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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 author: E-mail, [email protected]; Fax, +64 (3) 364 2590. 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. All rights reserved. For permissions, please email: [email protected] 1826 Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 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, Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 1827 E. J. Wiltshire and D. A. Collings 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. 1828 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. Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 1829 E. J. Wiltshire and D. A. Collings 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. 1830 Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 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 Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 1831 E. J. Wiltshire and D. A. Collings 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 1832 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 Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 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. 1833 E. J. Wiltshire and D. A. Collings 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). 1834 Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 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 References Abrahams, S., Lee, E., Walker, A.R., Tanner, G.J., Larkin, P.J. and Ashton, A.R. (2003) The Arabidopsis TDS4 gene encodes leucoanthocyanidin dioxygenase (LDOX) and is essential for proanthocyanidin synthesis and vacuole development. Plant J. 35: 624–636. Ashford, A.E. (2002) Tubular vacuoles in arbuscular mycorrhizas. New Phytol. 154: 545–547. Bartosz, G. (1997) Oxidative stress in plants. Acta Physiol. Plant. 19: 47–64. Canny, M.J. (1987) Locating active proton extrusion pumps in leaves. Plant Cell Environ. 10: 271–274. Cole, L., Orlovich, D.A. and Ashford, A.E. (1998) Structure, function, and motility of vacuoles in filamentous fungi. Fungal Genet. Biol. 24: 86–100. Collings, D.A., Harper, J.D.I., Marc, J., Overall, R.L. and Mullen, R.T. (2002) Life in the fast lane: actin-based motility of plant peroxisomes. Can. J. Bot. 80: 430–441. Cutler, S.R., Ehrhardt, D.W., Griffitts, J.S. and Somerville, C.R. (2000) Random GFP::cDNA fusions enable visualization of subcellular structures in cells of Arabidopsis at a high frequency. Proc. Natl Acad. Sci. USA 97: 3718–3723. Delhaize, E., Kataoka, T., Hebb, D.M., White, R.G. and Ryan, P.R. (2003) Genes encoding proteins of the cation diffusion facilitator family that confer manganese tolerance. Plant Cell 15: 1131–1142. di Sansebastiano, G.-P., Paris, N., Marc-Martin, S. and Neuhaus, J.-M. (1998) Specific accumulation of GFP in a non-acidic vacuolar compartment via a C-terminal propeptide-mediated sorting pathway. Plant J. 15: 449–457. Dixit, R. and Cyr, R.J. (2003) Cell damage and reactive oxygen species production induced by fluorescence microscopy: effect on mitosis and guidelines for non-invasive fluorescence microscopy. Plant J. 36: 280–290. Donner, H., Gao, L. and Mazza, G. (1997) Separation and characterization of simple and malonylated anthocyanins in red onions, Allium cepa L. Food Res. Int. 30: 637–643. Drabent, R., Pliska, B. and Olswewska, T. (1999) Fluorescent properties of plant anthocyanin pigments. I. Fluorescence of anthocyanins in Brassica oleracea L. extracts. J. Photochem. Photobiol. B. Biol. 50: 53–58. Dubrovsky, J.G., Guttenberger, M., Saralegui, A., Napsucialy-Mendivil, S., Voigt, B., Baluška, F., et al. (2006) Neutral red as a probe for confocal laser scanning microscope studies of plant roots. Ann. Bot. 97: 1127–1138. Emans, N., Zimmermann, S. and Fischer, R. (2002) Uptake of a fluorescent marker in plant cells is sensitive to brefeldin A and wortmannin. Plant Cell 14: 71–86. Finer, J.J., Vain, P., Jones, M.W. and McMullen, M.D. (1992) Development of the particle inflow gun for DNA delivery to plant cells. Plant Cell Rep. 11: 323–328. Gould, K.S. (2004) Nature’s Swiss army knife: the diverse protective roles of anthocyanins in leaves. J. Biomed. Biotechnol. 2004: 314–320. Grotewold, E. (2006) The genetics and biochemistry of floral pigments. Annu. Rev. Plant Biol. 57: 61–80. Guilliermond, A. (1929) The recent development of our idea of the vacuome in plant cells. Amer. J. Bot. 16: 1–22. Gunning, B. E. S. (2007) Plant Cell Biology on DVD. 1838 Haseloff, J., Siemering, K.R., Prasher, D.C. and Hodge, S. (1997) Removal of a cryptic intron and subcellular localization of green fluorescent protein are required to mark transgenic Arabidopsis plants brightly. Proc. Natl. Acad. Sci. USA 94: 2122–2127. Hashimoto, K., Igarashi, H., Mano, S., Nishimura, N., Shimmen, T. and Yokota, E. (2005) Peroxisomal localization of a myosin XI isoform in Arabidopsis thaliana. Plant Cell Physiol. 45: 782–789. Hecht, K. (1912) Studien über den Vorgang der Plasmolyse. Beitrage zur Biologie der Pflanzen 11: 133–192. Hicks, G.R., Rojo, E., Hong, S., Carter, D.G. and Raikhel, N.V. (2004) Germinating pollen has tubular vacuoles, displays highly dynamic biogenesis, and requires VACUOLESS1 for proper function. Plant Physiol. 134: 1227–1239. Higaki, T., Kutsuna, N., Okubo, E., Sano, T. and Hasezawa, S. (2006) Actin microfilaments regulate vacuolar structures and dynamics: dual observation of actin microfilaments and vacuolar membrane in living tobacco BY-2 cells. Plant Cell Physiol. 47: 839–852. Hillmer, S., Quader, H., Robert-Nicoud, M. and Robinson, D.G. (1989) Lucifer Yellow uptake in cells and protoplasts of Daucus carota visualised by laser scanning microscopy. J. Exp. Bot. 40: 417–423. Iglesias, V.A. and Meins, F. (2000) Movement of plant viruses is delayed in a b-1,3-glucanase-deficient mutant showing a reduced plasmodesmatal size exclusion limit and enhanced callose deposition. Plant J. 21: 157–166. Irani, N.G. and Grotewold, E. (2005) Light-induced morphological alterations in anthocyanin-accumulating vacuoles of maize cells. BMC Plant Biol. 5: 7. Kost, B., Spielhofer, P. and Chua, N.-H. (1998) A GFP-mouse talin fusion protein labels plant actin filaments in vivo and visualizes the actin cytoskeleton in growing pollen tubes. Plant J. 16: 393–401. Kutsuna, N., Kumagai, F., Sato, M.H. and Hasezawa, S. (2003) Threedimensional reconstruction of tubular structure of vacuolar membrane throughout mitosis in living tobacco cells. Plant Cell Physiol. 44: 1045–1054. Lazzaro, M.D. and Thomson, W.W. (1996) The vacuolar-tubular continuum in living trichomes of chickpea (Cicer arietinum) provides a rapid means of solute delivery from base to tip. Protoplasma 193: 181–190. Li, J.-F. and Nebenführ, A. (2007) Organelle targeting of myosin XI is mediated by two globular tail subdomains with separate cargo binding sites. J. Biol. Chem. 282: 20593–20602. Lichtscheidl, I.K. and Weiss, D.G. (1988) Visualization of submicroscopic structures in the cytoplasm in Allium cepa inner epidermal cells by video-enhanced contrast light microscopy. Eur. J. Cell Biol. 46: 376–382. Liebe, S. and Quader, H. (1994) Myosin in onion (Allium cepa) bulb scale epidermal cells: involvement in dynamics of organelles and cytoplasmic reticulum. Physiol. Plant. 90: 114–124. Marc, J., Granger, C.L., Brincat, J., Fisher, D.D., Kao, T.-H., McGrubbin, A.G., et al. (1998) A GFP-MAP4 reporter gene for visualizing cortical microtubule rearrangements in living epidermal cells. Plant Cell 10: 1927–1939. Marty, F. (1978) Cytochemical studies on GERL, provacuole, and vacuoles in root meristematic cells of Euphorbia. Proc. Natl Acad. Sci. USA 75: 852–856. Marty, F. (1999) Plant vacuoles. Plant Cell 11: 587–599. Nebenführ, A., Gallagher, L.A., Dunahy, T.G., Frohlick, J.A., Mazurkiewicz, A.M., Meehl, J.B., et al. (1999) Stop-and-go movements of plant Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. Tubular vacuoles in red onion epidermal cells Golgi stacks are mediated by the acto-myosin system. Plant Physiol. 121: 1127–1141. Newell, J.M., Leigh, R.A. and Hall, J.L. (1998) Vacuole development in cultured evacuolated oat mesophyll protoplasts. J. Exp. Bot. 49: 817–827. Oda, Y., Hirata, A., Sano, T., Fujita, T., Hiwatashi, Y., Sato, Y., et al. (2009) Microtubules regulate dynamic organization of vacuoles in Physcomitrella patens. Plant Cell Physiol. 50: 855–868. Ovečka, M., Lang, I., Baluška, F., Ismail, A., Illeš, P. and Lichtscheidl, I. (2005) Endocytosis and vesicle trafficking during tip growth of root hairs. Protoplasma 226: 39–54. Palevitz, B.A. and O’Kane, D.J. (1981) Epifluorescence and video analysis of vacuole motility and development in stomatal cells of Allium. Science 214: 443–445. Palevitz, B.A., O’Kane, D.J., Kobres, R.E. and Raikhel, N.V. (1981) The vacuole system in stomatal cells of Allium. Vacuole movements and changes in morphology in differentiating cells as revealed by epifluorescence, video and electron microscopy. Protoplasma 109: 23–55. Parthasarathy, M.V., Perdue, T.D., Witztum, A. and Alvernaz, J. (1985) Actin network as a normal component of the cytoskeleton in many vascular plant cells. Amer. J. Bot. 72: 1318–1323. Poustka, F., Irani, N.G., Feller, A., Lu, Y., Pourcel, L., Frame, K., et al. (2007) A trafficking pathway for anthocyanins overlaps with the endoplasmic reticulum-to-vacuole protein-sorting route in Arabidopsis and contributes to the formation of vacuolar inclusions. Plant Physiol. 145: 1323–1335. Quader, H., Hofmann, A. and Schnepf, E. (1989) Reorganization of the endoplasmic reticulum in epidermal cells of onion bulb scales after cold stress: involvement of cytoskeletal elements. Planta 177: 273–280. Quader, H. and Schnepf, E. (1986) Endoplasmic reticulum and cytoplasmic streaming: fluorescence microscopical observations in adaxial epidermis cells of onion bulb scales. Protoplasma 131: 250–252. Rees, B., Shepherd, V.A. and Ashford, A.E. (1994) Presence of a motile tubular vacuole system in different phyla of fungi. Mycol. Res. 98: 985–992. Riesen, D. and Hanson, M.R. (2007) Association of six YFP-myosin XI-tail fusions with mobile plant cell organelles. BMC Plant Biol. 7: 6. Reisen, D., Marty, F. and Leborgne-Castel, N. (2005) New insights into the tonoplast architecture of plant vacuoles and vacuolar dynamics during osmotic stress. BMC Plant Biol. 5: 13. Saito, C., Ueda, T., Abe, H., Wada, Y., Kuroiwa, T., Hisada, A., et al. (2002) A complex and mobile structure forms a distinct subregion within the continuous vacuolar membrane in young cotyledons of Arabidopsis. Plant J. 29: 245–255. 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 thaliana fimbrin highlights new features of a dynamic actin cytoskeleton in live plant cells. Plant Physiol. 136: 3968–3978. Shimmen, T. and Yoshida, S. (1993) Analysis of temperature dependence of cytoplasmic streaming using tonoplast-free cells of Characeae. Protoplasma 176: 174–177. Silverstone, A.L., Ciampaglio, C.N. and Sun, T.-P. (1998) The Arabidopsis RGA gene encodes a transcriptional regulator repressing the gibberellin signal transduction pathway. Plant Cell 10: 155–169. Slimestad, R., Fossen, T. and Vågen, I.M. (2007) Onions: a source of unique dietary flavonoids. J. Agric. Food Chem. 55: 10067–10080. Swanson, S.J., Bethke, P.C. and Jones, R.L. (1998) Barley aleurone cells contain two types of vacuoles: characterization of lytic organelles by use of fluorescent probes. Plant Cell 10: 685–698. Takemoto, D., Jones, D.A. and Hardham, A.R. (2003) GFP-tagging of cell components reveals the dynamics of subcellular re-organization in response to infection of Arabidopsis by oomycete pathogens. Plant J. 33: 775–792. Tamura, K., Shimada, T., Ono, E., Tanaka, Y., Nagatani, A., Higashi, S., et al. (2003) Why green fluorescent fusion proteins have not been observed in the vacuoles of higher plants. Plant J. 35: 545–555. Tanaka, Y., Kutsuna, N., Kanazawa, Y., Kondo, N., Hasezawa, S. and Sano, T. (2007) Intra-vacuolar reserves of membranes during stomatal closure: the possible role of guard cell vacuoles estimated by 3-D reconstruction. Plant Cell Physiol. 48: 1159–1169. Timmers, A.C.J., Tirlapur, U.K. and Schel, J.H.N. (1995) Vacuolar accumulation of acridine orange and neutral red in zygotic and somatic embryos of carrot (Daucus carota L.). Protoplasma 188: 236–244. Url, W. (1964) Phasenoptische Untersuchungen an Innerepidermen der Zweibelschuppe von Allium cepa L. Protoplasma 58: 294–311. van Gestel, K., Köhler, R.H. and Verbelen, J.-P. (2002) Plant mitochondria move on F-actin, but their positioning in the cortical cytoplasm depends on both F-actin and microtubules. J. Exp. Bot. 53: 659–667. Verbelen, J.-P. and Tao, W. (1998) Mobile arrays of vacuole ripples are common in plant cells. Plant Cell Rep. 17: 917–920. von Arnim, A.G., Deng, X.-W. and Stacey, M.G. (1998) Cloning vectors for the expression of green fluorescent protein fusion proteins in transgenic plants. Gene 221: 35–43. (Received July 26, 2009; Accepted September 1, 2009) Plant Cell Physiol. 50(10): 1826–1839 (2009) doi:10.1093/pcp/pcp124 © The Author 2009. 1839