Journal of Experimental Botany, Vol. 65, No. 7, pp. 1681–1688, 2014 doi:10.1093/jxb/ert445 Advance Access publication 13 January, 2014
Review paper
Phloem imaging Elisabeth Truernit* Department of Biology, ETH Zürich, Universitätsstrasse 2, 8092 Zürich, Switzerland * To whom correspondence should be addressed. E-mail: [email protected]
Abstract The phloem is the long-distance solute-conducting tissue of plants. The observation of phloem cells is particularly challenging for several reasons and many recent advances in microscopy are, therefore, especially beneficial for the study of phloem anatomy and physiology. This review will give an overview of the imaging techniques that have been used for studying different aspects of phloem biology. It will also highlight some new imaging techniques that have emerged in recent years that will certainly advance our knowledge about phloem function. Key words: Development, microscopic techniques, microscopy, phloem, plant, sugar transport.
Introduction: the phloem—still an enigmatic tissue The phloem consists of several different cell types of which the sieve element (SE) cells have received the most attention due to their solute-conducting function. SE cells are long narrow cells with perforated end walls—the sieve plates. In angiosperms, several SE cells together form continuous tubes, the sieve tubes, in which solutes are transported from cell to cell over long distances. Sieve tube members in differentiated tissue usually form a functional unit with companion cells (CCs). Due to their transport function, sieve tube members undergo dramatic changes during differentiation and as a consequence of this become genetically and metabolically dependent on the associated CC (Oparka and Turgeon, 1999). In 1837, the forest botanist Hartig discovered the SE using bright-field light microscopy. In a publication addressing the progress that had been made in phloem research until 1956, Cheadle wrote: ‘Truly the field of phloem anatomy is a wide open one. We need more investigators; only a few botanists […] seem interested in solving the problems presented by this extraordinary assemblage of cells. […] there are problems enough for scores of researchers […]’ (Cheadle, 1956). Since then, quite a few plant scientists, albeit not ‘scores of’, have investigated and tried to understand the structure of the phloem in relation to its function as a long-distance
transporting tissue. The advent of electron microscopy (EM) has greatly increased our knowledge of phloem cell structure. To date, most of our textbook wisdom about phloem anatomy is derived from EM studies, and from an extensive collection of these data assembled by Katherine Esau (Esau, 1969). While we have a better idea about phloem structure today, quite a few questions remain unanswered (Knoblauch and Peters, 2010; Knoblauch and Oparka, 2012). With the availability of new imaging techniques, there are exciting new possibilities to address these questions, and phloem anatomy has been revisited in recent years with some interesting new approaches.
A short overview of microscopic techniques relevant to phloem imaging The story of phloem imaging is tightly linked to the availability of microscopic techniques. Prior to summarizing how different aspects of phloem biology have been visualized and studied, this review will briefly describe the technical aspects of some microscopic techniques that have been used for phloem imaging.
Abbreviations: 3D-SIM, three-dimensional structured illumination microscopy; CC, companion cell; CLEM, correlative light and electron microscopy; CLSM, confocal laser-scanning microscopy; EM, electron microscopy; FM, fluorescence microscopy; GFP, green fluorescent protein; NMR, nuclear magnetic resonance; PET, positron emission tomography; SBEM, serial block face scanning electron microscopy; SE, sieve element; SEM, scanning electron microscopy; TEM, transmission electron microscopy. © The Author 2014. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected]
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Received 22 July 2013; Revised 14 November 2013; Accepted 26 November 2013
1682 | Truernit block face scanning electron microscopy (SBEM), a scanning electron microscope images the block face of a resin-embedded specimen. After each scan, a thin section of the block is cut away by an integrated ultra-microtome and the next image is then taken. With this method, three-dimensional images of electron microscopic resolution can be obtained (Denk and Horstmann, 2004). In correlative microscopy, different microscopic techniques are used in combination on the same specimen. Combining the information from images taken with different methods is quite obviously superior to relying on just one system. For example, the rough localization of a fluorescently labelled protein in a CLSM image can be correlated with a certain subcellular structure in a TEM image taken of the same cell (Lidke and Lidke, 2012).
Challenges of phloem imaging There are several problems associated specifically with the observation of phloem cells. As part of the vasculature, phloem, together with xylem tissue, is located inside plant organs. Phloem cells are, therefore, always surrounded by several layers of other cells and thus cannot be looked at directly. To overcome this issue, the most common approach for the study of vascular cells has involved the preparation of tissue sections and their analysis with light and electron microscopy. However, SE cells are very long and narrow. They can reach a length of more than 1 mm while their radius can be 100 times shorter (Thompson and Holbrook, 2003). It therefore requires a lot of patience, luck, and effort to obtain a tissue section comprising a whole SE cell cut along its longitudinal axis. Similarly, many horizontal sections through phloem tissue are necessary to obtain only a few sections containing sieve plates. Moreover, unlike xylem cells, SE cells are living cells with relatively thin cell walls. This increases the risk of introducing dehydration artefacts during tissue treatments. In addition, SE cells form a long symplasmic continuum in which a considerable turgor pressure is generated. The sudden release of this pressure which is inevitable when cutting through the tissue, may lead to artefacts due to the displacement of cell contents. Thus, there was always considerable debate about whether the intracellular structures seen in EM pictures really reflected the in vivo state of sieve tube elements (Knoblauch and Oparka, 2012).
Catching the phloem alive To understand phloem function, imaging phloem cells in an undisturbed living state would be of great advantage. Sieve tube cells undergo considerable changes during their differentiation. A mature sieve tube cell contains only a very limited number of organelles (mitochondria, sieve tube plastids, and endoplasmic reticulum). Moreover, filamentous P-proteins (for phloem proteins) can be found in sieve tube cells (Esau, 1969). Due to contradictory results and the high possibility of introducing artefacts during tissue preparation, the location of these organelles and proteins within the sieve tube cells is not really clear. A major issue of debate is the state of
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The resolution limit of an optical microscope is defined as the shortest distance at which two point-like structures can be distinguished as separate entities. According to calculations by Abbe, the resolution limit of a microscope cannot be smaller than half the wavelength of the illuminating beam (Abbe, 1873). For conventional light microscopy, this means that the maximal achievable resolution is about 200 nm in the lateral (x–y) dimension. Since microscopes are usually designed to collect light from one side of the specimen only, the axial (z) resolution limit is even higher (>500 nm). The same basic principles apply to EM, but since electron beams have shorter wavelengths, the resolution limit of an electron microscope can be more than 1000 times lower (Erni et al., 2009). Therefore, from the 1950s until the present day, transmission electron microscopy (TEM) has been the medium of choice when subcellular detail needs to be studied. However, TEM requires elaborate tissue preparation and sectioning and only static images of dynamic processes can be obtained. Scanning electron microscopy (SEM) requires less effort in sample preparation, but is limited to the visualization of structures exposed on the surface of a specimen. The use of fluorescent proteins in plants began about 20 years ago (Chiu et al., 1996) and shifted the focus back to light microscopy—more specifically fluorescence microscopy (FM). Using fluorescent proteins of different colours to label different cell compartments, to monitor gene expression or to follow individual proteins in time and space is now a routine technique in many laboratories (see Berg and Beachy, 2008, for a review). In parallel, confocal laser-scanning microscopy (CLSM) has developed into a very important microscopic tool for cell biology, since it greatly improved the quality of images taken from fluorescent samples. CLSM is restricted by the resolution limit of light microscopy, but the combination of a focused laser beam for excitation and a pinhole in the detection path allows out-of-focus fluorescent light to be filtered out. Confocal imaging thus allows for image acquisition at defined z-positions and for the three-dimensional reconstruction of a specimen from stacks of images taken at different positions along its z-axis. More recently, several different routes have been taken to overcome the resolution limit of FM defined by Abbe. These novel developments are collectively referred to as ‘superresolution’ microscopy (for detailed reviews see Huang et al., 2009; Schermelleh et al., 2010). Only a few of these techniques have found their entry into plant research so far. Of these, three-dimensional structured illumination microscopy (3D-SIM) is one of the most promising, as it reduces the resolution limit in both the x–y dimension (to ≥100 nm) as well as in the z-dimension (to ≥200 nm) (Schermelleh et al., 2010). In this method, a series of structured light patterns are used to excite a fluorescent sample. The information from interference of the illumination structure with the fluorescent sample is then used to generate a high-resolution image. At present, image acquisition with 3D-SIM is too slow to allow for live imaging of moving samples, although some recent reports indicate that this may change in the future (Shao et al., 2011). Other recent developments have improved the amount of information that can be obtained from microscopy. In serial
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Fig. 1. Imaging phloem cells in an undisturbed state. (A) TEM of Arabidopsis sieve tubes in leaves. Tissue was preserved by plunge freezing of entire plants. (B) CLSM live imaging of V. faba sieve elements translocating the fluorescent dye CDCF (green). Tissue is counterstained with RH-160 (red). (C) CLSM live imaging of Arabidopsis root sieve elements translocating CF (red). AtSEOR1YFP filaments are shown in cyan. The arrow indicates the position of sieve plates. PP, phloem parenchyma. Scale bars: (A) 500 nm; (B, C) 20 µm. Images are reproduced with permission from (A, C) Froelich et al. (2012). Phloem ultrastructure and pressure flow: sieve-element-occlusion-related agglomerations do not affect translocation. The Plant Cell 23, 4428–4445; and (B) Knoblauch and van Bel (1998). Sieve tubes in action. The Plant Cell 10, 35–50; www.plantcell.org, Copyright American Society of Plant Biologists.
(Micro-ROCs) to observe Arabidopsis roots non-invasively (Froelich et al., 2012). In these chambers, roots grow on the surface of a cover slip, but are in direct contact with soil at the same time. This set-up was successfully used to get very detailed images of YFP fused to the sieve tube-specific filamentous proteins AtSEOR1 and 2 in actively translocating SEs (Fig. 1C) (Anstead et al., 2012; Froelich et al., 2012).
Imaging sieve plates Besides the fact that sieve pores may be obstructed under certain circumstances, the size and structure of sieve pores as well as the number of sieve pores per sieve plate will
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sieve pores in sieve plates: in microscopic images sieve pores often appeared to be occluded, which would interfere with long-distance transport (van Bel et al., 2002; Knoblauch and Oparka, 2012). During tissue preparation (mostly for EM), the usual methods for preserving cells in their natural state (rapid chemical fixation or freezing) are not as effective on phloem cells. Due to their location in the centre of the plant, it would take too long for the treatment to reach the phloem cells, even in the smallest of plants. For bigger plants, cutting before fixation would already introduce artefacts. Instead, some investigators tried to reduce phloem pressure by limiting solute production (Evert et al., 1973) or by using wilting plants (Anderson and Cronshaw, 1970). Others have used phloem cells generated from callus tissue to avoid tissue sectioning, with the caveat that they were not really studying cells that played an active role in solute transport (Sjolund, 1996). Recently, a successful plunge freezing approach with subsequent freeze-substitution in fixative was used for the small plant Arabidopsis thaliana (Anstead et al., 2012). In this experiment, phloem cells appeared to be very well preserved and sieve pores were mostly unobstructed (Fig. 1A). However, this method has only worked for the SEs in source leaves so far, most likely because they have the highest sugar content. This probably prevents the formation of water crystals in these cells during rapid freezing, and the cells are thus naturally anti-freeze protected (Froelich et al., 2012). Apart from wide field microscopy, CLSM is still the only microscopic technique that allows live-cell imaging to date. Although it is possible non-invasively to take images of cells in deeper tissue layers with CLSM, the phloem is, nevertheless, too deep inside almost all plant organs to obtain highresolution images of its subcellular structure. The first CLSM images of living phloem cells have been made from conifer bark. Bark pieces were infiltrated with fluorescent dyes that stained either the endomembrane system or the cytoplasm. In addition, cell wall autofluorescence was used for imaging (Schulz, 1992). While this approach still required cutting the bark, Knoblauch and van Bel came up with a technique some years later that did not injure the sieve tube continuum at all. Leaving the phloem intact, they took off a few cell layers above the lower side of the main vein of Vicia faba leaves at two locations. They then used one of these ‘windows’ to apply different fluorescent dyes, while the other one was used for CLSM imaging (Knoblauch and van Bel, 1998). In these fascinating images, intact phloem displayed mostly open sieve plate pores while deliberately damaged tissue showed the formation of plugs at the sieve plates, similar to what had been seen in many EM pictures (Fig. 1B). This method of imaging intact SE cells was later also used by others (Furch et al., 2009). Also, a very similar technique is currently in use for a study on the subcellular structure of phloem cells. This study is taking advantage of phloem-specific promoters and fluorescent organelle marker lines in the model plant Arabidopsis (S Dinant, personal communication). Being very small and transparent, the Arabidopsis root is probably one of the very few plant organs where live CLSM imaging of phloem cells is possible without any tissue preparation. Froelich et al. developed special rhizosphere chambers
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Imaging phloem development The events leading to the development of mature phloem cells are consecutive cell divisions resulting in the final number of phloem cells, followed by the terminal differentiation of these cells into different phloem cell types. Just a few cells are actively engaged in phloem development at any given time during plant growth. These cells are located at the tip of growing roots, in young leaves or at the vascular cambium. Only a small number of genetic determinants of phloem
Fig. 2. Visualizing sieve plates. (A, B) CLSM images of mPS-PI stained Arabidopsis stem sections. (A) Sieve plate in a cross section. (B) 3D reconstruction of one sieve plate. (C, D) Images of tobacco sieve plates labelled with callose-specific antibody and fluorescent secondary antibody (red colour). (C) is an image taken with CLSM while (D) is taken with 3D-SIM. (E, F) SEM images of sieve plates from (E) P. vulgaris and (F) R. communis. Scale bars: (A–E) 5 µm; (F) 20 µm. Images are reproduced with permission from (A, B) Truernit et al. (2008). High-resolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. The Plant Cell 20, 1494–1503; (C, D) Fitzgibbon et al. (2010). Super-resolution imaging of plasmodesmata using three-dimensional structured illumination microscopy. Plant Physiology 153, 1453–1463; and (E, F) Mullendore et al. (2010). Sieve tube geometry in relation to phloem flow. The Plant Cell 22, 579–593, Copyright American Society of Plant Biologists.
development are known to date (see Lucas et al., 2013, for a recent review). This is, at least partly, due to difficulties identifying deviations from normal phloem development on a large scale which is directly related to limitations in microscopic imaging. Due to its indeterminate growth, small size, and transparency, the Arabidopsis root is the model of choice for many studies on phloem development. In this organ, the phloem is arranged in two parallel files with the youngest cells developing from vascular stem cells located at the root tip. Protophloem
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obviously influence solute transport. Therefore, good statistical data about sieve plate anatomy is crucial for understanding the physical properties of sieve tube flow. These data are currently scarce. Sieve pores develop from plasmodesmata connecting young, developing sieve element cells (Esau, 1969). They are lined with callose in many plant species, which makes them discernible with callose-specific stains, most commonly the fluorescent dye aniline blue (Thompson and Wolniak, 2008). Sieve pore geometry has been determined mainly with TEM. With some exceptions, sieve pores from most plant species are smaller than 0.5 µm (Esau, 1969; Thompson and Holbrook, 2003), which means that they are difficult to resolve with light microscopy. On the other hand, although TEM produces high-resolution images of sieve pores, it requires extremely thin tissue sections. Analysis of a series of consecutive sections would be necessary to reconstruct the structure of one individual sieve plate. Thus, it is too time-consuming to obtain good statistical data of pore parameters using TEM. Hence, despite their small size there have been several attempts to estimate sieve pore sizes using fluorescent stains and FM (Thompson and Wolniak, 2008) or CLSM (Truernit et al., 2008) (Fig. 2A, B). Since their size is close to the resolution limit of light microscopy, sieve pores are an especially good example of the improvements in image quality achieved with super-resolution microcopy. Recently, some clear and well-resolved images of pores in sieve plates have been obtained using 3D-SIM on tobacco petiole sections treated with an anti-callose antibody and a fluorescently labelled secondary antibody (Fig. 2C, D) (Fitzgibbon et al., 2010). Another very promising approach has used SEM to image sieve plate anatomy (Mullendore et al., 2010). In this work, a method for clearing the cytoplasmic constituents from cells was developed, so that the bare cell wall could be imaged (Fig. 2E, F). The advantage of this method is that it can be used with rather thick (1 mm) vertical tissue sections, and thus the likelihood that several complete sieve plates are present in a single sample is relatively high. A potential limitation, however, is that pore sizes may be slightly over-estimated because of the missing protoplast. Nevertheless, this method has the highest potential for obtaining large data sets of sieve plate anatomy to date. Knowing the degree of pore size variance during different developmental stages, in different organs or under different environmental stimuli will greatly enhance our understanding of phloem transport.
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Fig. 3. Visualizing Arabidopsis root phloem development with CLSM. (A) Aniline blue-stained embryonic root. The developing phloem file is indicated with arrowheads. (B) mPS-PI-stained root. The developing protophloem file is indicated with arrowheads. (C) 3D-reconstruction of an mPS-PI-stained embryonic root. (D) Mature protophloem-specific GFP marker expression in a living root counterstained with propidium iodide. Scale bars: 50 µm. Reproduced from (A) (Bauby et al., 2007. Protophloem differentiation in early Arabidopsis thaliana development. Plant and Cell Physiology 48, 97–109), by permission of Oxford University Press, and (B, C) reproduced from Truernit et al. (2008). Highresolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. The Plant Cell 20, 1494–1503), www.plantcell.org, Copyright American Society of Plant Biologists.
by outer tissue layers). In cases where live imaging is not essential for the investigation, a way of preserving fluorescence during tissue optimization for microscopy would be most useful. There have been some promising approaches leading in this direction. Thompson and Wolniak (2008) described the retention of mCitrine fluorescence located at the SE plasma membrane, after fixing and embedding the tissue in methacrylate while Bell et al. (2013) developed a method that successfully conserved fluorescence during tissue embedding. They took CLSM and 3D-SIM images of fluorescently-labelled subcellular structures in the phloem after tissue fixation. Subsequently they used correlative light and electron microscopy (CLEM) unambiguously to identify the fluorescently-labelled cellular compartments in TEM images.
Imaging phloem flow The most important criterion for obtaining good physiological data about phloem flow is that the plants are treated in the least invasive way possible. At the same time, a detectable
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SE cells differentiate at a distance from these stem cells and metaphloem SE cells and CCs differentiate further up in the differentiation zone of the root. Our knowledge about the cell divisions leading to mature phloem cell types comes from the analysis of sequential cross-sections of the Arabidopsis root with bright-field light microscopy (Mähönen et al., 2000; Bonke et al., 2003). Similarly, a careful investigation of tissue sections through Arabidopsis embryos and very young seedlings using bright-field light microscopy and TEM has provided us with insight into the differentiation pattern of the first vascular elements in this plant (Busse and Evert, 1999). It is, however, very tedious to create a three-dimensional model of vascular development using tissue sections and thus these methods are rather unsuitable for large-scale analysis. CLSM in combination with 3D reconstruction software eliminates the need for physical sectioning, provided that the tissues are transparent enough for the clear resolution of vascular cells in the centre. Tissue staining and clearing techniques have been assessed over the past years regarding their suitability for phloem cell imaging with CLSM. Bauby et al. (2007) used an aniline blue staining technique (Bougourd et al., 2000) to characterize the early events of phloem differentiation in Arabidopsis embryos and young seedlings (Fig. 3A). The aniline blue in this case stains components in the cytoplasm and, therefore, can only be used for young cells with small vacuoles. Another method using a pseudo-Schiff propidium iodide staining (mPS-PI) technique was successfully used to visualize phloem cells in several plant organs (Fig. 3B, C) (Truernit et al., 2008). The mPS-PI technique stains cell walls and, unlike aniline blue, can be applied to all tissue types. It also allows for the easy visualization of the three-dimensional arrangement of vascular cells. In the root, for example, phloem cell files can be followed along their longitudinal axis from the vascular initial to the differentiated phloem cell. Thus, compared with tissue sectioning, it is a straightforward method for the production of a large quantity of data describing phloem development in wild-type and mutant plants. It is, however, unsuitable for monitoring the intracellular events of phloem differentiation as cell contents are largely removed during the clearing process. In this case, SBEM would be the preferred technique since it combines the high resolution of EM with the possibility of automatically reconstructing a three-dimensional image of a (small) sample. A characterization of the subcellular processes during phloem cell differentiation using SBEM is currently underway (Y Helariutta, personal communication). Another issue in studying phloem development is the problem associated with the imaging of marker gene expression. Marker genes labelling different cell compartments, cell types or developmental stages of a certain cell type are indispensable tools for studying development and several specific markers are available for Arabidopsis phloem cells (see Fig. 3D for an example). These markers are mostly transcriptional or translational fusions to fluorescent proteins and require imaging with FM. Commonly, fluorescent proteins do not withstand fixation or tissue clearing, and thus microscopy needs to be performed on live tissue samples with the aforementioned problems associated with it (low image quality due to light scattering caused
1686 | Truernit physiological processes over a relatively large field of view, and can image entire small plants (Kiser et al., 2008). Nuclear magnetic resonance (NMR) flow imaging (also called MRI velocimetry) has also been applied to understand phloem dynamics (Köckenberger, 2001; Windt et al., 2006). NMR flow imaging has the advantage of being completely non-invasive, as water flow in the phloem can be detected directly. Its resolution limit is in the range of several microns (Köckenberger, 2001), thus it provides higher resolution than PET, but it is not possible to determine phloem flow in individual sieve tubes. Köckenberger et al. (1997) were the first to measure phloem flow in intact seedlings using NMR (Fig. 4D). Windt et al. (2006) developed an NMR imager especially suited for experiments with plants. It consisted of an electromagnet with an open structure, in which potted plants of up to 2 m in height can be placed with minimal physical constraints and in an upright position. With this, they were able to measure phloem flow in non-stressed plants over long periods of time. With the increased availability of FM, the use of membranepermeable fluorescent dyes as phloem flow tracers became popular. The most commonly used dyes are 5(6)carboxyfluorescein diacetate (CFDA) and 8-hydroxypyrene-1,3,6trisulphonic acid (HPTS-acetate). These dyes can pass the membrane in their acetate form which means that there is no need for injecting them directly into phloem cells and, consequently, interfering with phloem turgor pressure. Once inside the cell, they are being cleaved by esterases, leading to the generation of membrane impermeable fluorescent compounds
Fig. 4. Visualizing phloem flow. (A) Arrival of CF in intact Arabidopsis roots. Images were taken with CLSM every 6 min (time-course from left to right) and are false coloured (highest fluorescence intensity is red). (B) CLSM image on the right shows movement of soluble GFP (green) expressed under control of the CC specific AtSUC2 promoter in a root of a pAtSUC2::GFP Arabidopsis plant. Image on the left shows AtSUC2 promoter specific expression of GFP targeted to the membrane (i.e. not soluble) as a control. (C) Arabidopsis stem section of pAtSUC2::GFP plant imaged with wide field FM and showing strong GFP fluorescence in the phloem (Ph). (D) Image of a castor bean hypocotyl overlaid with a colour image of a flow-sensitive NMR experiment. False colours indicate bidirectional flow in the xylem and phloem tissues of the vascular bundles. Scale bars: (A–C) 200 µm; (D) 1 mm. Images are reproduced with permission from (A) Oparka et al. (1994). Real-time imaging of phloem unloading in the root tip of Arabidopsis. The Plant Journal 6, 759–766, Copyright © 2003, John Wiley and Sons; (B) Stadler et al. (2004). Expression of GFP-fusions in Arabidopsis companion cells reveals non-specific protein trafficking into sieve elements and identifies a novel post-phloem domain in roots. The Plant Journal 41, 319–331, Copyright © 2004, John Wiley and Sons; (C) Imlau et al. (1999). Cell-to-cell and long-distance trafficking of the green fluorescent protein in the phloem and symplastic unloading of the protein into sink tissues. The Plant Cell 11, 309–322, www.plantcell.org, Copyright American Society of Plant Biologists; and (D) Köckenberger et al. (1997). A non-invasive measurement of phloem and xylem water flow in castor bean seedlings by nuclear magnetic resonance microimaging. Planta 201, 53–63, Copyright Springer-Verlag.
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substance needs to be present in the phloem solute stream. This can either be introduced into the plant or needs to be present in the plant already. If phloem tracers are applied locally, the distribution of solutes originating from specific regions of the plant can be analysed. Since very high resolution is not really required, a couple of uncommon plant imaging techniques can be used for monitoring phloem flow. Pulse-labelling experiments with radioactive carbon isotopes directly display the distribution of photosynthetic products in the plant. Experiments using 14CO2 to analyse solute transport reach back to the 1950s (Vernon and Aronoff, 1952). To get an idea about the spatial distribution of the isotopes within the plant, the plants are usually radio-autographed, and thus a static image of phloem flow is obtained with this method. While 14C has a very long half-life (c. 5700 years) and emits beta particles, the decay of the short-lived (c. 20 min) 11 C isotope emits gamma radiation. Its short half-life makes 11 C more suitable for the analysis of short-term effects on phloem transport and its radiation quality makes it possible to detect the presence of 11C non-invasively (Minchin and Thorpe, 2003). For example, 11C tracer measurements have been used to study changes in carbon transport and partitioning after sink removal or following different stresses (reviewed in Minchin and Thorpe, 2003). Considerable advances have also been made in the field of gamma irradiation detection. Imaging techniques based on positron emission tomography (PET) can now achieve a spatial resolution of a few millimetres. They provide quantitative information about dynamic
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Conclusions and outlook We are currently experiencing a rapid increase in information about phloem anatomy and function due to the fast evolution of microscopic techniques. While this review is dedicated to phloem imaging, the reader is referred to another review in this issue, which specifically deals with newly gained insights on sieve tube structure, and the impact of these findings on our understanding of long-distance transport (Knoblauch et al., 2013, this issue). It is quite clear that there will not be a single ‘best way’ of imaging phloem cells. The technique of choice will always depend on the specific question asked, but also quite simply on the availability of equipment in laboratories. While different questions need to be answered with different microscopic techniques, it will become increasingly important to approach one specific question with different methods. In this respect, correlative microscopy which, in theory, can be used with any combination of microscopic techniques, will be a very important approach that will certainly see more applications in the future. In addition, the rapid advances in the development of super-resolution microscopes are likely to continue. A critical step forward would be the possibility to use super-resolution microscopy for live cell imaging of phloem cells.
Acknowledgements I thank David Seung and Mario Coiro for critical reading of the manuscript.
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(CF, HPTS) that are delivered to the phloem, most likely through the symplast. Distribution of CF within the plant phloem accurately reflected the distribution of 14C-labelled sucrose (Grignon et al., 1989). Depending on the tissue type, CF and HPTS (to a lesser extent) can also be sequestered by the vacuole which may influence their overall distribution pattern (Wright and Oparka, 1996). Despite this relatively minor problem, these fluorescent dyes have been used widely and successfully in combination with FM and CLSM to increase our understanding of solute partitioning in plants (Fig. 4A) (Oparka et al., 1994; Knoblauch and van Bel, 1998). A genetic tool not requiring physical plant manipulation is the expression of soluble green fluorescent protein (GFP) under the control of the CC-specific AtSUC2 promoter (Imlau et al., 1999). In transgenic plants with this construct, GFP expressed in the CCs moves with the solute stream into SEs and then accumulates in sink tissue. Despite the relatively large molecular size of GFP compared with solutes and fluorescent dyes, it seems to be able to move freely with the solute stream in transgenic Arabidopsis and tobacco plants and can be monitored with FM or CLSM (Fig. 4B, C) (Imlau et al., 1999; Wright, 2003). pAtSUC2::GFP plants were used to observe solute partitioning at the whole plant level and under different environmental conditions. For example, they were used to investigate symplasmic domains in Arabidopsis seeds and embryos (Stadler, 2005). Screens aimed at identifying mutants with a reduction of GFP accumulation in sink tissues were also performed with these plants in order to find mutants impaired in phloem transport or symplasmic connectivity. The result of one of these screens was the identification of a callose synthase involved in plasmodesmata-localized callose production (Vatén et al., 2011).
1688 | Truernit sieve-element-occlusion-related agglomerations do not affect translocation. The Plant Cell 23, 4428–4445.
Oparka KJ, Turgeon R. 1999. Sieve elements and companion cells: traffic control centers of the phloem. The Plant Cell 11, 739–750.
Furch ACU, van Bel AJE, Fricker MD, Felle HH, Fuchs M, Hafke JB. 2009. Sieve element Ca2+ channels as relay stations between remote stimuli and sieve tube occlusion in Vicia faba. The Plant Cell 21, 2118–2132.
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Grignon N, Touraine B, Durand M. 1989. 6(5) Carboxyfluorescein as a Tracer of Phloem Sap Translocation. American Journal of Botany 76, 871–877. Huang B, Bates M, Zhuang X. 2009. Super-resolution fluorescence microscopy. Annual Review of Biochemistry 78, 993–1016.
Kiser MR, Reid CD, Crowell AS, Phillips RP, Howell CR. 2008. Exploring the transport of plant metabolites using positron emitting radiotracers. Human Frontier Science Program Journal 2, 189–204. Knoblauch M, Froelich DR, Pickard WF, Peters WS. 2014. SEORious business: structural proteins in sieve tubes and their involvement in sieve element occlusion. Journal of Experimental Botany 65, 1879–1893. Knoblauch M, Oparka K. 2012. The structure of the phloem: still more questions than answers. The Plant Journal 70, 147–156. Knoblauch M, Peters WS. 2010. Münch, morphology, microfluidics: our structural problem with the phloem. Plant, Cell and Environment 33, 1439–1452. Knoblauch M, van Bel AJ. 1998. Sieve tubes in action. The Plant Cell 10, 35–50. Köckenberger W. 2001. Functional imaging of plants by magnetic resonance experiments. Trends in Plant Science 6, 286–292. Köckenberger W, Pope JM, Xia Y, Jeffrey KR, Komor E, Callaghan PT. 1997. A non-invasive measurement of phloem and xylem water flow in castor bean seedlings by nuclear magnetic resonance microimaging. Planta 201, 53–63. Lidke DS, Lidke KA. 2012. Advances in high-resolution imaging - techniques for three-dimensional imaging of cellular structures. Journal of Cell Science 125, 2571–2580. Lucas WJ, Groover A, Lichtenberger R, et al. 2013. The plant vascular system: evolution, development and functions. Journal of Integrative Plant Biology 55, 294–388. Mähönen AP, Bonke M, Kauppinen L, Riikonen M, Benfey PN, Helariutta Y. 2000. A novel two-component hybrid molecule regulates vascular morphogenesis of the Arabidopsis root. Genes and Development 14, 2938–2943. Minchin PEH, Thorpe MR. 2003. Using the short-lived isotope C-11 in mechanistic studies of photosynthate transport. Functional Plant Biology 30, 831–841. Mullendore DL, Windt CW, Van As H, Knoblauch M. 2010. Sieve tube geometry in relation to phloem flow. The Plant Cell 22, 579–593. Oparka KJ, Duckett CM, Prior D, Fisher DB. 1994. Real-time imaging of phloem unloading in the root tip of Arabidopsis. The Plant Journal 6, 759–766.
Shao L, Kner P, Rego EH, Gustafsson MGL. 2011. Superresolution 3D microscopy of live whole cells using structured illumination. Nature Methods 8, 1044–1046. Sjolund RD. 1996. Phloem in plant tissue cultures. Progress in Botany : Springer, 356–367. Stadler R. 2005. Cell-to-cell movement of green fluorescent protein reveals post-phloem transport in the outer integument and identifies symplastic domains in Arabidopsis seeds and embryos. Plant Physiology 139, 701–712. Stadler R, Wright KM, Lauterbach C, Amon G, Gahrtz M, Feuerstein A, Oparka KJ, Sauer N. 2004. Expression of GFPfusions in Arabidopsis companion cells reveals non-specific protein trafficking into sieve elements and identifies a novel post-phloem domain in roots. The Plant Journal 41, 319–331. Thompson MV, Holbrook NM. 2003. Scaling phloem transport: water potential equilibrium and osmoregulatory flow. Plant, Cell and Environment 26, 1561–1577. Thompson MV, Wolniak SM. 2008. A plasma membraneanchored fluorescent protein fusion illuminates sieve element plasma membranes in Arabidopsis and tobacco. Plant Physiology 146, 1599–1610. Truernit E, Bauby H, Dubreucq B, Grandjean O, Runions J, Barthelemy J, Palauqui J-C. 2008. High-resolution wholemount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. The Plant Cell 20, 1494–1503. van Bel AJE, Ehlers K, Knoblauch M. 2002. Sieve elements caught in the act. Trends in Plant Science 7, 126–132. Vatén A, Dettmer J, Wu S, et al. 2011. Callose biosynthesis regulates symplastic trafficking during root development. Developmental Cell 21, 1144–1155. Vernon LP, Aronoff S. 1952. Metabolism of soybean leaves. IV. Translocation from soybean leaves. Archives of Biochemistry and Biophysics 36, 383–398. Windt CW, Vergeldt FJ, De Jager PA, Van As H. 2006. MRI of long-distance water transport: a comparison of the phloem and xylem flow characteristics and dynamics in poplar, castor bean, tomato and tobacco. Plant, Cell and Environment 29, 1715–1729. Wright KM. 2003. Structural and functional vein maturation in developing tobacco leaves in relation to AtSUC2 promoter activity. Plant Physiology 131, 1555–1565. Wright KM, Oparka KJ. 1996. The fluorescent probe HPTS as a phloem-mobile, symplastic tracer: an evaluation using confocal laser scanning microscopy. Journal of Experimental Botany 47, 439–445.
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Review paper
Phloem imaging Elisabeth Truernit* Department of Biology, ETH Zürich, Universitätsstrasse 2, 8092 Zürich, Switzerland * To whom correspondence should be addressed. E-mail: [email protected]
Abstract The phloem is the long-distance solute-conducting tissue of plants. The observation of phloem cells is particularly challenging for several reasons and many recent advances in microscopy are, therefore, especially beneficial for the study of phloem anatomy and physiology. This review will give an overview of the imaging techniques that have been used for studying different aspects of phloem biology. It will also highlight some new imaging techniques that have emerged in recent years that will certainly advance our knowledge about phloem function. Key words: Development, microscopic techniques, microscopy, phloem, plant, sugar transport.
Introduction: the phloem—still an enigmatic tissue The phloem consists of several different cell types of which the sieve element (SE) cells have received the most attention due to their solute-conducting function. SE cells are long narrow cells with perforated end walls—the sieve plates. In angiosperms, several SE cells together form continuous tubes, the sieve tubes, in which solutes are transported from cell to cell over long distances. Sieve tube members in differentiated tissue usually form a functional unit with companion cells (CCs). Due to their transport function, sieve tube members undergo dramatic changes during differentiation and as a consequence of this become genetically and metabolically dependent on the associated CC (Oparka and Turgeon, 1999). In 1837, the forest botanist Hartig discovered the SE using bright-field light microscopy. In a publication addressing the progress that had been made in phloem research until 1956, Cheadle wrote: ‘Truly the field of phloem anatomy is a wide open one. We need more investigators; only a few botanists […] seem interested in solving the problems presented by this extraordinary assemblage of cells. […] there are problems enough for scores of researchers […]’ (Cheadle, 1956). Since then, quite a few plant scientists, albeit not ‘scores of’, have investigated and tried to understand the structure of the phloem in relation to its function as a long-distance
transporting tissue. The advent of electron microscopy (EM) has greatly increased our knowledge of phloem cell structure. To date, most of our textbook wisdom about phloem anatomy is derived from EM studies, and from an extensive collection of these data assembled by Katherine Esau (Esau, 1969). While we have a better idea about phloem structure today, quite a few questions remain unanswered (Knoblauch and Peters, 2010; Knoblauch and Oparka, 2012). With the availability of new imaging techniques, there are exciting new possibilities to address these questions, and phloem anatomy has been revisited in recent years with some interesting new approaches.
A short overview of microscopic techniques relevant to phloem imaging The story of phloem imaging is tightly linked to the availability of microscopic techniques. Prior to summarizing how different aspects of phloem biology have been visualized and studied, this review will briefly describe the technical aspects of some microscopic techniques that have been used for phloem imaging.
Abbreviations: 3D-SIM, three-dimensional structured illumination microscopy; CC, companion cell; CLEM, correlative light and electron microscopy; CLSM, confocal laser-scanning microscopy; EM, electron microscopy; FM, fluorescence microscopy; GFP, green fluorescent protein; NMR, nuclear magnetic resonance; PET, positron emission tomography; SBEM, serial block face scanning electron microscopy; SE, sieve element; SEM, scanning electron microscopy; TEM, transmission electron microscopy. © The Author 2014. Published by Oxford University Press on behalf of the Society for Experimental Biology. All rights reserved. For permissions, please email: [email protected]
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Received 22 July 2013; Revised 14 November 2013; Accepted 26 November 2013
1682 | Truernit block face scanning electron microscopy (SBEM), a scanning electron microscope images the block face of a resin-embedded specimen. After each scan, a thin section of the block is cut away by an integrated ultra-microtome and the next image is then taken. With this method, three-dimensional images of electron microscopic resolution can be obtained (Denk and Horstmann, 2004). In correlative microscopy, different microscopic techniques are used in combination on the same specimen. Combining the information from images taken with different methods is quite obviously superior to relying on just one system. For example, the rough localization of a fluorescently labelled protein in a CLSM image can be correlated with a certain subcellular structure in a TEM image taken of the same cell (Lidke and Lidke, 2012).
Challenges of phloem imaging There are several problems associated specifically with the observation of phloem cells. As part of the vasculature, phloem, together with xylem tissue, is located inside plant organs. Phloem cells are, therefore, always surrounded by several layers of other cells and thus cannot be looked at directly. To overcome this issue, the most common approach for the study of vascular cells has involved the preparation of tissue sections and their analysis with light and electron microscopy. However, SE cells are very long and narrow. They can reach a length of more than 1 mm while their radius can be 100 times shorter (Thompson and Holbrook, 2003). It therefore requires a lot of patience, luck, and effort to obtain a tissue section comprising a whole SE cell cut along its longitudinal axis. Similarly, many horizontal sections through phloem tissue are necessary to obtain only a few sections containing sieve plates. Moreover, unlike xylem cells, SE cells are living cells with relatively thin cell walls. This increases the risk of introducing dehydration artefacts during tissue treatments. In addition, SE cells form a long symplasmic continuum in which a considerable turgor pressure is generated. The sudden release of this pressure which is inevitable when cutting through the tissue, may lead to artefacts due to the displacement of cell contents. Thus, there was always considerable debate about whether the intracellular structures seen in EM pictures really reflected the in vivo state of sieve tube elements (Knoblauch and Oparka, 2012).
Catching the phloem alive To understand phloem function, imaging phloem cells in an undisturbed living state would be of great advantage. Sieve tube cells undergo considerable changes during their differentiation. A mature sieve tube cell contains only a very limited number of organelles (mitochondria, sieve tube plastids, and endoplasmic reticulum). Moreover, filamentous P-proteins (for phloem proteins) can be found in sieve tube cells (Esau, 1969). Due to contradictory results and the high possibility of introducing artefacts during tissue preparation, the location of these organelles and proteins within the sieve tube cells is not really clear. A major issue of debate is the state of
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The resolution limit of an optical microscope is defined as the shortest distance at which two point-like structures can be distinguished as separate entities. According to calculations by Abbe, the resolution limit of a microscope cannot be smaller than half the wavelength of the illuminating beam (Abbe, 1873). For conventional light microscopy, this means that the maximal achievable resolution is about 200 nm in the lateral (x–y) dimension. Since microscopes are usually designed to collect light from one side of the specimen only, the axial (z) resolution limit is even higher (>500 nm). The same basic principles apply to EM, but since electron beams have shorter wavelengths, the resolution limit of an electron microscope can be more than 1000 times lower (Erni et al., 2009). Therefore, from the 1950s until the present day, transmission electron microscopy (TEM) has been the medium of choice when subcellular detail needs to be studied. However, TEM requires elaborate tissue preparation and sectioning and only static images of dynamic processes can be obtained. Scanning electron microscopy (SEM) requires less effort in sample preparation, but is limited to the visualization of structures exposed on the surface of a specimen. The use of fluorescent proteins in plants began about 20 years ago (Chiu et al., 1996) and shifted the focus back to light microscopy—more specifically fluorescence microscopy (FM). Using fluorescent proteins of different colours to label different cell compartments, to monitor gene expression or to follow individual proteins in time and space is now a routine technique in many laboratories (see Berg and Beachy, 2008, for a review). In parallel, confocal laser-scanning microscopy (CLSM) has developed into a very important microscopic tool for cell biology, since it greatly improved the quality of images taken from fluorescent samples. CLSM is restricted by the resolution limit of light microscopy, but the combination of a focused laser beam for excitation and a pinhole in the detection path allows out-of-focus fluorescent light to be filtered out. Confocal imaging thus allows for image acquisition at defined z-positions and for the three-dimensional reconstruction of a specimen from stacks of images taken at different positions along its z-axis. More recently, several different routes have been taken to overcome the resolution limit of FM defined by Abbe. These novel developments are collectively referred to as ‘superresolution’ microscopy (for detailed reviews see Huang et al., 2009; Schermelleh et al., 2010). Only a few of these techniques have found their entry into plant research so far. Of these, three-dimensional structured illumination microscopy (3D-SIM) is one of the most promising, as it reduces the resolution limit in both the x–y dimension (to ≥100 nm) as well as in the z-dimension (to ≥200 nm) (Schermelleh et al., 2010). In this method, a series of structured light patterns are used to excite a fluorescent sample. The information from interference of the illumination structure with the fluorescent sample is then used to generate a high-resolution image. At present, image acquisition with 3D-SIM is too slow to allow for live imaging of moving samples, although some recent reports indicate that this may change in the future (Shao et al., 2011). Other recent developments have improved the amount of information that can be obtained from microscopy. In serial
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Fig. 1. Imaging phloem cells in an undisturbed state. (A) TEM of Arabidopsis sieve tubes in leaves. Tissue was preserved by plunge freezing of entire plants. (B) CLSM live imaging of V. faba sieve elements translocating the fluorescent dye CDCF (green). Tissue is counterstained with RH-160 (red). (C) CLSM live imaging of Arabidopsis root sieve elements translocating CF (red). AtSEOR1YFP filaments are shown in cyan. The arrow indicates the position of sieve plates. PP, phloem parenchyma. Scale bars: (A) 500 nm; (B, C) 20 µm. Images are reproduced with permission from (A, C) Froelich et al. (2012). Phloem ultrastructure and pressure flow: sieve-element-occlusion-related agglomerations do not affect translocation. The Plant Cell 23, 4428–4445; and (B) Knoblauch and van Bel (1998). Sieve tubes in action. The Plant Cell 10, 35–50; www.plantcell.org, Copyright American Society of Plant Biologists.
(Micro-ROCs) to observe Arabidopsis roots non-invasively (Froelich et al., 2012). In these chambers, roots grow on the surface of a cover slip, but are in direct contact with soil at the same time. This set-up was successfully used to get very detailed images of YFP fused to the sieve tube-specific filamentous proteins AtSEOR1 and 2 in actively translocating SEs (Fig. 1C) (Anstead et al., 2012; Froelich et al., 2012).
Imaging sieve plates Besides the fact that sieve pores may be obstructed under certain circumstances, the size and structure of sieve pores as well as the number of sieve pores per sieve plate will
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sieve pores in sieve plates: in microscopic images sieve pores often appeared to be occluded, which would interfere with long-distance transport (van Bel et al., 2002; Knoblauch and Oparka, 2012). During tissue preparation (mostly for EM), the usual methods for preserving cells in their natural state (rapid chemical fixation or freezing) are not as effective on phloem cells. Due to their location in the centre of the plant, it would take too long for the treatment to reach the phloem cells, even in the smallest of plants. For bigger plants, cutting before fixation would already introduce artefacts. Instead, some investigators tried to reduce phloem pressure by limiting solute production (Evert et al., 1973) or by using wilting plants (Anderson and Cronshaw, 1970). Others have used phloem cells generated from callus tissue to avoid tissue sectioning, with the caveat that they were not really studying cells that played an active role in solute transport (Sjolund, 1996). Recently, a successful plunge freezing approach with subsequent freeze-substitution in fixative was used for the small plant Arabidopsis thaliana (Anstead et al., 2012). In this experiment, phloem cells appeared to be very well preserved and sieve pores were mostly unobstructed (Fig. 1A). However, this method has only worked for the SEs in source leaves so far, most likely because they have the highest sugar content. This probably prevents the formation of water crystals in these cells during rapid freezing, and the cells are thus naturally anti-freeze protected (Froelich et al., 2012). Apart from wide field microscopy, CLSM is still the only microscopic technique that allows live-cell imaging to date. Although it is possible non-invasively to take images of cells in deeper tissue layers with CLSM, the phloem is, nevertheless, too deep inside almost all plant organs to obtain highresolution images of its subcellular structure. The first CLSM images of living phloem cells have been made from conifer bark. Bark pieces were infiltrated with fluorescent dyes that stained either the endomembrane system or the cytoplasm. In addition, cell wall autofluorescence was used for imaging (Schulz, 1992). While this approach still required cutting the bark, Knoblauch and van Bel came up with a technique some years later that did not injure the sieve tube continuum at all. Leaving the phloem intact, they took off a few cell layers above the lower side of the main vein of Vicia faba leaves at two locations. They then used one of these ‘windows’ to apply different fluorescent dyes, while the other one was used for CLSM imaging (Knoblauch and van Bel, 1998). In these fascinating images, intact phloem displayed mostly open sieve plate pores while deliberately damaged tissue showed the formation of plugs at the sieve plates, similar to what had been seen in many EM pictures (Fig. 1B). This method of imaging intact SE cells was later also used by others (Furch et al., 2009). Also, a very similar technique is currently in use for a study on the subcellular structure of phloem cells. This study is taking advantage of phloem-specific promoters and fluorescent organelle marker lines in the model plant Arabidopsis (S Dinant, personal communication). Being very small and transparent, the Arabidopsis root is probably one of the very few plant organs where live CLSM imaging of phloem cells is possible without any tissue preparation. Froelich et al. developed special rhizosphere chambers
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Imaging phloem development The events leading to the development of mature phloem cells are consecutive cell divisions resulting in the final number of phloem cells, followed by the terminal differentiation of these cells into different phloem cell types. Just a few cells are actively engaged in phloem development at any given time during plant growth. These cells are located at the tip of growing roots, in young leaves or at the vascular cambium. Only a small number of genetic determinants of phloem
Fig. 2. Visualizing sieve plates. (A, B) CLSM images of mPS-PI stained Arabidopsis stem sections. (A) Sieve plate in a cross section. (B) 3D reconstruction of one sieve plate. (C, D) Images of tobacco sieve plates labelled with callose-specific antibody and fluorescent secondary antibody (red colour). (C) is an image taken with CLSM while (D) is taken with 3D-SIM. (E, F) SEM images of sieve plates from (E) P. vulgaris and (F) R. communis. Scale bars: (A–E) 5 µm; (F) 20 µm. Images are reproduced with permission from (A, B) Truernit et al. (2008). High-resolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. The Plant Cell 20, 1494–1503; (C, D) Fitzgibbon et al. (2010). Super-resolution imaging of plasmodesmata using three-dimensional structured illumination microscopy. Plant Physiology 153, 1453–1463; and (E, F) Mullendore et al. (2010). Sieve tube geometry in relation to phloem flow. The Plant Cell 22, 579–593, Copyright American Society of Plant Biologists.
development are known to date (see Lucas et al., 2013, for a recent review). This is, at least partly, due to difficulties identifying deviations from normal phloem development on a large scale which is directly related to limitations in microscopic imaging. Due to its indeterminate growth, small size, and transparency, the Arabidopsis root is the model of choice for many studies on phloem development. In this organ, the phloem is arranged in two parallel files with the youngest cells developing from vascular stem cells located at the root tip. Protophloem
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obviously influence solute transport. Therefore, good statistical data about sieve plate anatomy is crucial for understanding the physical properties of sieve tube flow. These data are currently scarce. Sieve pores develop from plasmodesmata connecting young, developing sieve element cells (Esau, 1969). They are lined with callose in many plant species, which makes them discernible with callose-specific stains, most commonly the fluorescent dye aniline blue (Thompson and Wolniak, 2008). Sieve pore geometry has been determined mainly with TEM. With some exceptions, sieve pores from most plant species are smaller than 0.5 µm (Esau, 1969; Thompson and Holbrook, 2003), which means that they are difficult to resolve with light microscopy. On the other hand, although TEM produces high-resolution images of sieve pores, it requires extremely thin tissue sections. Analysis of a series of consecutive sections would be necessary to reconstruct the structure of one individual sieve plate. Thus, it is too time-consuming to obtain good statistical data of pore parameters using TEM. Hence, despite their small size there have been several attempts to estimate sieve pore sizes using fluorescent stains and FM (Thompson and Wolniak, 2008) or CLSM (Truernit et al., 2008) (Fig. 2A, B). Since their size is close to the resolution limit of light microscopy, sieve pores are an especially good example of the improvements in image quality achieved with super-resolution microcopy. Recently, some clear and well-resolved images of pores in sieve plates have been obtained using 3D-SIM on tobacco petiole sections treated with an anti-callose antibody and a fluorescently labelled secondary antibody (Fig. 2C, D) (Fitzgibbon et al., 2010). Another very promising approach has used SEM to image sieve plate anatomy (Mullendore et al., 2010). In this work, a method for clearing the cytoplasmic constituents from cells was developed, so that the bare cell wall could be imaged (Fig. 2E, F). The advantage of this method is that it can be used with rather thick (1 mm) vertical tissue sections, and thus the likelihood that several complete sieve plates are present in a single sample is relatively high. A potential limitation, however, is that pore sizes may be slightly over-estimated because of the missing protoplast. Nevertheless, this method has the highest potential for obtaining large data sets of sieve plate anatomy to date. Knowing the degree of pore size variance during different developmental stages, in different organs or under different environmental stimuli will greatly enhance our understanding of phloem transport.
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Fig. 3. Visualizing Arabidopsis root phloem development with CLSM. (A) Aniline blue-stained embryonic root. The developing phloem file is indicated with arrowheads. (B) mPS-PI-stained root. The developing protophloem file is indicated with arrowheads. (C) 3D-reconstruction of an mPS-PI-stained embryonic root. (D) Mature protophloem-specific GFP marker expression in a living root counterstained with propidium iodide. Scale bars: 50 µm. Reproduced from (A) (Bauby et al., 2007. Protophloem differentiation in early Arabidopsis thaliana development. Plant and Cell Physiology 48, 97–109), by permission of Oxford University Press, and (B, C) reproduced from Truernit et al. (2008). Highresolution whole-mount imaging of three-dimensional tissue organization and gene expression enables the study of phloem development and structure in Arabidopsis. The Plant Cell 20, 1494–1503), www.plantcell.org, Copyright American Society of Plant Biologists.
by outer tissue layers). In cases where live imaging is not essential for the investigation, a way of preserving fluorescence during tissue optimization for microscopy would be most useful. There have been some promising approaches leading in this direction. Thompson and Wolniak (2008) described the retention of mCitrine fluorescence located at the SE plasma membrane, after fixing and embedding the tissue in methacrylate while Bell et al. (2013) developed a method that successfully conserved fluorescence during tissue embedding. They took CLSM and 3D-SIM images of fluorescently-labelled subcellular structures in the phloem after tissue fixation. Subsequently they used correlative light and electron microscopy (CLEM) unambiguously to identify the fluorescently-labelled cellular compartments in TEM images.
Imaging phloem flow The most important criterion for obtaining good physiological data about phloem flow is that the plants are treated in the least invasive way possible. At the same time, a detectable
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SE cells differentiate at a distance from these stem cells and metaphloem SE cells and CCs differentiate further up in the differentiation zone of the root. Our knowledge about the cell divisions leading to mature phloem cell types comes from the analysis of sequential cross-sections of the Arabidopsis root with bright-field light microscopy (Mähönen et al., 2000; Bonke et al., 2003). Similarly, a careful investigation of tissue sections through Arabidopsis embryos and very young seedlings using bright-field light microscopy and TEM has provided us with insight into the differentiation pattern of the first vascular elements in this plant (Busse and Evert, 1999). It is, however, very tedious to create a three-dimensional model of vascular development using tissue sections and thus these methods are rather unsuitable for large-scale analysis. CLSM in combination with 3D reconstruction software eliminates the need for physical sectioning, provided that the tissues are transparent enough for the clear resolution of vascular cells in the centre. Tissue staining and clearing techniques have been assessed over the past years regarding their suitability for phloem cell imaging with CLSM. Bauby et al. (2007) used an aniline blue staining technique (Bougourd et al., 2000) to characterize the early events of phloem differentiation in Arabidopsis embryos and young seedlings (Fig. 3A). The aniline blue in this case stains components in the cytoplasm and, therefore, can only be used for young cells with small vacuoles. Another method using a pseudo-Schiff propidium iodide staining (mPS-PI) technique was successfully used to visualize phloem cells in several plant organs (Fig. 3B, C) (Truernit et al., 2008). The mPS-PI technique stains cell walls and, unlike aniline blue, can be applied to all tissue types. It also allows for the easy visualization of the three-dimensional arrangement of vascular cells. In the root, for example, phloem cell files can be followed along their longitudinal axis from the vascular initial to the differentiated phloem cell. Thus, compared with tissue sectioning, it is a straightforward method for the production of a large quantity of data describing phloem development in wild-type and mutant plants. It is, however, unsuitable for monitoring the intracellular events of phloem differentiation as cell contents are largely removed during the clearing process. In this case, SBEM would be the preferred technique since it combines the high resolution of EM with the possibility of automatically reconstructing a three-dimensional image of a (small) sample. A characterization of the subcellular processes during phloem cell differentiation using SBEM is currently underway (Y Helariutta, personal communication). Another issue in studying phloem development is the problem associated with the imaging of marker gene expression. Marker genes labelling different cell compartments, cell types or developmental stages of a certain cell type are indispensable tools for studying development and several specific markers are available for Arabidopsis phloem cells (see Fig. 3D for an example). These markers are mostly transcriptional or translational fusions to fluorescent proteins and require imaging with FM. Commonly, fluorescent proteins do not withstand fixation or tissue clearing, and thus microscopy needs to be performed on live tissue samples with the aforementioned problems associated with it (low image quality due to light scattering caused
1686 | Truernit physiological processes over a relatively large field of view, and can image entire small plants (Kiser et al., 2008). Nuclear magnetic resonance (NMR) flow imaging (also called MRI velocimetry) has also been applied to understand phloem dynamics (Köckenberger, 2001; Windt et al., 2006). NMR flow imaging has the advantage of being completely non-invasive, as water flow in the phloem can be detected directly. Its resolution limit is in the range of several microns (Köckenberger, 2001), thus it provides higher resolution than PET, but it is not possible to determine phloem flow in individual sieve tubes. Köckenberger et al. (1997) were the first to measure phloem flow in intact seedlings using NMR (Fig. 4D). Windt et al. (2006) developed an NMR imager especially suited for experiments with plants. It consisted of an electromagnet with an open structure, in which potted plants of up to 2 m in height can be placed with minimal physical constraints and in an upright position. With this, they were able to measure phloem flow in non-stressed plants over long periods of time. With the increased availability of FM, the use of membranepermeable fluorescent dyes as phloem flow tracers became popular. The most commonly used dyes are 5(6)carboxyfluorescein diacetate (CFDA) and 8-hydroxypyrene-1,3,6trisulphonic acid (HPTS-acetate). These dyes can pass the membrane in their acetate form which means that there is no need for injecting them directly into phloem cells and, consequently, interfering with phloem turgor pressure. Once inside the cell, they are being cleaved by esterases, leading to the generation of membrane impermeable fluorescent compounds
Fig. 4. Visualizing phloem flow. (A) Arrival of CF in intact Arabidopsis roots. Images were taken with CLSM every 6 min (time-course from left to right) and are false coloured (highest fluorescence intensity is red). (B) CLSM image on the right shows movement of soluble GFP (green) expressed under control of the CC specific AtSUC2 promoter in a root of a pAtSUC2::GFP Arabidopsis plant. Image on the left shows AtSUC2 promoter specific expression of GFP targeted to the membrane (i.e. not soluble) as a control. (C) Arabidopsis stem section of pAtSUC2::GFP plant imaged with wide field FM and showing strong GFP fluorescence in the phloem (Ph). (D) Image of a castor bean hypocotyl overlaid with a colour image of a flow-sensitive NMR experiment. False colours indicate bidirectional flow in the xylem and phloem tissues of the vascular bundles. Scale bars: (A–C) 200 µm; (D) 1 mm. Images are reproduced with permission from (A) Oparka et al. (1994). Real-time imaging of phloem unloading in the root tip of Arabidopsis. The Plant Journal 6, 759–766, Copyright © 2003, John Wiley and Sons; (B) Stadler et al. (2004). Expression of GFP-fusions in Arabidopsis companion cells reveals non-specific protein trafficking into sieve elements and identifies a novel post-phloem domain in roots. The Plant Journal 41, 319–331, Copyright © 2004, John Wiley and Sons; (C) Imlau et al. (1999). Cell-to-cell and long-distance trafficking of the green fluorescent protein in the phloem and symplastic unloading of the protein into sink tissues. The Plant Cell 11, 309–322, www.plantcell.org, Copyright American Society of Plant Biologists; and (D) Köckenberger et al. (1997). A non-invasive measurement of phloem and xylem water flow in castor bean seedlings by nuclear magnetic resonance microimaging. Planta 201, 53–63, Copyright Springer-Verlag.
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substance needs to be present in the phloem solute stream. This can either be introduced into the plant or needs to be present in the plant already. If phloem tracers are applied locally, the distribution of solutes originating from specific regions of the plant can be analysed. Since very high resolution is not really required, a couple of uncommon plant imaging techniques can be used for monitoring phloem flow. Pulse-labelling experiments with radioactive carbon isotopes directly display the distribution of photosynthetic products in the plant. Experiments using 14CO2 to analyse solute transport reach back to the 1950s (Vernon and Aronoff, 1952). To get an idea about the spatial distribution of the isotopes within the plant, the plants are usually radio-autographed, and thus a static image of phloem flow is obtained with this method. While 14C has a very long half-life (c. 5700 years) and emits beta particles, the decay of the short-lived (c. 20 min) 11 C isotope emits gamma radiation. Its short half-life makes 11 C more suitable for the analysis of short-term effects on phloem transport and its radiation quality makes it possible to detect the presence of 11C non-invasively (Minchin and Thorpe, 2003). For example, 11C tracer measurements have been used to study changes in carbon transport and partitioning after sink removal or following different stresses (reviewed in Minchin and Thorpe, 2003). Considerable advances have also been made in the field of gamma irradiation detection. Imaging techniques based on positron emission tomography (PET) can now achieve a spatial resolution of a few millimetres. They provide quantitative information about dynamic
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Conclusions and outlook We are currently experiencing a rapid increase in information about phloem anatomy and function due to the fast evolution of microscopic techniques. While this review is dedicated to phloem imaging, the reader is referred to another review in this issue, which specifically deals with newly gained insights on sieve tube structure, and the impact of these findings on our understanding of long-distance transport (Knoblauch et al., 2013, this issue). It is quite clear that there will not be a single ‘best way’ of imaging phloem cells. The technique of choice will always depend on the specific question asked, but also quite simply on the availability of equipment in laboratories. While different questions need to be answered with different microscopic techniques, it will become increasingly important to approach one specific question with different methods. In this respect, correlative microscopy which, in theory, can be used with any combination of microscopic techniques, will be a very important approach that will certainly see more applications in the future. In addition, the rapid advances in the development of super-resolution microscopes are likely to continue. A critical step forward would be the possibility to use super-resolution microscopy for live cell imaging of phloem cells.
Acknowledgements I thank David Seung and Mario Coiro for critical reading of the manuscript.
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