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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]
Received 22 July 2013; Revised 14 November 2013; Accepted 26 November 2013
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]
1682 | Truernit
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
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
Phloem imaging | 1683
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
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
1684 | Truernit
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.
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
Phloem imaging | 1685
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
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
1686 | Truernit
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
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.
Phloem imaging | 1687
(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).
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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