Download plant cell biology in the new millennium: new tools and new

American Journal of Botany 87(11): 1547–1560. 2000.
INVITED SPECIAL PAPER
PLANT
CELL BIOLOGY IN THE NEW MILLENNIUM: NEW
TOOLS AND NEW INSIGHTS1
ELISON B. BLANCAFLOR2
AND
SIMON GILROY3,4
Plant Biology Division, The Samuel Roberts Noble Foundation Inc., 2510 Sam Noble Parkway, Ardmore, Oklahoma 73401 USA;
and 3Biology Department, 208 Mueller Laboratory, The Pennsylvania State University, University Park, Pennsylvania 16802 USA
2
The highly regulated structural components of the plant cell form the basis of its function. It is becoming increasingly recognized
that cellular components are ordered into regulatory units ranging from the multienzyme complexes that allow metabolic channeling
during primary metabolism to the ‘‘transducon’’ complexes of signal transduction elements that allow for the highly efficient transfer
of information within the cell. Against this structural background the highly dynamic processes regulating cell function are played
out. Recent technological advances in three areas have driven our understanding of the complexities of the structural and functional
dynamics of the plant cell. First, microscope and digital camera technology has seen not only improvements in the resolution of the
optics and sensitivity of detectors, but also the development of novel microscopy applications such as confocal and multiphoton
microscopy. These technologies are allowing cell biologists to image the dynamics of living cells with unparalleled three-dimensional
resolution. The second advance has been in the availability of increasingly powerful and affordable computers. The computer control/
analysis required for many of the new microscopy techniques was simply unavailable until recently. Third, there have been dramatic
advances in the available probes to use with these new microscopy approaches. Thus the plant cell biologist now has available a vast
array of fluorescent probes that will report cell parameters as diverse as the pH of the cytosol, the oxygen level in a tissue, or the
dynamics of the cytoskeleton. The combination of these new approaches has led to an increasingly detailed picture of how plant cells
regulate their activities.
Key words: caged probes; cell biology (plants); confocal microscopy; green fluorescent protein; laser tweezers; light microscopy,
ratio imaging; signal transduction.
OVERVIEW
The past decade has seen an explosion in the development
of new techniques in cell biology, particularly in the field of
light microscopy and imaging. New microscopes are continuously being developed that have either improved optics or
the ability to image new features of the cell. High performance
cameras with improved detection capabilities are also becoming a common feature of many cell biology laboratories. Since
its conception by Marvin Minsky in the 1950s, confocal microscopy has now become a routine technique and indispensable tool not only for cell biological studies but also for molecular investigations (Lichtman, 1994). In addition, although
used by only a handful of research laboratories worldwide
since its recent introduction, applications of multiphoton fluorescence microscopy to biological questions are likely to grow
rapidly within the next few years. This technique is poised to
provide a very real alternative to confocal microscopy in revealing the three-dimensional dynamics of the plant cell
(Denk, Strickler, and Webb, 1990). Computers are also fast
evolving to meet the demanding requirements for image capture and analysis that have become essential to make full use
of these new instruments. For example, approaches such as
real-time computational image deconvolution (Scalettar et al.,
1996) were until recently only available to microscopsists with
access to a supercomputer. All these developments have translated into a repertoire of advanced microscopy and imaging
Manuscript received 3 February 2000; revision accepted 5 May 2000.
Funding for this work was through the Samuel Roberts Noble Foundation
Inc. (E.B.B.) and grants from the National Aeronautics and Space Administration and the National Science Foundation (S.G.). The authors thank Dr.
Richard S. Nelson and Dr. Sarah Swanson for comments on the manuscript.
4
Author for reprint requests (e-mail: [email protected]).
1
techniques that can be exploited to aid in further understanding
the inner workings of plant cells.
The power of modern light microscopy to view plant cellular structure relies almost as much on improving fluorescent
probes as it does on faster computers, sensitive photodetectors,
and advanced lasers. The green fluorescent protein (GFP), a
new cellular marker cloned from the jellyfish Aequoria victoria, has allowed investigators to monitor cytoskeletal and
organelle dynamics in living plant cells (Kohler, 1998; Kost,
Spielhofer, and Chua, 1998; Marc et al., 1998) as well as longdistance transport of macromolecules and viruses (e.g., Imlau,
Truernit, and Sauer, 1999; Oparka et al., 1999; reviewed by
Thompson and Schulz, 1999; Lazarowitz, 1999). Several spectral variants of this protein have also enabled researchers to
monitor the dynamics of two or more molecules in the cell
simultaneously (Ellenberg, Lippincott-Schwartz, and Presley,
1999; Palm and Wlodawer, 1999; Haseloff, 1999). These spectral variants of GFP have also led to the design of transgenic
intracellular ion sensors that take advantage of fluorescence
energy resonance transfer (FRET) between different spectral
forms of GFP (Miyawaki et al., 1997, 1999; Allen et al., 1999;
Gadella, van der Krogt, and Bisseling, 1999).
The combination of sophisticated instrumentation with new
fluorescent probes has transformed light microscopy into a
powerful analytical tool that has allowed researchers to gain
new insights on basic plant cellular structure and function. As
new approaches emerge and current technologies undergo refinement, we are constantly presented with a new suite of tools
to tackle previously intractable questions in plant cell biology.
Due to space limitations, this review will highlight just some
of the developments in optical microscopy and imaging technology outlined above and how they have revolutionized our
view of plant cell function. We will also suggest some of the
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directions these new tools could lead us as we take plant cell
biology into the new millennium.
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For more than three centuries, the light microscope has provided biologists
with a powerful visual and analytical tool to study cells and tissues. With the
introduction of the electron microscope in the 1930s, researchers were able
to take cell biology several steps further into understanding cell function by
resolving structures not previously visible with the light microscope. Although
electron microscopy continues to play an essential role in characterizing cellular function it can only yield static images of cells and tissues. Yet it has
always been clear to microscopists that the plant cell possesses an intricately
patterned, highly dynamic, three-dimensional structure, which is thought to
form the basis of cellular function. Conventional light microscopy in combination with other contrast-enhancing approaches such as differential interference contrast (DIC) microscopy and phase contrast have provided tools to
monitor the structural dynamics of the cell (for a full discussion of these
techniques, see Ruzin, 1999). These microscopy approaches have continued
to be instrumental in cataloging the structural basis of plant cellular processes
from Robert Hooke’s discovery in 1665 that cells formed a functional unit of
cork to modern high-resolution DIC video-enhanced microscopy that allows
individual plant microtubules to be imaged in solution (Moore et al., 1997).
However, cell biologists had to await significant technological advances to be
able to capture and, more recently, manipulate the dynamic, three-dimensional
nature of specific molecular components of the plant cell in vivo.
the surface of individual hyphae can then be captured at different planes
within the cell using the confocal microscope. Figure 1 shows that the confocal imaging approach not only produces blur-free images, but also captures
the three-dimensional architecture of the cell. Thus each plane in the colonized
cell has a unique pattern of hyphae. A projection of sequential optical sections
taken throughout the depth of the cell shows the complex pattern of hyphae
as they ramify throughout the single colonized cell (Fig. 2). Such an appreciation of the three-dimensional details and complexity that forms the basis
of cellular structure is now routinely possible in large part because of widespread availability of confocal microscopes.
Confocal microscopy has been applied to answer questions as varied as
determining cellulose microfibril orientation (Verbelen and Stickens, 1995),
in situ localization of RNA and DNA probes (Wymer et al., 1999), determining the spatial and temporal characteristics of cell wall pH during root growth
(Taylor, Slatter, and, Leopold, 1996; Bibikova et al., 1998) and for monitoring
fluorescent dye movement to study cell-to-cell communication in roots and
shoots (Zhu, Lucas, and Rost, 1998; Gisel et al., 1999). Three-dimensional
morphological examination of surface structures used to be the realm of scanning electron microscopy (SEM), but confocal microscopy is proving to be
an effective tool in these type of studies as well (Lemon and Posluszny, 1998;
Melville et al., 1998), with the important advantage of being applicable to
living cells. However, it is important to note that the confocal microscope has
only reached its place as an indispensable tool for the plant biologist because,
in parallel, researchers have been developing an ever increasing range of
fluorescent reagents to visualize defined cellular structures and processes (see
below).
Confocal microscopy—Confocal microscopy is proving to be one of the
most exciting advances in optical microscopy of the last century. Although
conventional wide-field epifluorescence microscopy has been a powerful tool
for locating specific molecular components of the cell, it suffers from the
problem of out-of-focus fluorescence interfering with the contrast and resolution of the final image (Webb, 1999). With the commercialization of confocal microscopes in 1988, biologists began seeing cells in three dimensions
and with much more clarity than previously possible. The impact that confocal
microscopy has had on cell biology is evident in the number of reviews and
books written on the subject (e.g., Pawley, 1995; Sheppard and Shotton, 1997;
Webb, 1999). The number of scientific papers employing confocal microscopy
in plant biology has also grown dramatically since it was first introduced and
is indicative of the importance of this technology to plant science research
(Hepler and Gunning, 1998; Wymer et al., 1999).
Confocal microscopes work by exciting fluorescence with a highly focused
beam of laser light. The laser illuminates the sample, and the emitted fluorescent light is collected by the microscope objective. Light emitted from the
focal plane of the microscope passes through a pinhole aperture positioned in
front of a photomultiplier detector. The photomultiplier then passes the detected light signal to a computer, which will use it to generate an image. The
critical feature of the confocal approach is that light emitted from points above
or below the plane of focus is blocked by the pinhole and so never reaches
the detector. Thus only an image of the fluorescence from the focal plane is
observed (i.e., the microscope has generated an ‘‘optical section’’). A single
two-dimensional view of the optical section is then obtained by scanning the
laser from point to point over the sample until the entire focal plane of the
sample is imaged (Lichtman, 1994). Because only the light at the focal plane
of the sample passes through the pinhole onto the detector, the resulting image
is free from out- of-focus fluorescence. By collecting a series of optical sections at different focal planes, a full three-dimensional image of the sample
can be reconstructed (Webb, 1999). As sectioning is performed using optics
rather than the physical sectioning of the sample, living cells can be analyzed.
A series of optical sections through a root cell of Medicago truncatula
colonized with a symbiotic fungus (Glomus versiforme) demonstrates the
quality of images that can be obtained using confocal microscopy (Figs. 1
and 2). A root was stained with wheat germ agglutinin (WGA) linked to the
fluorescent dye Texas-red. WGA selectively binds to the N-acetoglucosamine
acid residues on the hyphal surface. Fluorescence from the Texas red labeling
Multiphoton fluorescence microscopy—A new approach to live cell imaging that is likely to have as dramatic an impact as confocal microscopy is
multiphoton microscopy. This technique uses a laser to deliver a burst of lowenergy, long-wavelength photons in a short pulse. The photon flux density in
the pulses is high enough that some of the photons hit a target fluorochrome
simultaneously. When the target absorbs two photons at essentially the same
time, they produce a similar effect as a single short-wavelength photon with
twice the energy (i.e., half the wavelength). For example, two photons of
infrared radiation (e.g., 700 nm) when absorbed simultaneously can excite a
UV or blue light excitable dye (e.g., a dye normally excited by 350 nm light;
Xu et al., 1995; Sako et al., 1995). The probability that two photons will
arrive at the fluorochrome simultaneously drops off dramatically away from
the focal plane of the microscope. Thus fluorescence is only effectively excited at the focal plane, and an optical section analogous to a confocal optical
section is generated. Unlike the confocal approach, however, the optical section is produced by the excitation of the fluorochrome in a single focal plane.
In addition, the multiphoton microscope imposes much less demand on the
detector optics because, unlike the confocal microscope, the emitted light does
not have to be accurately focused through a pinhole to achieve the optical
sectioning effect. This helps improve the efficiency of detection and the ability
to image deep into tissues.
Multiphoton microscopy has the potential to circumvent some of the problems inherent in confocal imaging. The high-intensity, short-wavelength irradiation used to excite the samples in many confocal imaging situations can
be damaging when imaged over long periods. Also, the shorter wavelengths
of light tend to penetrate less far into a sample (Gilroy, 1997). Multiphoton
microscopy, on the other hand, limits excitation to the focal plane of the
sample, and, therefore, photobleaching of the fluorochrome outside the focal
plane is minimized. This feature, in addition to the fact that the longer wavelength photons used in multiphoton microscopy have less energy, cuts down
on phototoxicity to the sample (Denk, Piston, and Webb, 1995). Longer wavelength photons are also less prone to scattering, making it possible to image
deeper into the sample (Gilroy, 1997; Fricker and Oparka, 1999). Multiphoton
microscopy promises to generate confocal-like images while making longer
observation times possible and should provide an important complement to
the confocal microscope (Sako et al., 1995).
ADVANCED OPTICAL MICROSCOPY
INSTRUMENTATION AND TECHNIQUES
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Figs. 1–2. Confocal images of an inner cortical cell colonized by arbuscular-mycorrhizal fungi in a root of Medicago truncatula. 1. A series of optical
sections (a–f) taken at 1-mm intervals using a Bio-Rad MRC 1024 confocal microscope of a root section stained with Texas-Red conjugated to wheat germ
agglutinin, which binds to the surface of the hyphae. Single optical sections are free from out-of-focus fluorescence, a typical problem encountered with
conventional fluorescence microscopy. 2. Projection of 29 optical sections taken at 0. 2-mm intervals from the same cell depicted in Fig. 1. Confocal microscopy
allows the projection of several optical sections to reveal the intricate three-dimensional structure of a cell. Note the highly complex distribution of hypha in a
colonized cell (E. B. Blancaflor, L. Zhao, and M. J. Harrison, unpublished data). Scale bar 5 10 mm.
FLUORESCENT DYES
The new imaging technologies have provided unparalleled
opportunities to image the plant cell. However, the insights
they have revealed have resulted in large part because of the
explosion in the availability of optical probes that can be used
with these techniques. Chemists are continuously developing
new fluorescent labels to allow imaging of proteins, nucleic
acids, lipids, and just about any specific cell component. In
addition, targeted probes for organelles and even specific ions
are now commercially available (Haugland, 1999). Many of
these reagents have yet to be successfully applied to plant
cells, and we can expect some plant-specific problems with
the use of some probes. For example, although fluorescent
probes to image membrane potential are widely used in animal
cells (e.g., Montana, Farkas, and Loew, 1989; Haugland,
1999), plant biologists have had little success with equivalent
approaches. Similarly, the ubiquitous ‘‘ester-loading’’ method
employed to load fluorescent dyes into the animal cell cytoplasm has had very limited success in plants where dyes may
completely fail to penetrate the cell (Gilroy, 1997; Fricker et
al., 1999) or accumulate in the vacuole rather than the cytoplasm (Swanson and Jones, 1996). However, there is a vast
array of untried fluorescent probes available to the plant biologist, and their potential to reveal specific insights into how
the plant cell functions is enormous. This is especially true
when they are combined with image analysis approaches, such
as ratio imaging, which allows fluorescence images to yield
quantitative information about the dynamics of the cell.
Fluorescence ratio imaging—The fluorescence of many
fluorochromes is sensitive to their molecular environment. For
example, the fluorescence of perhaps the most widely used
fluorochrome, fluorescein, increases as the pH rises. This has
led to the design of a whole spectrum of dyes that are selective
indicators of specific cell parameters or molecules. Thus there
are dyes whose fluorescence reports parameters ranging from
lipid mobility to specific ion concentrations (e.g., Ca21, Mg21,
H1, K1, Na1, Cl-, Zn21; Haugland, 1999). These indicators
have driven much of the advances in our understanding of
cellular control as they provide an image of the dynamics of
a well-defined cell component. Their usage is perhaps most
prevalent in the study of ionic signals such as Ca21 and pH.
For example, Ca2 1-selective dyes such as Fluo-3 and Calcium
Green, whose fluorescence increases in response to Ca21 levels, have been instrumental in demonstrating a previously unsuspected role for Ca21 release from intracellular stores and
intracellular waves of Ca21 in pollen tube growth (reviewed
in Franklin-Tong, 1999). However, there have always been
limitations in interpreting fluorescence images from such dyes.
The signal from the dye may increase due to increased Ca21,
but also due to dye relocalization, differences in dye concentration, photobleaching, and differences in optics between regions of a tissue or even subregions of a cell. This limitation
was elegantly overcome by applying the ratio analysis approach (Tanasugarn et al., 1984; Tsien and Poenie, 1986). In
this approach a spectral shift in fluorescence is monitored rather than simply monitoring fluorescence intensity. A wavelength of fluorescence that increases with ion concentration is
compared with one that is invariant or decreases with ion level.
The ratio of these intensities is independent of dye concentration and many of the other optical artifacts that make the signal
from single-wavelength dyes hard to quantify accurately.
The ratio analysis approach has allowed reliable, quantitative, spatial, and temporal analysis of images from fluorescent
probes. Pollen tube growth and signaling in guard cells serve
to highlight the power of this technology and the biological
surprises it has revealed. Ratio imaging has shown a tip-focused gradient in Ca21 centered on the growing tip of the pollen tube (Fig. 3). The gradient is always associated with the
growing tip and altering the gradient redirects growth (Malho
and Trewavas, 1996; Bibikova, Zhigilei, and Gilroy, 1997).
When tip growth ceases, the gradient is lost. Such data led to
the idea that localized Ca21 influx across the plasma membrane
generated a tip-focused Ca21 gradient that promoted secretion
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Figs. 3–6. Optical probes used in modern light microscopy to visualize cellular dynamics. 3. Ratio imaging of cytoplasmic Ca21 levels in a pollen tube of
Trandescantia. The pollen tube was microinjected with Indo-1 and Ca21 levels determined from the emission ratio 400–430 nm/460–480 nm using a Zeiss
LSM 410 UV confocal microscope. Ca21 levels were pseudocolored according to the inset scale. Fluorescence ratio imaging reveals the highly localized calcium
gradient at the pollen tube tip. Scale bar 5 10 mm. 4. Transient expression of GFP in an epidermal cell of Nicotiana tabacum. The coding region of enhanced
GFP (eGFP) was linked to the cauliflower mosaic virus 35S promoter, biolistically bombarded, and the leaf imaged using a Bio-Rad 1024 confocal microscope.
Free GFP is distributed throughout the cell cytoplasm and can also be seen to enter the nucleus (n) but not the chloroplasts (c) (S. Yin, E. B. Blancaflor, and
N. L. Paiva, unpublished data). Scale bar 5 20 mm. 5. A leaf of Nicotiana benthamiana inoculated with tobacco mosaic virus that was genetically modified to
express GFP. The leaf shows green fluorescence associated with the veins when photographed under UV light. The advent of GFP technology has allowed the
visualization of the pathways by which viruses move in living plants (N. Cheng and R. S. Nelson, unpublished data). Bar 5 1 cm. 6. Reorientation of
Trandescantia pollen tube growth after local photoactivation of caged Ca21 ionophore (A23187). Uncaging of the ionophore was accomplished by UV irradiating
the region denoted by the white box using a 0.7-s scan of a Zeiss UV confocal microscope. Local activation of the ionophore allows Ca21 influx into the cell
at that point. The pollen tube reorients toward the new site of elevated calcium induced by the localized Ca21 influx. Scale bar 5 10 mm. Inset shows fluorescence
ratio imaging of Ca21 levels in Trandescantia pollen tubes during localized photoactivation of caged Ca21 ionophore (Bibikova et al., 1997). The pollen tube
was microinjected with calcium green/rhodamine and Ca21 levels pseudocolored according to the scale in Fig. 3. Note that Ca21 is elevated at the site of
ionophore uncaging. Scale bar 5 10 mm.
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and therefore drives growth. The link between the Ca21 gradient and growth was even more strongly made when it was
noted by several researchers that pollen tubes grow in pulses
and that the gradient also oscillated (reviewed in FranklinTong, 1999). However, a closer examination revealed that the
story is not so simple. The increase in the Ca21 gradient coincides with growth but comparing this to measurements made
using the self-referencing (vibrating) microelectrode showed
Ca21 influx into the tip lagged the elevation of the cytosolic
gradient by 10 s (Pierson et al., 1996; Holdaway-Clarke et al.,
1997; Messerli and Robinson, 1997). We are still trying to
understand how these observations fit into a model of the regulation of tip growth, but Ca21 incorporation into newly synthesized wall polymers, stretch-activated Ca21 channels, and
Ca21 release from internal stores all seem plausible features
that could be interacting to give these ion dynamics. Recent
imaging data suggest the Ca21 gradient is controlled in part
by the Rop class of monomeric GTPases (Li et al., 1999) and
possibly influenced by the microtubule cytoskeleton (Bibikova, Blancaflor, and Gilroy, 1999). Thus the ratio imaging data
have not only indicated that Ca21 is a critical player in the
growth machinery but has also shown us that we have only
just begun to characterize its action.
In guard cells, the subtlety of potential signaling systems is
dramatically revealed by the ratio analysis approach. Guard
cells respond to multiple environmental signals including red
and blue light, CO2, humidity, and the stress hormone ABA
(abscisic acid; Assmann and Shimazaki, 1999). This ability to
integrate numerous signals has led to intensive studies on signaling in these cells. Studies with inhibitors of Ca21 signaling
hinted at a Ca21-dependent system of signaling. However, only
after ratio imaging allowed the spatial and temporal dynamics
of the system to be seen and manipulated was the full extent
of the complex signaling network in this single cell revealed
(reviewed in Hetherington et al., 1998). These studies showed
the operation of Ca21-dependent and -independent signaling
systems in response to ABA that is influenced by the environmental history of the plant (Allan et al., 1994). Evidence has
also accumulated for information being encoded in the spatial
patterns, amplitude, and frequency of transient elevations in
Ca21 levels occurring within the guard cell cytoplasm (Gilroy,
Read, and Trewavas, 1990; Grabov and Blatt, 1998; Staxen et
al., 1999). For example, Staxen et al. (1999) presented evidence that the frequency of Ca21 transients might encode the
ambient ABA concentration. Recent work with calmodulindependent protein kinase II from mammalian cells shows a
single enzyme can decode Ca21 transients into specific frequency-related intermediate enzymatic activities (Dekoninck
and Schulman, 1998), providing a model of how plant cells
might use such frequency-related information. It is clear that
most cells respond to myriad signals with a remarkable ability
to integrate and respond appropriately. We can anticipate that
the guard cell story of a complex and dynamic regulatory network will be reprised in many other systems as more plant
cells receive equivalent, intensive analysis by cell physiologists.
Aequorin—No review of imaging and analysis of signaling
systems in plants would be complete without mentioning the
advances made in the use of aequorin as a luminescent Ca21
indicator. Aequorin, like GFP, is a protein originally from the
jellyfish A. victoria that has proven of immense use to cell
biologists. Aequorin is a luminescent protein consisting of an
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apoprotein (apoaequorin) and coelentrazine, its small cofactor.
Light emission from aequorin is triggered by Ca21 with the
rate of luminescence being proportional to the Ca21 concentration. The gene for apoaequorin has been cloned, and when
cells expressing this protein are treated with coelentrazine,
functional aequorin is reconstituted. The luminescence rate can
be calibrated to an absolute Ca21 concentration making it possible to use transgenically expressed aequorin as a Ca21 sensor.
Plants such as tobacco and Arabidopsis have been stably transformed with aequorin and their Ca21 dynamics monitored either by putting whole plants in a luminometer, or by visualizing light emission using a highly sensitive camera.
Aequorin has revealed some surprising insights into plant
signaling and regulation. It has been used to show that there
are proliferating waves of Ca21 and organ-specific responses
when challenged by a host of environmental signals such as
salinity, drought, cold, heat shock, blue light, elicitors, touch,
or anoxia (reviewed in Trewavas, 1999). With aequorin it was
also shown that there are oscillations in Ca21 associated with
the circadian rhythms of the plant (Johnson et al., 1995). Aequorin has been targeted to various organelles and has shown,
for example, that not only is chloroplast Ca21 regulated independently of the cytosol (Johnson et al., 1995), but also, that
despite the extensive pore system in the nuclear membrane,
nuclear Ca21 is also regulated independently of cytosolic levels
(Van der Luit et al., 1999).
Artificial coelentrazines of varying Ca21 affinities, aequorins
with various spectral properties, and even a ratioable aequorin
are available (Knight et al., 1993). However, imaging the rapid
dynamics of Ca21 at the cellular level using aequorin is very
difficult, and imaging with subcellular resolution has proven
impossible (unless the protein is specifically targeted to a subcellular site) due to the inherently low signal generated by this
luminescent protein. These applications are more suited to
analysis by fluorescent dyes and GFP-based sensors such as
the cameleon Ca21 probe (see below). However, aequorin has
a far superior dynamic range than these fluorescent sensors
and can therefore easily detect rapid Ca21 transients in whole
plants. Aequorin’s high signal-to-background also makes it usable in plant cells where autofluorescence makes fluorescent
sensors inapplicable. These advantages and disadvantages of
aequorin highlight that although there is a remarkable array of
probes and technologies available to the plant biologist to
monitor cell behavior, each is appropriate to answer a specific
kind of question and it is in their combination and complementary usage that the real power of these new approaches
lies.
GREEN FLUORESCENT PROTEINS: NOVEL INSIGHTS
ON CELLULAR STRUCTURE, DYNAMICS, AND
SIGNALING IN LIVING PLANT CELLS
The discovery of GFP and several of its spectral variants
from Aequorea victoria (Ellenberg, Lippincott-Schwartz, and
Presley, 1999; Palm and Wlodawer, 1999; Haseloff, 1999) and
more recently a red fluorescent protein from another anthozoan
species (Matz et al., 1999) has provided a unique complement
to the battery of imaging devices that have been developed
during the latter part of the 20th century. GFP is a 27-kDa
protein that fluoresces bright green when excited by UV or
blue light (Fig. 4). GFP fluorescence depends on an internal
chromophore and so, unlike other biological fluorochromes, it
does not need the addition of a cofactor to fluoresce (Cubbit,
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Wollenweber, and Heim, 1999; Prendegast, 1999). In addition,
mutated forms of GFP have been generated with altered spectral characteristics (e.g., Blue-FP, Cyan-FP, and Yellow-FP), as
well as altered stabilities and solubilities (Heim and Tsien,
1996). The gene for GFP can be fused to other target genes
and the resulting construct genetically engineered into a cell.
The behavior of the resulting GFP-fusion protein can then be
followed in vivo. The entire GFP protein appears necessary to
generate the fluorochrome (Cubbit, Wollenweber, and Heim,
1999) and so the entire 27-kDa GFP coding region must be
fused to the gene of interest. Remarkably, attaching the 27kDa GFP to many proteins does not alter their activity, although this cannot be taken for granted and must be rigorously
proven in vitro and in vivo before results from the GFP tagged
protein can be easily interpreted. However, GFP fusions have
been applied with great success to follow the dynamics of the
plant cytoskeleton, endomembrane trafficking, organelle dynamics, and macromolecular transport. GFP has also been
used to mark specific cell types and to follow cell signaling
events. These topics are discussed below.
The plant cytoskeleton—Through the use of highly specific
antibodies against cytoskeletal proteins, researchers were able
to infer from fixed, static images how the plant cytoskeleton
reorganizes during various developmental stages and in response to a variety of environmental stimuli (Lloyd, 1987; Cyr,
1994; Hush and Overall, 1996; Nick, 1998). A more dynamic
element was added to this snapshot view of the cell through
the use of fluorescent analog cytochemistry, whereby a fluorescently labeled protein (e.g., rhodamine-tubulin) is introduced into the cell by microinjection. The labeled protein
could then be incorporated into the pool of normal cellular
proteins and their behavior monitored by fluorescence or confocal microscopy. Using this technique, the highly dynamic
nature of both the cortical and spindle microtubule cytoskeleton of living plant cells were directly observed and quantified
(Wasteneys, Gunning, and Hepler, 1993; Hush et al., 1994;
Yuan et al., 1994; Wymer et al., 1997; Himmelspach et al.,
1999). Similar progress has been made with the actin cytoskeleton. By microinjecting small amounts of fluorescently labeled phalloidin, a low molecular weight fungal metabolite
that binds preferentially to F-actin, it was possible to observe
the actin cytoskeleton in living plant cells undergoing division
(Schmit and Lambert, 1990; Cleary, 1995), in pollen tubes
(Miller, Lancelle, and Hepler, 1996), and in Transdescantia
stamen hairs (Ren et al., 1997).
The procedures to load fluorescently labeled proteins into
cells, however, can be a technically demanding and slow process, especially for walled plant cells. Microinjection, being
an invasive technique, can also perturb cellular function. Furthermore, the technique suffers from limited observation times
and is only applicable to isolated or exposed single cells (Kost,
Mathur, and Chua, 1999). These limitations have been partially
overcome with the use of GFP. By fusing the microtubulebinding domain of the microtubule-associated protein 4
(MAP4) with GFP and transiently expressing the recombinant
protein in plant cells, Marc et al. (1998) were able to label
cortical microtubules in living epidermal cells. A time course
analysis of the labeled microtubules revealed intricate dynamics including localized reorientations, lengthening, shortening,
and translocation of microtubules.
Kost, Spielhofer, and Chua (1998) have taken a similar approach for actin by transiently expressing GFP fused to the
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actin binding domain of talin, a mammalian actin-binding protein (designated GFP-mTn). GFP-mTn constructs decorated
actin filaments in tobacco pollen tubes and cultured BY2 cells,
which allowed the in vivo observation of actin dynamics in
these cells. More recently, stable transformation of Arabidopsis with the GFP-mTn and GFP-MAP4 fusions was used to
monitor actin filament and microtubule dynamics during trichome development (Mathur et al., 1999; Mathur and Chua,
2000).
Although these are the only reports to date on visualizing
the cytoskeleton in living plant cells using GFP fusions, we
can anticipate that studies on the plant cytoskeleton will advance rapidly in the next few years because of this technology. Additional advances will come by applying multiphoton
microscopy to intact plants expressing the GFP-cytoskeleton
binding protein fusions to image cytoskeletal dynamics in
cells located deeper in an intact plant organ (Potter et al.,
1996; Schwille et al., 1999). Studies with fluorescent analog
cytochemistry already suggest that microtubules on different
faces of a cell may have vastly different dynamics and organization (Yuan et al., 1992), and studies on fixed material
have shown that the cytoskeleton in the innermost cells of
roots reorganizes rapidly when exposed to a variety of hormones and other stimuli (Shibaoka, 1994; Blancaflor and
Hasenstein, 1995a, b; Blancaflor, Jones, and Gilroy, 1998;
Nick, 1998; Hasenstein, Blancaflor, and Lee, 1999). Imaging
the cytoskeleton deep into living tissues using multiphoton
and GFP technology should reveal the dynamic nature of
these responses. Furthermore, the availability of several spectral variants of GFP will be of enormous value in allowing
multiple tagging experiments to probe the interaction between different components of the plant cytoskeleton in living cells. For example, studies of the dynamics of the interaction between microtubules and actin should be high on the
priority list of plant cytoskeletal researchers (Tominaga et al.,
1997; Collings et al., 1998).
Visualizing intracellular organelle dynamics and
endomembrane trafficking—The use of GFP has also led to
new insights on the dynamics and functions of various cellular
organelles and the endomembrane system of plants. The creation of GFP fusions that contain specific localization sequences results in the retention of GFP in specific organelles and
membrane systems and allows their direct observation in vivo
(Kohler, 1998). Although membrane-permeable fluorescent
dyes have been available to observe the dynamics of some
cellular components such as the endoplasmic reticulum (ER)
and mitochondria in living plant cells, questions as to their
specificity have been a major concern (Hepler and Gunning,
1998). Furthermore, rapid photobleaching and the possible
toxic effects of these fluorescent dyes have made long-term
observation of these organelles in living plant cells difficult
(Scott et al., 1999).
So far, the most common and readily available GFP-based
subcellular markers are constructs that have the ER peptide
retention signal, KDEL. By fusing the KDEL sequence to the
coding sequence of GFP and expressing this construct in
plants, the dynamic cortical network of membranous tubules
representing the ER could be visualized (Boevink et al., 1996,
1998). This new vital ER marker has already provided new
information on structure/function of the endomembrane system as well as vesicle trafficking in plants (Hawes, Brandizzi,
and Andreeva, 1999). For example, GFP fused to the ER re-
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tention signal has not only demonstrated the highly dynamic
nature of the plant endomembrane system, but has also revealed the existence of rapidly moving organelles (Haseloff
and Siemering, 1998). Although these organelles have been
suggested to be proplastids, an alternative explanation as to
the identity of these organelles has been proposed (Gunning,
1998). Using a GFP fusion targeted to the Golgi apparatus
(made by linking GFP to the transmembrane domain of a rat
sialyl transferase) in combination with a second construct consisting of GFP fused to the Arabidopsis H/KDEL homologue
of the yeast HDEL receptor, aERD2, the simultaneous visualization of the Golgi and ER was possible. This provided the
first evidence that the ER and Golgi in plants are closely associated (Boevink et al., 1998).
In addition to the ER and Golgi, GFP has also been targeted
to the mitochondria (Kohler et al., 1997b), vacuole (Sansebastiano et al., 1998), plastids (Kohler et al., 1997a; Tirlapur
et al., 1999), and the nucleus (Chytilova, Macas, and Galbraith, 1999). Among other findings, this work has allowed
the monitoring of the mobility of thin tubular projections originating from plastids (Kohler et al., 1997a; Tirlapur et al.,
1999) and the movement and shape changes of the nucleus
(Chytilova, Macas, and Galbraith, 1999). Furthermore, by generating a protein fusion between GFP and a spliceosomal protein, the dynamics of coiled bodies, nuclear organelles involved in RNA processing, have been observed in higher
plants for the first time (Boudonck, Dolan, and Shaw, 1999).
By creating GFP fusions to known cellular proteins or their
putative targeting peptides, it has also been possible to determine the dynamics and localization patterns of these proteins
in living plant cells. For example, GFP linked to phragmoplastin, a protein known to be associated with cell plate formation, has allowed the dynamics of early cell plate development to be observed in vivo (Gu and Verma, 1997). A
unique family of calmodulin-dependent Ca21-ATPases were localized to the ER and nuclear envelope of Arabidopsis (Hong
et al., 1999) and a calmodulin-binding transporter protein was
shown to be associated with the plasma membrane of barley
aleurone protoplasts (Schuurink et al., 1998). Various proteins
(e.g., CRY2, COP1, phyB) implicated in light signal transduction were linked to GFP and shown to translocate to the
nucleus (Kleiner et al., 1999; Stacey and von Arnim, 1999;
Yamaguchi et al., 1999; Stacey, Hicks, and von Arnim, 1999),
while an isocitrate dehydrogenase isoenzyme fused to GFP
was demonstrated to localize exclusively to the mitochondria
(Galvez et al., 1998). GFP fusions to transit peptides of
phage-type organellar RNA polymerases also allowed localization of this protein to the mitochondria and plastids. These
data provided additional support for the involvement of these
polymerases in the transcriptional machinery of plant mitochondria and plastids (Hedtke et al., 1999). Therefore, with
the availability of several spectral variants of GFP and the
ability to target GFP to almost any cellular organelle desired,
we predict that the direct interactions between these different
organelles and/or protein fusions will be an area of intense
research.
Macromolecular transport and virus movements in
plants—Virus movement in plants is a complex process that
is generally considered to involve both cell- to-cell movement
via the plasmodesmata (Reichel, Mas, and Beachy, 1999) and
systemic transport via the vascular system (Nelson and van
Bel, 1998). Despite being intensively studied, the pathways
CELL BIOLOGY IN NEW MILLENNIUM
1553
and mechanisms by which viruses moved in plants were poorly understood primarily because of an inability to observe the
infection process in live tissue over time (Nelson and van Bel,
1998; Santa Cruz, 1999). However, through the observation of
GFP fluorescence from chimeric viruses, researchers can effectively monitor the vein classes by which viruses move after
infection in near-real time conditions (Fig. 5; Roberts et al.,
1997; Cheng et al., 2000).
New insights as to the manner in which viruses move from
cell to cell are also beginning to come to light. Although the
association of viral components with some of the host cell
components has been reported previously, these associations
were demonstrated mainly through electron and light microscopy of fixed plant tissues (Nelson and van Bel, 1998). With
modified viruses expressing GFP fusions to viral proteins, specific localization of viral proteins to plasmodesmata (Itaya et
al., 1997; Oparka et al., 1997) and other subcellular components of the plant host have been elegantly demonstrated. For
example, by linking the tobacco mosaic virus movement protein (MP) to GFP, colocalization of MP-GFP to cellular actin
was observed and MP was demonstrated to bind actin in vitro
(McLean, Zupan, and Zambryski, 1995). MP-GFP has also
been shown to localize to fluorescent aggregates and filamentous structures in infected BY2 cells. Treatment of infected
cells with microtubule-depolymerizing compounds disrupted
the filamentous strands providing evidence for MP-microtubule association (Heinlein et al., 1995, 1998). These studies
all indicate that the local spread of viruses could be facilitated
by interactions between a virus and these components of the
host plant (Reichel, Mas, and Beachy, 1999).
The progress made from studies on plant virus movement
has had a significant impact on plant cell biology and plant
physiology. The ability of viruses to penetrate the physical
barriers presented by the plant has opened new ideas on the
fundamental mechanisms by which macromolecules are transported throughout the plant. These transport stories have revealed perhaps one of the major discoveries of the past decade,
that plant nucleic acid can be transported from cell to cell.
This was originally reported for the Knotted gene in meristems
(Lucas et al., 1995), but there are now an ever-expanding set
of results pointing to a remarkable informational macromolecular exchange between plant cells that are likely to coordinate
many aspects of plant development and function (Lucas and
Wolf, 1999; Oparka et al., 1999). The reader is referred to the
several excellent reviews published recently that have thoroughly addressed these issues (Lazarowitz, 1999; Lazarowitz
and Beachy, 1999; Santa Cruz, 1999; Thompson and Schulz,
1999).
Plant cellular signaling—Another exciting use of GFP fusion proteins is to allow the visualization and quantification of
protein–protein interactions. This technique is called fluorescence resonance energy transfer (FRET) and relies on the phenomenon whereby a donor fluorescent molecule, when excited
with the appropriate wavelength of light, transfers some of its
emission energy to an adjacent acceptor chromophore. Since
FRET is dependent on the proximity of the donor to the acceptor, measurement of the efficiency of FRET can be used to
estimate the distance between donor and acceptor fluorophores. Therefore if two different proteins or molecules are
tagged with either a donor or acceptor fluorophore, FRET can
determine whether these proteins are in close association (Gordon et al., 1998).
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The concept of FRET in combination with GFP technology
offers new methods for elucidating signal transduction pathways in plants. For example, spectral variants of GFP were
recently used to construct a novel intracellular Ca21 sensor
referred to as ‘‘cameleon.’’ This probe was designed so that
two spectral variants of GFP with overlapping emission and
excitation spectra were fused at opposite ends of calmodulin
(a Ca21-binding protein) and the M13 peptide, (the calmodulin-binding domain from myosin light chain kinase). The two
varieties of cameleon constructs are the yellow-cameleon,
which consists of cyan fluorescent protein (CFP) and the yellow fluorescent protein (YFP), and the blue cameleon, which
consists of blue fluorescent protein (BFP) and GFP (Miyawaki
et al., 1997).
Cameleons report Ca21 in the following manner. Binding of
Ca21 to the calmodulin within the cameleon induces a conformational change causing it to bind to the M13 peptide. The
cameleon molecule is folded in the process such that the donor
(e.g., CFP) and acceptor (e.g., YFP) are brought in to closer
proximity to each other, thereby increasing FRET efficiency.
The donor:acceptor fluorescence ratio can then be calibrated
to the cytoplasmic Ca21 concentration (Miyawaki et al., 1997,
1999). Although the BFP constructs suffer from low signal
strength in plants, plant-expressed CFP constructs are bright
(Blancaflor, Dowd, and Gilroy, unpublished data) making the
CFP-YFP cameleon the appropriate choice for use with plant
tissues.
Cameleons were first used to measure intracellular calcium
in mammalian cells (Miyawaki et al., 1997; Emmanouilidou
et al., 1999), but the yellow cameleon construct has now been
successfully engineered into plant cells (Allen et al., 1999;
Fricker and Oparka, 1999; Gadella, van der Krogt, and Bisseling, 1999). Allen et al. (1999) constitutively expressed this
yellow cameleon in Arabidopsis and used this to investigate
cytoplasmic Ca21 changes in guard cells. Extracellular Ca21 or
ABA application induced cytoplasmic Ca21 transients in guard
cells, which were similar to the changes observed using fluorescent dyes. These results indicate that GFP-based Ca21 indicators are suitable for plants and therefore will be of enormous value in understanding the complex role of Ca21 in plant
cell signal transduction.
In addition to Ca21, pH has also been shown to tightly regulate numerous cellular signaling events and physiological
processes in plant cells (Guern et al., 1991). Like previous
studies with Ca21, measurement of cytoplasmic pH has relied
on fluorescent ratiometric dyes (Bibikova et al., 1998; Fricker
et al., 1999; Scott and Allen, 1999). GFP-based pH indicators
have also been recently developed for monitoring cytoplasmic
and organelle pH in mammalian cells (Kneen et al., 1998;
Llopis et al., 1998). Various laboratories are attempting to engineer these GFP-based pH indicators into plants, and it will
be exciting to see whether these pH sensors function in living
plant cells.
The use of GFP-based ion sensors to study plant signal
transduction is still in its infancy, but is likely to generate
information not previously obtainable with the traditional fluorescent dyes. This is due to the fact that these GFP-based ion
sensors can be easily targeted to organelles (see discussion
above); therefore the subtle changes in ions that appear to be
part of the signaling cascades of some plant processes such as
gravitropism can now be determined (Legue et al., 1997; Rosen, Chen, and Masson, 1999; Scott and Allen, 1999). Furthermore, the cameleon indicators can now be imaged with
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video-rate scanning two-photon microscopy (Fan et al., 1999);
therefore it will be possible to quantify rapid changes in Ca21
deep in an intact plant organ.
GFP as a cell- and tissue-specific marker—In developmental biology, labeling of cells to follow cell fate has relied
mainly on the microinjection of dyes into specific cell types
or immunocytochemical detection of specific cell types after
tissue fixation (Brand, 1999). With GFP, it is now possible to
mark specific cells or tissues in living Arabidopsis plants and
follow their fates directly during development (Fricker and
Oparka, 1999; Haseloff, 1999). This was accomplished by using the targeted gene expression approach that uses the GAL4
yeast transcription activator. In this ‘‘enhancer-trap’’ strategy,
the GAL4 gene is randomly integrated into the Arabidopsis
genome, bringing it under the control of different genomic
enhancer sequences that can direct a wide range of expression
patterns in the plant. However, in order to obtain efficient expression, it was necessary to alter codon usage by employing
a derivative sequence (GAL4-VP19). With GAL4-VP19
linked to mGFP5, which codes for ER-targeted GFP, the generated expression patterns of GFP in Arabidopsis roots could
be readily visualized with a confocal microscope (Haseloff,
1999).
Using the enhancer trap approach, root-cap-specific promoters were recently identified and used to genetically ablate
(i.e., selectively kill through the localized expression of a cytotoxic protein) root caps of Arabidopsis to study the consequences that such ablations have on root development (Haseloff, 1999). With the availability of specifically marked cell
lines, researchers were also able to follow cellular fate and
infer positional information that contributes to a specific type
of patterning in roots and shoots (Berger et al., 1998a, b; Sabatini et al., 1999). Cell-specific marking using GFP has also
been useful in the identification of endodermal protoplasts for
patch-clamp experiments (Maathuis et al., 1998) and companion cells for single-cell detection of gene transcripts (Brandt
et al., 1999).
One of the more exciting prospects of the enhancer trap
approach is the ability to activate other genes of interest in the
GFP-marked cell lines. This can be accomplished by subcloning the desired gene behind a tandem array of five optimized
GAL4 binding sites (the Upstream Activation Sequences) and
transforming plants with this construct. In the absence of the
GAL4 transcriptional activator, however, the desired gene is
not expressed. By crossing this plant with a line with a known
pattern of GAL4 expression, known because of the GFP expression patterns, the silenced gene is activated only in the
cells where GAL4 is expressed (Haseloff, 1999). Using this
approach, it should be possible, for example, to express the
GFP-based cameleon or pH indicators in specific cells or tissues in the plant and monitor signaling-related ion fluxes in
these cells. Since the fluorescence signal will be coming exclusively from specific cells, the problem of interfering signals
from other tissues is eliminated. Furthermore, it should also
be possible to express particular GFP-protein fusions such as
the GFP:MAP4 or GFP:talin fusions described above in specific cells and follow the dynamics of the cytoskeleton as these
cells progress developmentally.
OPTICAL TECHNIQUES FOR MICROMANIPULATION
OF PLANT CELLS
In addition to the improved ability to visualize dynamic
structures and quantify various physiological processes in
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BLANCAFLOR
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CELL BIOLOGY IN NEW MILLENNIUM
1555
Fig. 7. Displacement of mineral-containing vesicles (statoliths) using optical tweezers. The arrow indicates the initial trapping of statoliths on one side of
the rhizoid. Similar to the graviresponse, rhizoids bend toward the side of statolith accumulation (E. B. Blancaflor, and S. Gilroy, unpublished data). Scale bar
5 20 mm.
cells, optical microscopy is now being used as a precise tool
to manipulate both the physical and chemical properties of the
cell. The fields of chemistry and physics have contributed substantially to the development of these new optical approaches
and a representative from each field is discussed below.
Caged probe technology—Caged or photoactivatable
probes provide the opportunity to manipulate cell biochemistry
or signaling activities in situ using light. All caged probes
operate on a similar theme. A biologically active molecule is
derivatized by a caging group, which then sterically blocks the
activity of the molecule. The caging group is chosen carefully
to be photoactivatable. Thus, when the caging group absorbs
light (usually only UV light is energetic enough for these applications), it photolyzes, releasing the biologically active molecule. Some photolysis by-products are also released, and their
potential biological activity has to be carefully checked. However, the caged molecules used in plant cells to date have revealed no indication of biological activity or cytotoxicity of
these by products.
When the caged molecule is loaded into the cell (e.g., by
microinjection), then illumination using a standard fluorescence microscope is all it takes to release the molecule in the
cell. More sophisticated setups use a flash UV illuminator or
localized UV laser irradiation to produce highly controlled
temporal or spatial uncaging. In addition, the more UV light
that is used, the more probe is activated. Thus the cell biologist
can now control the spatial, temporal, and amplitude aspects
of a molecule’s activity in the living cell. As with fluorescent
probes, the range of caged probes that are available has grown
dramatically in the last few years. There are now caged hormones, amino acids, peptides, drugs (such as caged taxol),
nucleotides, ionophores, fluorochromes, and a range of cagedchelators and even caged-nitric oxide (Haugland, 1999). Most
are commercially available, although some of the more specialized reagents must be obtained from the laboratory where
they were synthesized.
Not surprisingly this technology has been applied to manipulate a wide range of control features of animal cells (Adams
and Tsien, 1993). However, though used in several plant studies, its potential has yet to be fully explored by plant cell
biologists. For example, caged Ca21, Ca21 chelators, and ionophores have been used to show a requirement for Ca21 in the
response to GA and ABA in aleurone cells (Gilroy, 1996),
phytochrome action (Shacklock, Read, and Trewavas, 1992;
Fallon, Shacklock, and Trewavas, 1993), the signaling pathway of stomatal closure (Gilroy, Read, and Trewavas, 1990)
and in the localization of growth in pollen tubes (Fig. 6; Malho
and Trewavas, 1996; Bibikova, Zhigilei, and Gilroy, 1997).
Caged inositol triphosphate (IP3) has revealed a role for an IP3
mobilizable internal Ca21 pool in the stomatal closure response
(Blatt, Thiel, and Trentham, 1990; Gilroy, Read, and Trewavas, 1990) and pollen tube growth (Franklin-Tong, 1999) and
caged ABA has been used to localize one site of ABA perception to an intracellular site in the guard cell (Allan et al.,
1994). Given the ability to quantitatively regulate the spatial
and temporal release of a caged molecule with subcellular precision, we can predict an increasing application of such approaches to probe the spatial and temporal requirements of
regulation in the plant cell.
Laser micromanipulation—The use of lasers is not only
limited to providing a coherent light source for microscopic
imaging of cells, but has also been used for the micromanipulation of whole cells and organelles. Laser micromanipulation
can be divided into two types. The first type, called laser tweezers or optical trapping, makes use of a continuous-wave, lowpower infrared laser beam. The laser beam is focused on an
object at the focal plane of the microscope, which results in
the refraction of the incident laser beam. The refraction of the
laser produces forces that pull the object toward the laser
beam, which can be used to hold the object in place. When
applied to cells, moving the laser beam can pull organelles or
whole cells from one place to another (Berns, 1998; Greulich
and Pilarczyk, 1998).
One example of the use of laser tweezers in biology has
been to address the problem of gravity sensing in the green
alga Chara. Chara is anchored to its substratum via singlecell, root-like structures called rhizoids. The tips of these rhizoids contain numerous vesicles that are filled with heavy mineral deposits (Fig. 7). Upon tilting the rhizoids by 908, these
vesicles sediment rapidly to the new lower flank of the cell.
Curvature commences shortly after sedimentation, and it is
believed that the sedimentation of these vesicles is responsible
for the initial perception of gravity leading to the redirection
of rhizoid growth (Sievers, Buchen and Hodick, 1996). Laser
tweezers have been used to mimic the gravitational displacement of these vesicles and have been shown to alter the direction of rhizoid growth (Fig. 7; Leitz, Schnepf, and Greulich,
1995). By combining laser trapping experiments and imaging
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techniques such as fluorescence ratio analysis, it is now possible to link displacement of the mineral containing vesicles
to other cell regulatory elements (e.g., changes in Ca21 distribution) that have been proposed to drive growth reorientation
during rhizoid gravitropism (Sinclair and Trewavas, 1997).
Optical trapping techniques have also been applied to study
the physical properties of the actin cytoskeleton in living plant
cells. In this case, light from an argon-ion laser is focused onto
a transvacuolar strand and the force needed to displace strandassociated vesicles is measured to give an indication of the
viscoelastic properties of actin (Schindler, 1995). Using this
technique, it was demonstrated that various factors including
lipids, Ca21, pH, aluminum, and the plant hormones auxin and
cytokinin alter the tension of the actin cytoskeleton in soybean
suspension cells (Grabski et al., 1994; Grabski and Schindler,
1995, 1996). Furthermore, Ca21-regulated kinases and phosphatases have been identified as possible signaling molecules
that modulate the changes in the viscoelastic properties of the
actin cytoskeleton (Grabski et al., 1998). However, whether
the transvacuolar strands from which tension measurements
were taken actually represent the actin cytoskeleton remains
questionable. With the availability of GFP to directly visualize
the cytoskeleton in living plant cells (see above; Kost, Spielhofer, and Chua, 1998; Marc et al., 1998), it would be interesting to combine optical trapping techniques and fluorescence
imaging to probe further into the physical properties of the
cytoskeleton.
The second type of laser micromanipulation is called optical
scissors or laser ablation. This technique relies on the precise
delivery of a high-powered laser beam to a target cell or organelle without affecting the areas surrounding the target (Ponelies et al., 1994; Berns, 1998; Greulich and Pilarczyk, 1998).
With this technique, whole plant cells have been ablated to
precisely define the gravity-sensing cells in root caps of Arabidopsis (Blancaflor, Fasano, and Gilroy, 1998, 1999). Signals
that affect cell fate determination during root development
have also been studied with the use of laser ablation (van den
Berg et al., 1997; Berger, 1998; Sabatini et al., 1999).
Other applications of laser ablation include exposing the
plasma membrane of plant cells by ablating out a small region
of the cell wall to facilitate genetic transformation (Guo,
Liang, and Berns, 1995) or to allow easy access of micropipettes for patch-clamp experiments (Henriksen and Assmann,
1997; Bouget, Berger, and Brownlee, 1998). Laser ablation
was also used to cut a hole in the wall of a single pollen grain
to isolate and eventually amplify its genomic DNA (Matsunaga et al., 1999). It is also now possible to combine laser
tweezers and laser ablation to obtain an even more subtle manipulation of plant cells. Buer et al. (1998) demonstrated that
cells of Agrobacterium rhizogenes could be inserted into callus
cells of Gingko biloba. They accomplished this by ablating a
2–4 mm hole through the cell wall with a pulsed laser beam
and moving one or two bacterial cells through the hole by
optical tweezers. Using optimum osmotic conditions, they
achieved a 95% cell survival rate. Therefore this technique
promises to be an important tool to address questions in plant
biology ranging from plant–microbe interactions to cellular organelle dynamics (Buer et al., 1998). For example, it would
be interesting to insert dense particles into statolith-free Chara
rhizoids (Kiss, 1994) to probe further into the gravity-sensing
mechanisms of these cells (Sievers, Buchen, and Hodick,
1996).
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CONCLUSIONS AND PROSPECTS
The rapid advances in optical probes and imaging technology make these exciting times for plant cell biology. The three
lines of technological advances that have brought us to this
unprecedented view of the dynamic plant cell show no sign
of slowing. First we can predict a continuing impact of everincreasing computing power. Image analysis is inherently a
computer-based science and as computing power increases we
can predict that techniques to improve resolution and extract
even more complex information from raw fluorescence images, such as real-time image deconvolution, will become
commonplace. The second thread of advance will be new and
exciting imaging technologies. We have highlighted multiphoton microscopy as a novel imaging approach with enormous
promise, but this is but one of the many technologies, such as
fluorescence lifetime imaging and correlation imaging, on the
horizon. The third line of advance will be in fluorescent and
luminescent probes. GFP technology is clearly an important
step towards the ultimate goal of noninvasively monitoring the
dynamics of the components of the plant cell, but there will
be an increasing need for fluorescent probes to monitor not
only levels and distribution of regulators but also their activity.
For example, recently, calmodulin activity has been imaged
using novel fluorescent dyes (Hahn, Waggoner, and Taylor,
1990; Torok et al., 1998) and a GFP-based probe (Persechini
and Cronk, 1999). These successes suggest that developing
probes to image the activities of cellular regulators is an attainable goal.
The most significant advances will be made by combining
these three areas of technology. For example, fluorescence correlation microscopy uses powerful computational analysis of
fluorescence detected by very sensitive imaging sensors
(Haupts et al., 1998; Schwille et al., 1999). It has been used
to look at the changes in parameters such as molecular diffusional constants of GFP fusion proteins to image their interactions with other cell components (Brock et al., 1999; Trier
et al., 1999). Computation, imaging, and fluorescent probes
have come together to reveal the dynamics of the environment
around a single molecule and have provided a very powerful
tool to monitor the activities of the living cell. The application
of such technologies to the vast array of unanswered questions
in plant development, function, and regulation will be a major
goal for plant cell biology researchers in the new millennium.
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