Download Dynamic Tubular Vacuoles Radiate Through the

New Dynamics in an Old Friend: Dynamic
Tubular Vacuoles Radiate Through the Cortical
Cytoplasm of Red Onion Epidermal Cells
Regular Paper
Elizabeth J. Wiltshire and David A. Collings∗
School of Biological Sciences, University of Canterbury, Private Bag 4800, Christchurch, New Zealand
The textbook image of the plant vacuole sitting passively
in the centre of the cell is not always correct. We observed
vacuole dynamics in the epidermal cells of red onion
(Allium cepa) bulbs, using confocal microscopy to detect
autofluorescence from the pigment anthocyanin. The
central vacuole was penetrated by highly mobile
transvacuolar strands of cytoplasm, which were also visible
in concurrent transmitted light images. Tubular vacuoles
also extended from the large central vacuole and radiated
through the cortical cytoplasm. These tubules were thin,
having a diameter of about 1.5 µm, and were connected to
the central vacuole as shown by fluorescence recovery
after photobleaching (FRAP) experiments. The tubules
were bounded by the tonoplast, as revealed by transient
expression of green fluorescent protein (GFP) targeted
to the vacuolar membrane and through labeling with the
dye MDY-64. Expression of endoplasmic reticulumtargeted GFP demonstrated that the vacuolar tubules
were distinct from the cortical endoplasmic reticulum.
Movement of the tubular vacuoles depended on actin
microfilaments, as microfilament disruption blocked
tubule movement and caused their collapse into
minivacuoles. The close association of the tubules with
GFP-tagged actin microfilaments suggests that the tubules
are associated with myosin, and that tubules likely move
along microfilaments. Tubular vacuoles do not require
anthocyanin for their formation, as tubules were also
present in white onion cells that lack anthocyanin. The
function of these tubular vacuoles remains unknown, but
as they greatly increase the surface area of the tonoplast,
they might increase transport rates between the cytoplasm
and vacuole.
Keywords: actin microfilaments • Allium cepa • anthocyanin
• onion epidermis • vacuolar tubules • vacuole
∗Corresponding
Abbreviations: ER, endoplasmic reticulum; FRAP,
fluorescence recovery after photobleaching; GFP, green
fluorescent protein; YFP, yellow fluorescent protein.
Introduction
Textbook images of the plant vacuole show a large,
static organelle, surrounded by the vacuolar membrane
(tonoplast), which sits passively in the centre of the cell. This
central vacuole performs multiple functions, and is important not only for the generation of turgor pressure but also
as a store of ions, metabolites and pigments, and as a site of
detoxification (Marty 1999). Recent research into vacuolar
structure has shown that the image of a single, passive central vacuole is not always accurate. Vacuoles are diverse:
while the central vacuole is acidic and lytic, other non-acidic
vacuoles can act as sites of storage, and different types of
vacuole can co-exist in a single cell (Swanson et al. 1998).
In most active and growing cells, transvacuolar strands penetrate the vacuole. Bounded by the tonoplast, these strands
contain cytoplasm, organelles such as endoplasmic reticulum (ER) and mitochondria, and the actin microfilaments
that drive both cytoplasmic streaming and the dynamic
reorganization of the strands (Parthasarathy et al. 1985, Kost
et al. 1998). The vacuoles of expanding Arabidopsis cotyledon epidermal (Saito et al. 2002) and guard (Tanaka et al.
2007) cells, and tobacco suspension culture cells (Reisen
et al. 2005) also contain membrane-bound inclusions.
Ripples have been reported in the surface of tobacco and
onion vacuoles (Verbelen and Tao 1998), and vacuoles can
also exist as tubules. These tubular vacuoles were first
observed in the margins of rose leaves. As the cells matured,
the accumulation of the red pigment anthocyanin rendered
tubular vacuoles visible, prior to their fusion to form the
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Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124, available online at www.pcp.oxfordjournals.org
© The Author 2009. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
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Tubular vacuoles in red onion epidermal cells
central vacuole (Guilliermond 1929). This development of
a large central vacuole from tubular pre-vacuoles has also
been demonstrated by electron microscopy in other cell
types (Marty 1978, Marty 1999). More recently, tubular vacuoles have also been observed in onion epidermal (Url 1964)
and guard (Palevitz and O’Kane 1981) cells, in Arabidopsis
pollen tubes (Hicks et al. 2004) and root hairs (Ovečka et al.
2005), and in cultured tobacco cells during cytokinesis
(Kutsuna et al. 2003).
This re-evaluation of vacuolar structure has relied on the
application of confocal microscopy and advances in green
fluorescent protein (GFP) and dye-based fluorescence
methods. While vacuolar-targeted GFP has been observed in
non-acidic tobacco protoplast vacuoles (di Sansebastiano
et al. 1998), expressing GFP in acidic vacuoles has been more
difficult: GFP could only be observed in acidic vacuoles of
Arabidopsis when it was allowed to accumulate in darkness
(Tamura et al. 2003). Thus, direct GFP visualization of the
vacuole is challenging. It is more reliably achieved by
targeting GFP constructs to the tonoplast through fusions
with tonoplast-resident proteins including tonoplast intrinsic proteins (TIPs) (Cutler et al. 2000, Saito et al. 2002), syntaxins such as AtVam3p (Kutsuna et al. 2003), aquaporins
(Cutler et al. 2000, Reisen et al. 2005) and cation transporters
(Delhaize et al. 2003).
There are three pathways that can be utilized to dye the
vacuole for fluorescence imaging. These include the endocytosis of the styryl dyes (FM4-64 and FM1-43) that initially
label endosomes but which label the tonoplast after several
hours (Emans et al. 2002, Ovečka et al. 2005, Tanaka et al.
2007), and of membrane-impermeant dyes such as Lucifer
Yellow (Hillmer et al. 1989, Reisen et al. 2005) and Alexa 568
hydrazide (Emans et al. 2002, Kutsuna et al. 2003) that may
also accumulate in the vacuole via endocytosis. Some membrane-permeant dyes naturally accumulate in acidic vacuoles because of charge effects. These include acridine orange
(Timmers et al. 1995, Verbelen and Tao 1998, Reisen et al.
2005), neutral red (Guilliermond 1929, Palevitz et al. 1981,
Timmers et al. 1995, di Sansebastiano et al. 1998, Reisen et al.
2005, Dubrovsky et al. 2006, Poustka et al. 2007), sulforhodamine (Canny 1987, D. Liu and L. Cantrill, personal communication) and Lysosensor yellow (Swanson et al. 1998).
Finally, some membrane-permeant dyes are chemically
modified in the cytoplasm into forms that are pumped into
the vacuole. These include fluorescein diacetate and its
derivates, which are de-esterified into their fluorescent,
anionic forms (Swanson et al. 1998, Kutsuna et al. 2003,
Reisen et al. 2005) and monochlorobimane, which reacts
with glutathione (Swanson et al. 1998, Reisen et al. 2005).
With the difficulties inherent in viewing vacuolar GFP,
and with dye-based approaches being problematic due to
difficulties in loading cells and the relative non-specificity of
many of the dyes, it is significant that the vacuole can also be
imaged directly through autofluorescence of constituent
molecules. These molecules include a range of flavonoids,
both colorless (Palevitz et al. 1981, Palevitz and O’Kane 1981,
H. Berg, personal communication) and the pigmented
anthocyanins (Poustka et al. 2007). As part of ongoing
research into the organization and structure of Allium epidermal cells, vacuoles were imaged using weak anthocyanin
fluorescence. The anthocyanin cyanidin-3-glucoside is the
predominant pigment in the epidermal cell layers of red
onion bulb scales (Donner et al. 1997) although at least 25
different anthocyanins have been reported (Slimestad et al.
2007). Our research has demonstrated vacuole dynamics,
and revealed thin tubular extensions of the central vacuole
that ramify through the cortical cytoplasm. These tubular
vacuoles remained connected to the central vacuole,
and their movement and structure depends upon actin
microfilaments and not microtubules. The function of these
vacuolar tubules remains unknown.
Results
Anthocyanin fluorescence reveals the red onion
vacuole
The vacuoles of red onion epidermal cells contain anthocyanins which can be used to visualize vacuole morphology
and dynamics (Fig. 1). Using 488-, 514- or 561-nm excitation,
we followed the dynamics of the central vacuole in cells in
inner epidermal peels. Numerous cytoplasmic strands
through the subcortex and vacuole were visible as dark, nonfluorescent regions (Fig. 1A, asterisks), as were nuclei
(arrows). Transmitted light images showing cellular organization and the large central vacuole were best collected with
red light (633 nm) as both blue and green laser light were
absorbed by the anthocyanin. However, by combining red,
green (561 nm) and blue (488 nm) images collected concurrently line by line, we generated pseudocolor transmitted
light images whose color matched that of cells viewed by eye
(Fig. 1A). Scans of white onion inner epidermal cells made
under similar imaging conditions showed no fluorescence
(see also Fig. 3B).
Fluorescence emission spectra for six different excitation
wavelengths ranging from 405 to 633 nm were collected by
scanning fluorescence emission wavelengths (λ scanning).
Normalized curves showed the same, single emission peak at
approximately 630 nm for all wavelengths in inner epidermal
cells suggesting the presence of only a single fluorophore
(Fig. 1B). By correlating the relative absorbance of transmitted green light at 561 nm, we confirmed that there was
a linear relationship between the intensity of the red anthocyanin pigmentation and the intensity of fluorescence, with
reddest cells being most fluorescent (correlation visible in
Fig. 1A). Our data do not prove that the single fluorescent
emission peak at 630 nm relates directly to red anthocyanins,
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Fig. 1 Anthocyanin autofluorescence reveals vacuolar dynamics in inner epidermal cells. (A) Using cyan excitation (514 nm), anthocyanin
autofluorescence (580–780 nm; anthocyanin, upper image) demonstrated vacuolar morphology. Non-fluorescent transvacuolar strands and
aggregates of cytoplasm (asterisks) were dynamic while dark regions representing nuclei (arrows) were stationary. Concurrent transmitted light
imaging with the 514-nm laser was difficult because of anthocyanin absorbance. However, addition of red (633 nm), blue (488 nm) and green
(561 nm) transmitted light images generated a full color image (bright-field, lower image). (B) Red onion epidermal cells contain a compound
that fluoresced with excitation in the visible spectrum. Emission spectra (λ scans) recorded at six different excitation wavelengths from 405 to
633 nm showed only a single, broad emission peak at 630 nm. Normalized emission curves for seven cells shown in (A) were averaged, with the
curve for 633 nm excitation adjusted as this overlapped with the emission maximum. Outer epidermal cells gave comparable data. (C) Higher
magnification images of the region shown boxed in (A). Optical sections at 5 µm intervals through the epidermal cells reveal the presence of
dynamic vacuolar tubules (asterisk) whereas tubules associated with nuclei (arrows) were less dynamic. (D) A red/green stereo surface
reconstruction of the optical series shown in (C) created in ImageJ. This image demonstrates variations in the density of vacuolar tubules,
and that the cells with more anthocyanin as seen in (A), have more tubules. (E) While tubules were visible in fluorescence (anthocyanin, left),
they were not usually visible in transmitted light images although on occasions there were suggestions of similar structures (bright-field, right).
Bars: (A) 100 µm; (C) 20 µm for (C) and (D), (E) 20 µm.
as this peak might result from some minor anthocyanin
component or a reaction intermediate in the anthocyanin
biosynthetic pathway. However, as fluorescence was used
solely to investigate vacuolar organization, we use the
term ‘anthocyanin fluorescence’ to refer to this 630 nm
emission peak.
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Higher magnification images of the vacuole revealed
hitherto unsuspected complexity in its structure. Optical
sectioning through the epidermal cell showed extensive
tubules that ramified through the cortical cytoplasm
[Fig. 1C; Movie 1 (Supplementary data)]. These structures
were distinct from the ripples in the vacuolar surface
Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009.
Tubular vacuoles in red onion epidermal cells
previously reported by Verbelen and Tao (1998) as they were
separated completely from the surface of the central vacuole. This was most notable adjacent to the nucleus (Fig. 1C,
arrows) but distinct tubules were present across much of
the surface of the vacuole. These tubules were thin, having
a diameter of about 1.5 µm, and accounted for only a small
percentage of total vacuole volume. Two forms of tubule
were visible. Stable reticulate arrays of tubules were common
around the nucleus and in regions where cytoplasmic
streaming was less dynamic (Fig. 1C, arrows) while dynamic
tubules were present in the cell cortex where they were often
associated with subcortical strands of cytoplasm (Fig. 1C
asterisks; see also Fig. 2). Interestingly, cells with higher levels
of anthocyanin also had a greater number of tubules, as
shown by a comparison of tubule numbers in the lighter and
darker cells in the color transmitted light image (Fig. 1A)
and in a stereo reconstruction of the cells (Fig. 1D).
Although tubules were not generally visible with the
transmitted light system on our confocal microscope, on
some occasions transmitted light images showed suggestions of structures matching tubule patterns (Fig. 1E) and
we suspect that a system optimized for light microscopy
might resolve them.
Most epidermal cells from all developmental stages of
bulb leaves contained vacuolar tubules in both the inner
and outer epidermis. Fig. 2 and Supplementary Movie 2,
taken from a region adjacent to the nucleus of an outer
epidermal cell from the outermost leaf of an onion bulb,
show tubule dynamics over several minutes. This cell showed
several examples of rapidly translocating vacuolar tubules
(arrows) moving along subcortical cytoplasmic strands
(dark strands, lacking anthcyanin fluorescence). Some cells,
however, were less dynamic, contained few transvacuolar
cytoplasmic strands and generally lacked visible tubules.
Fig. 2. Anthocyanin autofluorescence revealed vacuolar dynamics in
the outer epidermis of the outermost living leaf of a red onion bulb.
Time-course of tubule dynamics in the outer cortex, adjacent to the
nucleus showing four images each separated by 60 s. Arrows indicate
vacuolar tubules that lay within cytoplasmic strands. Bar, 20 µm.
Tubular vacuoles and other organelles
We used a combination of fluorescent dyes and transient
expression of GFP fusion proteins to compare the organization of tubular vacuoles with the other organelles present in
epidermal cells. When cytosolic yellow fluorescent protein
(YFP) was transiently expressed in outer epidermal cells,
organelles that excluded YFP appeared as dark regions. These
organelles consisted of small, round to elongate structures
that were not identified but which might be mitochondria,
peroxisomes and oil droplets, and long tubular structures.
These matched the location of anthocyanin-containing
tubular vacuoles (Fig. 3A, arrows). Similar tubular structures
lacking cytosolic YFP were also seen in inner epidermal cells
of white onions, showing that vacuolar tubules do not
require anthocyanin to form (Fig. 3B). We also used particle
Fig. 3 Tubular vacuoles were observed as structures that excluded
cytoplasmic labels. (A) Transiently expressed cytosolic YFP and
anthocyanin imaged concurrently in an outer epidermal cell. Paired
arrows indicate locations of long vacuolar tubules from which YFP was
excluded. Smaller dark regions indicate other organelles such as
mitochondria. (B) Similar long tubules excluding YFP were observed in
the inner epidermis of white onion cells, suggesting that these may
also contain vacuolar tubules. Imaging conditions were identical to
(A) and showed no anthocyanin fluorescence. (C) Oregon green
conjugated to 70 kDa dextran was delivered to the cytoplasm of an
outer epidermal cell using particle bombardment. At the surface of
the cell, vacuolar tubules were clearly observed as regions excluding
the cytosolic dye. An optical section 15 µm further into the cell shows
that the Oregon green–dextran conjugate is excluded from the
nucleus (n) and that tubules exclude the cytoplasm. Bar in (C) 20 µm
for all images.
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bombardment of Oregon green conjugated to 70 kDa dextran
(Iglesias and Meins 2000) to load dye into the cytoplasm of
outer epidermal cells. Again, tubular vacuoles excluded dye.
As the 70 kDa dextran was excluded from the nucleus, this
allowed the vacuolar tubules around the nucleus to be
observed as distinct from perinculear cytoplasmic strands
(Fig. 3C; Supplementary Movie 3).
The organization of the tubular vacuole system is distinct
from the other cellular systems. Although the ER is superficially similar to tubular vacuoles, being also a combination of
stable reticulate arrays and rapidly streaming subcortical
strands, expression of GFP–HDEL in outer epidermal cells
showed that the two networks were distinct, with the tubular vacuoles having a less branched pattern than reticulate
cortical ER (Fig. 4; Supplementary Movie 4). However,
extensive subcortical ER strands often lay parallel and close
to vacuolar tubules (arrows).
The vacuole is surrounded by the vacuolar membrane or
tonoplast. We successfully counterstained the tonoplast
with MDY-64, a novel dye that has previously been used to
label the tonoplast of yeast vacuoles (Cole et al. 1998).
MDY-64 labeled the tonoplast around the central vacuole,
and also the plasma membrane where it revealed potential
pit-fields (Fig. 5A, asterisks). MDY-64 also labeled a mass of
closely furled membranes that correspond to the vacuolar
tubules seen with anthocyanin fluorescence (arrows). Labeling of the tonoplast around vacuolar tubules was also confirmed with expression of ShMTP1–GFP (Delhaize et al.
2003). This construct labeled the tonoplast around tubules
in the outer cortex of inner epidermal cells (Fig. 5B;
Supplementary Movie 5), although it also labeled unidentified and highly dynamic organelles. GFP expression also
demonstrated that tubular vacuoles were distinct from both
mitochondria and the Golgi apparatus (data not shown).
Fig. 4 An outer epidermal cell transiently expressing ER-targeted GFP (GFP–HDEL) was imaged with low laser power at 488 nm sequentially (line
by line) with excitation of anthocyanin with high laser power at 561 nm. Single optical sections at three different time points (times in seconds)
demonstrate differences between cortical ER organization and vacuolar tubules. Subcortical strands of ER that lie parallel to vacuolar tubules in
cytoplasmic strands are indicated (arrows). Bar, 20 µm.
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Tubular vacuoles in red onion epidermal cells
Fig. 5 Vacuolar tubules are enclosed by the tonoplast. (A) Inner epidermal cells were labeled with the dye MDY-64 (10 µM, 10 min), washed for
10 min, and confocal optical sections recorded for anthocyanin and MDY-64 fluorescence. The MDY-64 labeled the plasma membrane and
tonoplast around the central vacuole, and often showed four parallel lines (black arrows) although occasionally the inner two lines that represent
the plasma membrane fused (asterisks). Transmitted light images, which appear darker as they were collected with the 568-nm laser which is
strongly absorbed by the anthocyanin, suggest that these fusion sites were pit-fields where the cell wall was thinner. The MDY-64 also labeled
a confusing mass of membranes that corresponded to the vacuolar tubules (arrows) thus demonstrating that the tubules were indeed surrounded
by the tonoplast. (B) The tonoplast labeling construct ShMTP1–GFP was transiently expressed in inner epidermal cells. Imaging in the outer
cortex showed that as anthocyanin-containing tubules moved (times indicated in seconds in ShMTP1–GFP images), tubes that were faintly
visible with the ShMTP1–GFP construct moved in similar patterns. The ShMTP1–GFP also labeled highly dynamic punctate organelles whose
identity was not established but which might be pre-vacuolar compartments. These observations confirmed that the tubules are surrounded
by the vacuolar membrane. A cluster of gold particles present in the outer cortex is indicated with an asterisk. Bar in (B) 20 µm for both images.
Tubular vacuoles may be induced by confocal
imaging
Vacuolar tubules often increased in number and complexity
during confocal imaging, suggesting that their presence is,
in part, due to laser irradiation (Fig. 6; Supplementary
Movie 5). In this example from the outer epidermis, few
tubules were present as imaging began but there were
numerous non-fluorescent inclusions within the vacuole
(Fig. 6A, arrow). As imaging progressed, these inclusions
quickly disappeared, and extensive tubulation developed in
the outer cortex within 7 min. High-magnification images of
the tubular vacuoles suggest that they originate from the
central vacuole (Fig. 6B, arrow). Furthermore, image
sequences demonstrate that the tubules can elongate from
either the central vacuole (asterisk at 100 s) or other tubules
(arrowhead at 110 s) before eventually fusing with other
tubules to increase the complexity of the network. The fate
of the vacuolar inclusions as they disappeared was not
resolved in these sequences, and whether fusion of inclusions with the central vacuolar membrane provided the
necessary increase in membrane content for the formation
of the tubules was not determined.
Where tubules were induced by imaging, they remained
present at least 30 min after imaging had ceased (data not
shown). Nevertheless, examples existed where tubules were
prominent at the start of imaging, where imaging did not
induce vacuolar tubules and where, on occasions, tubules
decreased during imaging. An extensive survey of different
imaging wavelengths (488, 514 and 561 nm) and laser intensities showed induction to be inconsistent.
Vacuolar tubules are connected to the
central vacuole
High-magnification images of the tubular vacuoles suggest
that they originate from the central vacuole, and this was
confirmed by running fluorescence recovery after photobleaching (FRAP) experiments (Fig. 7; Supplementary
Movie 6). As anthocyanin was resistant to photobleaching,
Oregon green–dextran was loaded into the vacuole of
outer epidermal cells by particle bombardment where its
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Fig. 6 Confocal imaging may induce the formation of vacuolar tubules. (A) Continuous imaging over 7 min resulted in a significant increase in the
presence of vacuolar tubules, and the concurrent loss of small, non-fluorescent inclusions within the vacuole (arrow at 1 min). Numbers indicate
times in minutes after the start of imaging. (B) Higher magnification view of the cell showing the area around the nucleus that is boxed in (A) at
10-s intervals, beginning at 90 s. Tubules around the nucleus appear to pull away from the surface of the central vacuole (arrow), implying that
the tubular network and the central vacuole are connected. A tubule also grows from the surface of the central vacuole, beginning at 100 s, before
fusing with another tubule (asterisk), while another tubule branches and grows away from the tubules around the vacuole (arrowhead at 110 s).
Bars: (A) 20 µm; (B) 10 µm.
fluorescence precisely matched anthocyanin fluorescence in
both the central vacuole and tubules (Fig. 7A). Irradiating a
small region of tubules caused the rapid bleaching of Oregon
green but not anthocyanin (Fig. 7B). This loss was reversible,
with Oregon green fluorescence in the tubules recovering
within 10 s of high-intensity bleaching ceasing (Fig. 7A, C).
As anthocyanin did not bleach under these conditions, it
provided a reference for tubule structure (Fig. 7A, B, D).
Similar experiments were conducted on anthocyanin-free
inner epidermal cells which not only confirmed the presence
of tubules in these cells in the absence of anthocyanin, but
also demonstrated that these tubules were connected to the
central vacuole.
Tubular vacuoles are retained during plasmolysis
Tubules were unaffected during plasmolysis with 0.5 M
sucrose, and remained during the subsequent recovery of
cells in distilled water (Fig. 8). Cytoplasmic streaming
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continued during plasmolysis, although in some cases it was
lost in areas of cells. In these locations, tubular vacuoles
collapsed into round, non-dynamic mini-vacuoles. Tubular
vacuoles were not observed inside Hechtian strands,
although anthocyanin’s weak fluorescence and the thin
nature of these strands would impede detection of any
tubular vacuoles present there.
Tubular vacuoles require actin microfilaments
and not microtubules
The dynamism and structure of tubular vacuoles depends
on actin microfilaments and not microtubules. In epidermal
peels incubated with the actin-binding compound latrunculin B (2 µM), cytoplasmic streaming ceased within 5 min
and tubular vacuoles collapsed into small, round minivacuoles (Fig. 9A, B). Expression of the actin-labeling probes
GFP–hTalin and GFP–fABD2 in both inner and outer
epidermal cells demonstrated that dynamic and elongated
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Tubular vacuoles in red onion epidermal cells
Fig. 7 FRAP showed that the tubular vacuoles are connected to the central vacuole. (A) Oregon green conjugated to 70 kDa dextran (central
image, green in overlay) was loaded into the vacuole of an individual cell by particle bombardment and showed a similar pattern to anthocyanin
fluorescence found in all the epidermal cells (top image, red in overlay). A representative pair of images taken after 30 s of continual imaging with
moderate laser power is shown (labeled –30) immediately prior to the bleaching of a small region of tubules using full laser power at 514 and
561 nm for 30 s (boxed green). Selected images are also shown for the first 30 s of recovery. The rapid recovery of Oregon green fluorescence
within the tubules immediately after photobleaching indicates that the tubules are connected to the central vacuole. Times shown in the overlay
image are in seconds after the cessation of bleaching. (B) During 30 s of bleaching, anthocyanin (Anth.) remained unbleached while Oregon
green–dextran (OG) bleached within the first 10 s. (C) Oregon green and (D) anthocyanin intensities were recorded for two regions of the cell,
marked 1 (central vacuole) and 2 (tubules) in (A). While anthocyanin fluorescence was undiminished by 30 s of full laser power, Oregon green
bleached but recovered rapidly. Oregon green fluorescence also decreased in the central vacuole, consistent with a slow bleaching of the entire
vacuolar contents of the dye, and with diffusion into tubules. Bars: (A) 20 µm; (B) 10 µm.
Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009.
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Fig. 8 Vacuolar tubules survive plasmolysis.
Inner epidermal cells containing
anthocyanin were plasmolysed with 0.5 M
sucrose. Tubules, which had developed
over 20 min imaging, remained present
during 30 min in sucrose as the cell
plasmolysed. Times after the addition of
sucrose are indicated in minutes.
Fluorescence images using 514-nm
excitation and transmitted light images
(633 nm excitation due to absorbance of
green and blue laser light by anthocyanin)
were recorded sequentially. Bar, 50 µm.
Fig. 9 Tubule dynamics depends on actin microfilaments and not microtubules. (A) and (B) In cells exposed to 2 µM latrunculin, vacuolar tubules
collapsed into spherical mini-vacuoles. (A) shows an entire cell prior to drug addition while (B) shows inset images 20 s apart taken at 0, 2 and
4 min. Changes in tubule patterns indicate cytoplasmic streaming slowed by 100 s and ceased by 210 s. (C) Transient expression of GFP–hTalin.
Tubular vacuoles often lay parallel to and closely associated with the labeled actin microfilament bundles; arrows indicate a tubule that lies
beside an actin bundle. (D) and (E) In cells treated with 10-µM oryzalin, tubular vacuoles were not affected. (D) The entire cell. (E) Inset from (D),
showing images 20 s apart at about 1, 10 and 18 min after oryzalin addition. Changes in pattern indicate cytoplasmic streaming and tubular
vacuole motility. (F) Transient expression of GFP–MBD. Tubular vacuoles formed patterns distinct from the labeled cortical microtubules.
Bars: in (A) 50 µm for (A), (B), (D) and (E); in (F) 20 µm for (C) and (F).
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Tubular vacuoles in red onion epidermal cells
vacuolar tubules were often closely associated with and
parallel to bundled actin microfilaments (Fig. 9C, arrows;
Supplementary Movie 7). By contrast, epidermal peels
incubated with the microtubule depolymerizing herbicide
oryzalin (20 µM, 20 min) showed no changes to cell streaming or tubule dynamics (Fig. 9D, E) and tubules did not colocalize with microtubules labeled with the marker GFP–MBD2
(Fig. 9F).
Discussion
Onion epidermal peels have been extensively used as a model
research system for the last century. Initial observations of
plasmolysis (Hecht 1912), innovative observations of endomembrane structure (Url 1964, Lichtscheidl and Weiss 1988)
and the earliest observations of plant ER using DIOC6(3)
(Quader and Schnepf 1986, Quader et al. 1989) were all conducted in the onion epidermis. In recent years, onion epidermal cells have become a model system for testing GFP fusion
protein expression by particle bombardment (Silverstone
et al. 1998, von Arnim et al. 1998). With this research focus on
the onion epidermis, it is remarkable that while tubular
vacuoles were reported in onion epidermal cells (Url 1964),
they have not been subsequently investigated.
Viewing the plant vacuole using anthocyanin
autofluorescence
Numerous methods have been applied to view the vacuole.
These include GFP labeling of the tonoplast (Cutler et al. 2000,
Saito et al. 2002) and of the vacuole itself (di Sansebastiano
et al. 1998, Tamura et al. 2003), and the use of fluorescent
dyes. These dyes have been used to view both the vacuole and
vacuolar tubules, with uptake occurring via three distinct
routes. Labeling pathways include the endocytosis of membrane-labeling or membrane-impermeant dyes. Alternatively,
dyes can label the vacuole based on accumulation there
through pH and charge effects, or through chemical modification in the cytoplasm and sequestration of the product in the
vacuole.
In this study, we describe three different ways in which
the vacuole and tubular vacuoles can be studied in the onion
epidermis. The dye MDY-64 labels the tonoplast and reveals
the complexity of vacuolar organization and, although it has
only rarely been used to label the vacuole in higher plants
(Abrahams et al. 2003), it has been used to image vacuolar
tubules in tomato trichomes (Gunning 2007). We also bombarded dyes into the vacuole with a gene gun (Iglesias and
Meins 2000) but despite confirming the presence of vacuolar
tubules in anthocyanin-free cells and studying tubules with
FRAP, we found bombardment to be an unreliable delivery
method. We also used anthocyanin fluorescence to view
tubules: anthocyanin pigmentation has previously been used
to view tubular vacuoles in rose leaves (Guilliermond 1929)
while autofluorescent flavonoids have also been used to
image tubular vacuoles in onion guard cells (Palevitz and
O’Kane 1981, Palevitz et al. 1981) and soybean root cells
(H. Berg, personal communication).
The different fluorescence methods for studying the
vacuole each have advantages and disadvantages, and anthocyanin is no exception to this. Anthocyanin’s primary
advantage is that it is produced by the cell while its main
disadvantage is that high levels of laser irradiation are
required to generate significant fluorescence. Although the
fluorescence properties of the different Allium anthocyanins
have not been measured, fluorescent anthocyanins have
their strongest excitation in the ultraviolet (Drabent et al.
1999) rather than in the visible wavelengths more suitable
for live cell imaging. For this reason, the imaging of anthocyanin in the onion vacuole requires high levels of irradiation. For example, when 488-or 514-nm excitation was used
to view vacuolar tubules, transiently expressed GFP, which is
normally relatively stable to excitation light, is bleached.
The only way that GFP can be imaged along with anthocyanin is sequential scanning, exciting the anthocyanin with
high power at 561 nm while using lower irradiation at 488 nm
for GFP.
While it is possible to image the vacuole of onion epidermal cells using high excitation intensities without causing
apparent damage, cellular changes do occur. Anthocyanin
strongly absorbs both blue and green light (Fig. 1A), and the
continued use of high-powered lasers to excite the anthocyanin in the vacuole will result in significant heating of the
cells during time-course experiments. Cytoplasmic streaming rates increase at higher temperatures (Shimmen and
Yoshida 1993), and although not quantified in this study,
streaming rates did appear to increase after prolonged
imaging. The continued presence of streaming within the
onion epidermal cells would indicate that they remained
healthy during the experiments. However, in other cell types,
high light intensities are known to cause cellular damage
or induce cell death, possibly through the generation of
reactive oxygen species (Bartosz 1997, Dixit and Cyr 2003).
Tubular vacuoles are present in the cortex of onion
epidermal cells
Both the inner and outer epidermal cells of red onions contain tubular extensions from the central vacuole that extend
through the cortical cytoplasm, as demonstrated by anthocyanin autofluorescence. Dye studies, bombardments and
transient expression of GFP fusion proteins suggest that similar structures are present in onion cells lacking anthocyanin.
FRAP observations demonstrated that these tubules connect to the central vacuole, while GFP fusion proteins and
the dye MDY-64 confirmed that they are surrounded by the
tonoplast. The tubules undergo rapid and continuous cytoplasmic streaming. Unlike the moss Physcomitrella, where
Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009.
1835
E. J. Wiltshire and D. A. Collings
tubular vacuole dynamics are mediated by microtubules
(Oda et al. 2009), the dynamics of onion tubular vacuoles are
not regulated by microtubules: microtubule depolymerization did not modify tubule dynamics or structure and there
was no alignment between tubules and the cortical microtubule cytoskeleton. Instead, vacuolar tubules rely on the actin
cytoskeleton for their mobility and organization. This was
demonstrated by microfilament disruption with latrunculin,
which caused tubules to collapse into spherical minivacuoles, and by the coalignment of tubules with GFP–hTalinand GFP–fABD2-labeled microfilament bundles.
The structural dependence of higher plant vacuolar
tubules on actin and myosin has been observed in other
systems (Verbelen and Tao 1998, Ovečka et al. 2005, Higaki
et al. 2006) and is similar to the actin-based motility of other
organelles including the Golgi apparatus (Nebenführ et al.
1999), mitochondria (van Gestel et al. 2002) and peroxisomes
(Collings et al. 2002). However, while these other organelles
interact directly with microfilaments through myosin motors
attached to the organelle surface (Hashimoto et al. 2005,
Li and Nebenführ 2007, Riesen and Hanson 2007), interactions between myosin and the higher plant tonoplast have
not been demonstrated. Immunolabeling experiments in
Allium epidermal cells demonstrated myosin labeling of
organelles and disrupted endomembranes (Liebe and Quader
1994), but did not distinguish between myosin associated
with ER and tonoplast. Thus, while the close association of
tubules with microfilaments would suggest that it is likely
that myosin-associated tubules actively move along microfilaments, our results do not prove this and it remains possible that tubule dynamics are controlled by bulk flow of the
cytoplasm.
The tubules seen by anthocyanin fluorescence in red onion
epidermal cells and by other methods in non-anthocyanincontaining cells, are similar to the structures observed by transmitted light in white onion epidermal cells (Url 1964). As our
imaging system was optimized for confocal fluorescence
microscopy rather than transmitted light, we did not consistently observe tubules by light microscopy. It would also seem
likely that these tubules are similar to the ‘ripples’ in the vacuolar surface of tobacco and onion epidermal cells reported by
Verbelen and Tao (1998); with the reduced z-resolution of confocal microscopes, perhaps these ripples were in fact tubules
running adjacent to the surface of the vacuole.
Most other reports of vacuolar tubules are associated
with specialist cell types including trichomes (Lazzaro and
Thomson 1996, Gunning 2007), root hairs (Ovečka et al.
2005), pollen tubes (Hicks et al. 2004), developing guard cells
(Palevitz and O’Kane 1981, Palevitz et al. 1981) or cells in
various forms of culture (Hillmer et al. 1989, Newell et al.
1998, Kutsuna et al. 2003). Vacuolar tubules are only rarely
reported in non-differentiated cells. There are, however,
several exceptions to this. D. Liu and L. Cantrill (personal
1836
communication) have observed extensive vacuolar tubules
in rice tapetum cells. Further, soybean root meristematic
cells have an extensive tubular vacuolar network, rendered
visible by the accumulation of isoflavonoids following
Agrobacterium infection (Gunning 2007, H. Berg, personal
communication).
Are vacuolar tubules induced, and do they have
a specific function?
Vacuolar tubules have been suggested to play functional
roles in specific systems. For example, chickpea trichomes
contain a continuum of connected vacuoles, ranging from
large central vacuoles at the base of the trichome to tubular
vacuoles near the tip. These vacuolar tubules are thought to
effect the transport of sugars and metabolites to the secretory tip of the cell (Lazzaro and Thomson 1996). Similarly,
tubular vacuoles are commonly found in fungal hyphae
(Rees et al. 1994) where they are suggested to function in
nitrogen and phosphorus storage and transport (Ashford
2002). Functional roles for tubular vacuoles need not exist,
however, in systems where they represent a stage in vacuolar
development (Guilliermond 1929, Palevitz and O’Kane 1981,
Newell et al. 1998).
We have considered various possibilities for roles of these
tubules in mature onion bulb epidermal cells. As a role in
nutrient transport would seem unlikely in a storage tissue
such as onion, other functions were considered. Anthocyanins are generated on the cytoplasmic face of the ER and
transported to the vacuole, where they are thought to provide various protective roles to the cell (Gould 2004,
Grotewold 2006). Vacuolar tubules might protect the cytoplasm against reactive oxygen species by allowing them to
be accessed by anthocyanins. However, as tubules are present in anthocyanin-free cells, this would seem unlikely to be
the primary role for the tubules.
Our interpretation of the apparent but inconsistent
induction of tubules by imaging does not relate directly
to specific light-induced effects. While light does cause
a rapid rearrangement of anthocyanin-containing vacuolar
inclusions in anthocyanin-accumulating vacuoles of maize
cells (Irani and Grotewold 2005), we consider that tubule
induction is most likely caused by an increase in cytoplasmic
streaming in the cells that resulted from temperature
increases due to laser absorption by the anthocyanin.
Tubules as a by-product of cytoplasmic streaming rather
than a functionally specific component of the anthocyanincontaining vacuole would also explain why tubules can be
found in anthocyanin-free cells, and why they are present in
cells immediately on imaging. It is not, however, impossible
that tubules do play a role in vacuole functioning for their
presence does greatly increase the surface are of the tonoplast, and would thus significantly increase the rate of transport between the cytoplasm and the vacuole.
Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009.
Tubular vacuoles in red onion epidermal cells
Conclusions
Contrary to the textbook image of the vacuole as a passive
sac, we have shown it to have dynamic tubular extrusions
that extend through the cytoplasm in onion epidermal cells.
While the function of these tubules has yet to be elucidated,
their presence is scientifically interesting and provides
a powerful example that novel observations remain possible
when new technology is applied to a well-studied system.
Materials and methods
Plant material
Red and white onion bulbs (Allium cepa L.) were purchased
from local markets. Observations of outer epidermal cells
were made using whole sections of leaf (about 10 mm
square). Anthocyanin-containing inner epidermal cells were
found towards the upper and lower ends of the bulb, but
would develop throughout the inner epidermis if 20–30-mm
squares of leaf were excised and kept moist and in the light
for several days. For uptake of fluorescent molecules and
inhibitory drugs, inner epidermal peels were floated on solutions made in distilled water.
Stains and inhibitors
Stock solutions that were prepared in dimethyl sulfoxide
(DMSO) included latrunculin B (2 mM; MP BioMedicals,
Sydney, NSW, Australia), oryzalin (20 mM; Lilley, Greenfield,
IN, USA) and MDY-64 (10 mM; Invitrogen, Carlsbad, CA,
USA). Stocks were stored at –20°C and diluted in distilled
water to 2 µM, 20 µM and 10 µM, respectively.
Particle bombardment for dye loading and transient
GFP expression
A particle inflow gun (Finer et al. 1992) (Kiwi Scientific, Levin,
New Zealand) was used for delivery of dye and DNA into
epidermal cells. For biolistic delivery of dyes (Iglesias and
Meins 2000), Oregon green conjugated to 70 kDa dextran
(Invitrogen) (0.5 mM, 20 µl) was mixed with 4 mg of washed
1.1-µm diameter tungsten particles (BioRad, Hercules, CA,
USA) and sonicated (20 min). Resuspended particles (4 µl)
were dried onto 13-mm Swinnex filter holders (Millipore,
Billerica, MA, USA), which were used as particle carriers
inside the delivery system and shot into epidermal cells using
a 30 ms pulse of 120 psi He gas. Cells were incubated overnight before viewing.
For biolistic delivery of plasmid DNA (Collings et al. 2002),
1 mg of 1.6-µm diameter gold particles were coated with
1 µg of plasmid DNA by calcium precipitation, stabilized
with spermidine and resuspended in ethanol. Resuspended
particles (10 µl) were dried onto filter carriers and shot into
epidermal cells using 60 psi He gas, and viewed after 8–24 h.
Constructs used included cytosolic YFP, GFP–HDEL, which
labeled the ER (Haseloff et al. 1997), GFP–MBD, which labeled
microtubules (Marc et al. 1998), GFP–hTalin and GFP–
fABD2, which labeled actin microfilaments (Takemoto et al.
2003, Sheahan et al. 2004), and ShMTP1–GFP (Delhaize et al.
2003), which labeled the tonoplast.
Microscopy
All experiments were conducted on an inverted confocal
microscope (Leica model SP5; Wetzlar, Germany) using
a 20× NA 0.7 glycerol immersion lens. Anthocyanin was
excited with 488-, 514- or 561-nm laser lines with emission
generally captured from 580 to 780 nm. As anthocyanin is
not strongly fluorescent, imaging required high laser power.
Fortunately, anthocyanin resists photobleaching. GFP and
Oregon green were excited at 488 nm with emission recorded
from 500 to 550 nm using lower laser power to avoid fluorophore bleaching and, if required, anthocyanin was concurrently imaged using 561 nm excitation at high laser power.
The dye MDY-64 was excited with 458 nm and fluorescence
collected from 470 to 535 nm. Transmitted light images
were routinely recorded with bright-field optics. These could
be generated as pseudocolor images by combining red
(631 nm), green (561 nm) and blue (488 nm) transmitted
light images with laser power adjusted such that background
transmitted light was even.
Three dimensional images (red/green anaglyphs) were
prepared in ImageJ (National Institutes of Health, Bethesda,
MD, USA; available at http://rsb.info.nih.gov/ij). Images were
adjusted in Adobe Photoshop using standard brightness,
contrast and gamma tools.
Supplementary Data
Supplementary Data are available at PCP Online.
Funding
This work was supported by a University of Canterbury College of Science research grant to D.A.C., and by a University
of Canterbury Summer Research Scholarship to E.J.W.
Acknowledgments
We thank Manfred Ingerfeld for assistance with confocal
microscopy, Manny Delhaize (CSIRO Plant Industry), Daigo
Takemoto (Australian National University) and Jan Marc
(Sydney University) for supplying GFP fusion protein constructs, and Brigitta Kurenbach (University of Canterbury) for
assistance with bacterial transformations and plasmid purification. We also thank Mark Lazzaro (College of Charleston),
Brian Gunning (Australian National University), Howard Berg
(Danforth Center) and Danny Liu (Sydney University) for discussions. Rosemary White (CSIRO Plant Industry) suggested
using MDY-64 while John Dalrymple-Alford (University of
Canterbury) kindly supplied tungsten particles and suggestions on particle bombardment of dyes into cells.
Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009.
1837
E. J. Wiltshire and D. A. Collings
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(Received July 26, 2009; Accepted September 1, 2009)
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