemental Movie 3. Cortical ER Network Remodeling in Tobacco
Epidermal Cells Treated with Latrunculin b.
Supplemental Movie 4. Cortical ER Network Remodeling in Tobacco
Epidermal Cells Expressing GFP-HDEL Treated with Oryzalin.
Supplemental Movie 5. Cortical ER Network Remodeling in Tobacco
Epidermal Cells Coexpressing GFP-HDEL and the eYFP-XIJ Tail
Domain.
Supplemental Movie 6. Cortical ER Network Remodeling in Tobacco
Epidermal Cells Treated with BFA.
ACKNOWLEDGMENTS
We thank Janet Evins for technical support during the work, D.W.
McCurdy for the GFP-FABD2 constructs, and T. Hashimoto for GFPTUA6 constructs. I.S. was supported by a Leverhulme Trust grant
(F/00382/G) to C.H. J.R. was funded by a Biotechnology and Biological
Science Research Council grant (BB/F014074/1), and L.G. was funded
by the International Research Travel/Award Grant and Faculty Development Leave programs at Texas A&M University.
Received October 16, 2009; revised November 19, 2009; accepted
December 7, 2009; published December 29, 2009.
REFERENCES
Adam, G., and Delbruck, M. (1968). Reduction of dimensionality in
biological diffusion processes. In Structural Chemistry in Molecular
Biology, A. Rich and N. Davidson, eds (San Francisco, CA: Freeman),
pp. 198–215.
Avisar, D., Abu-Abied, M., Belausov, E., Sadot, E., Hawes, C., and
Sparkes, I.A. (2009). A comparative study of the involvement of 17
Arabidopsis myosin family members on the motility of Golgi and other
organelles. Plant Physiol. 150: 700–709.
Avisar, D., Prokhnevsky, A.I., Makarova, K.S., Koonin, E.V., and
Dolja, V.V. (2008). Myosin XI-K is required for rapid trafficking of Golgi
stacks, peroxisomes and mitochondria in leaf cells of Nicotiana
benthamiana. Plant Physiol. 146: 1098–1108.
Batoko, H., Zheng, H.Q., Hawes, C., and Moore, I. (2000). A Rab1
GTPase is required for transport between the endoplasmic reticulum
and Golgi apparatus and for normal Golgi movement in plants. Plant
Cell 12: 2201–2218.
Boevink, P., Martin, B., Oparka, K., Santa Cruz, S., and Hawes, C.
(1999). Transport of virally expressed green fluorescent protein
through the secretory pathway in tobacco leaves is inhibited by
cold shock and brefeldin A. Planta 208: 392–400.
Boevink, P., Oparka, K., Santa Cruz, S., Martin, B., Betteridge, A.,
and Hawes, C. (1998). Stacks on tracks: The plant Golgi apparatus
traffics on an actin/ER network. Plant J. 15: 441–447.
Brandizzi, F., Snapp, E.L., Roberts, A.G., Lippincott-Schwartz, J.,
and Hawes, C. (2002). Membrane protein transport between the
endoplasmic reticulum and the Golgi in tobacco leaves is energy
dependent but cytoskeleton independent: Evidence from selective
photobleaching. Plant Cell 14: 1293–1309.
Brown, J.R., Stafford, P., and Langford, G.M. (2004). Short-range
axonal/dendritic transport by myosin-V: A model for vesicle delivery to
the synapse. J. Neurobiol. 58: 175–188.
Coue, M., Brenner, S.L., Spector, I., and Korn, E.D. (1987). Inhibition
of actin polymerization by latrunculin A. FEBS Lett. 213: 316–318.
daSilva, L.L.P., Snapp, E.L., Denecke, J., Lippincott-Schwartz, J.,
Hawes, C., and Brandizzi, F. (2004). Endoplasmic reticulum export
sites and Golgi bodies behave as single mobile secretory units in plant
cells. Plant Cell 16: 1753–1771.
De Craene, J.O., Coleman, J., Estrada de Martin, P., Pypaert, M.,
Anderson, S., Yates III, J.R., Ferro-Novick, S., and Novick, P.
(2006). Rtn1p is involved in structuring the cortical endoplasmic
reticulum. Mol. Biol. Cell 17: 3009–3020.
Estrada, P., Kim, J.W., Coleman, J., Walker, L., Dunn, B., Takizawa,
P., Novick, P., and Ferro-Novick, S. (2003). Myo4p and She3p are
required for cortical ER inheritance in Saccharomyces cerevisiae. J.
Cell Biol. 163: 1255–1266.
Graumann, K., Irons, S.L., Runions, J., and Hawes, C. (2007).
Retention and mobility of the mammalian lamin B receptor in the
plant nuclear envelope. Biol. Cell 99: 553–562.
Grigoriev, I., Gouveia, S.M., van der Vaart, B., Demmers, J., Smyth,
J.T., Honnappa, S., Splinter, D., Steinmetz, M.O., Putney, J.W., Jr.,
Hoogenraad, C.C., and Akhmanova, A. (2008). STIM1 is a MT-plusend-tracking protein involved in remodeling of the ER. Curr. Biol. 18:
177–182.
Gupton, S.L., Collings, D.A., and Allen, N.S. (2006). Endoplasmic
reticulum targeted GFP reveals ER organization in tobacco NT-1 cells
during cell division. Plant Physiol. Biochem. 44: 95–105.
Hawes, C., Osterrieder, A., Hummel, E., and Sparkes, I. (2008). The
Plant ER-Golgi interface. Traffic 9: 1571–1580.
Hepler, P.K., and Wolniak, S.M. (1984). Membranes in the mitotic
apparatus: Their structure and function. Int. Rev. Cytol. 90: 169–238.
Irons, S.L., Evans, D.E., and Brandizzi, F. (2003). The first 238 amino
acids of the human lamin B receptor are targeted to the nuclear
envelope in plants. J. Exp. Bot. 384: 943–950.
Kachar, B., and Reese, T.S. (1988). The mechanism of cytoplasmic
streaming in characean algal cells: Sliding of endoplasmic reticulum
along actin filaments. J. Cell Biol. 106: 1545–1552.
Klopfenstein, D.R., Kappeler, F., and Hauri, H.P. (1998). A novel direct
interaction of endoplasmic reticulum with microtubules. EMBO J. 17:
6168–6177.
Knebel, W.H., Quader, H., and Schnepf, E. (1990). Mobile and immobile endoplasmic reticulum in onion bulb epidermis cells: Short- and
long-term observations with a confocal laser scanning microscope.
Eur. J. Cell Biol. 52: 328–340.
Langhans, M., Niemes, S., Pimpl, P., and Robinson, D. (2009).
Oryzalin bodies: In addition to its anti-microtubules properties, the
dinitroaniline herbicide oryzalin causes nodulation of the endoplasmic
reticulum. Protoplasma 236: 73–84.
Lee, C., and Chen, L.B. (1988). Dynamic behavior of endoplasmic
reticulum in living cells. Cell 54: 37–46.
Lichtscheidl, I.K., and Url, W.G. (1990). Organisation and dynamics of
cortical endoplasmic reticulum in inner epidermal cells of onion bulb
scales. Protoplasma 157: 203–215.
Lodish, H., Berk, A., Kaiser, C.A., Krieger, M., Scott, M.P., Bretscher,
A., Ploegh, H., and Matsudaira, P. (2008). Molecular Cell Biology.
(New York: W.H. Freeman and Co.).
Loverdo, C., Benichou, O., Moreau, M., and Voituriez, R. (2008).
Enhanced reaction kinetics in biological cells. Nat. Phys. 4: 134–137.
Cytoskeletal Control of Plant Endoplasmic Reticulum
Mirny, L. (2008). Cell commuters avoid delays. Nat. Phys. 4: 93–95.
Nebenführ, A., Gallagher, L.A., Dunahay, T.G., Frohlick, J.A.,
Mazurkiewicz, A.M., Meehl, J.B., and Staehelin, L.A. (1999).
Stop-and-go movements of plant Golgi stacks are mediated by the
acto-myosin system. Plant Physiol. 121: 1127–1141.
Peremyslov, V.V., Prokhnevsky, A.I., Avisar, D., and Dolja, V.V.
(2008). Two class XI myosins function in organelle trafficking and
root hair development in Arabidopsis thaliana. Plant Physiol. 146:
1109–1116.
Pickard, W.F. (2003). The role of cytoplasmic streaming in symplastic
transport. Plant Cell Environ. 26: 1–15.
Prokhnevsky, A.I., Peremyslov, V.V., and Dolja, V.V. (2008). Overlapping functions of the four class XI myosins in Arabidopsis growth,
root hair elongation and organelle motility. Proc. Natl. Acad. Sci. USA
105: 19744–19749.
Quader, H., Hofmann, A., and Schepf, E. (1987). Shape and movement
of the endoplamic reticulum in onion bulb epidermis cells:possible
involvement of actin. Eur. J. Cell Biol. 44: 17–26.
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.
Runions, J., Brach, T., Kuhner, S., and Hawes, C. (2006). Photoactivation of GFP reveals protein dynamics within the endoplasmic
reticulum membrane. J. Exp. Bot. 57: 43–50.
Saint-Jore, C.M., Evins, J., Batoko, H., Brandizzi, F., Moore, I., and
Hawes, C. (2002). Redistribution of membrane proteins between the
Golgi apparatus and endoplasmic reticulum in plants is reversible and
not dependent on cytoskeletal networks. Plant J. 29: 661–678.
Sheahan, M.B., Staiger, C.J., Rose, R.J., and McCurdy, D.W. (2004).
A green fluorescent protein fusion to actin-binding domain 2 of
Arabidopsis fimbrin highlights new features of a dynamic actin cytoskeleton in live plant cells. Plant Physiol. 136: 3968–3978.
Sparkes, I.A., Frigerio, L., Tolley, N., and Hawes, C. (2009a). The plant
endoplasmic reticulum: A cell-wide web. Biochem. J. 423: 145–155.
Sparkes, I.A., Hawes, C., and Baker, A. (2005). AtPEX2 and AtPEX10
are targeted to peroxisomes independently of known endoplasmic
reticulum trafficking routes. Plant Physiol. 139: 690–700.
3949
Sparkes, I.A., Ketelaar, T., Ruijter, N.C.A., and Hawes, C. (2009b).
Grab a Golgi: Laser trapping of Golgi bodies reveals in vivo interactions with the endoplasmic reticulum. Traffic 10: 567–571.
Sparkes, I.A., Runions, J., Kearns, A., and Hawes, C. (2006). Rapid,
transient expression of fluorescent fusion proteins in tobacco plants
and generation of stably transformed plants. Nat. Protoc. 1: 2019–
2025.
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.
Staehelin, A. (1997). The plant ER: A dynamic organelle composed
of a large number of discrete functional domains. Plant J. 11:
1151–1185.
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.
Tolley, N., Sparkes, I.A., Hunter, P.R., Craddock, C.P., Nuttall, J.,
Roberts, L.M., Hawes, C., Pedrazzini, E., and Frigerio, L. (2008).
Overexpression of a plant reticulon remodels the lumen of the cortical
endoplasmic reticulum but does not perturb protein transport. Traffic
9: 94–102.
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 35nm steps at high
velocity. EMBO J. 6: 1263–1272.
Ueda, K., Matsuyama, T., and Hashimoto, T. (1999). Visualisation of
microtubules in living cells of transgenic Arabidopsis thaliana. Protoplasma 206: 201–206.
Voeltz, G.K., Prinz, W.A., Shibata, Y., Rist, J.M., and Rapoport, T.A.
(2006). A class of membrane proteins shaping the tubular endoplasmic reticulum. Cell 124: 573–586.
Vedrenne, C., and Hauri, H.P. (2006). Morphogenesis of the endoplasmic reticulum: Beyond active membrane expansion. Traffic 7:
639–646.
Wolfram, S. (2002). A New Kind of Science. (Champaign, IL: Wolfram
Media).
Movement and Remodeling of the Endoplasmic Reticulum in Nondividing Cells of Tobacco Leaves
I. Sparkes, J. Runions, C. Hawes and L. Griffing
Plant Cell 2009;21;3937-3949; originally published online December 29, 2009;
DOI 10.1105/tpc.109.072249
This information is current as of June 18, 2017
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References
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