plant cell biology in the new millennium - usp · 1547 american journal of botany 87(11):...

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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

2Plant 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

The highly regulated structural components of the plant cell form the basis of its function. It is becoming increasingly recognizedthat cellular components are ordered into regulatory units ranging from the multienzyme complexes that allow metabolic channelingduring primary metabolism to the ‘‘transducon’’ complexes of signal transduction elements that allow for the highly efficient transferof information within the cell. Against this structural background the highly dynamic processes regulating cell function are playedout. Recent technological advances in three areas have driven our understanding of the complexities of the structural and functionaldynamics of the plant cell. First, microscope and digital camera technology has seen not only improvements in the resolution of theoptics and sensitivity of detectors, but also the development of novel microscopy applications such as confocal and multiphotonmicroscopy. These technologies are allowing cell biologists to image the dynamics of living cells with unparalleled three-dimensionalresolution. 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 dramaticadvances in the available probes to use with these new microscopy approaches. Thus the plant cell biologist now has available a vastarray of fluorescent probes that will report cell parameters as diverse as the pH of the cytosol, the oxygen level in a tissue, or thedynamics of the cytoskeleton. The combination of these new approaches has led to an increasingly detailed picture of how plant cellsregulate 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 developmentof new techniques in cell biology, particularly in the field oflight microscopy and imaging. New microscopes are contin-uously being developed that have either improved optics orthe ability to image new features of the cell. High performancecameras with improved detection capabilities are also becom-ing a common feature of many cell biology laboratories. Sinceits conception by Marvin Minsky in the 1950s, confocal mi-croscopy has now become a routine technique and indispens-able tool not only for cell biological studies but also for mo-lecular investigations (Lichtman, 1994). In addition, althoughused by only a handful of research laboratories worldwidesince its recent introduction, applications of multiphoton fluo-rescence microscopy to biological questions are likely to growrapidly within the next few years. This technique is poised toprovide a very real alternative to confocal microscopy in re-vealing the three-dimensional dynamics of the plant cell(Denk, Strickler, and Webb, 1990). Computers are also fastevolving to meet the demanding requirements for image cap-ture and analysis that have become essential to make full useof these new instruments. For example, approaches such asreal-time computational image deconvolution (Scalettar et al.,1996) were until recently only available to microscopsists withaccess to a supercomputer. All these developments have trans-lated into a repertoire of advanced microscopy and imaging

1 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 Adminis-tration 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]).

techniques that can be exploited to aid in further understandingthe inner workings of plant cells.

The power of modern light microscopy to view plant cel-lular structure relies almost as much on improving fluorescentprobes as it does on faster computers, sensitive photodetectors,and advanced lasers. The green fluorescent protein (GFP), anew cellular marker cloned from the jellyfish Aequoria vic-toria, has allowed investigators to monitor cytoskeletal andorganelle dynamics in living plant cells (Kohler, 1998; Kost,Spielhofer, and Chua, 1998; Marc et al., 1998) as well as long-distance transport of macromolecules and viruses (e.g., Imlau,Truernit, and Sauer, 1999; Oparka et al., 1999; reviewed byThompson and Schulz, 1999; Lazarowitz, 1999). Several spec-tral variants of this protein have also enabled researchers tomonitor the dynamics of two or more molecules in the cellsimultaneously (Ellenberg, Lippincott-Schwartz, and Presley,1999; Palm and Wlodawer, 1999; Haseloff, 1999). These spec-tral variants of GFP have also led to the design of transgenicintracellular ion sensors that take advantage of fluorescenceenergy resonance transfer (FRET) between different spectralforms of GFP (Miyawaki et al., 1997, 1999; Allen et al., 1999;Gadella, van der Krogt, and Bisseling, 1999).

The combination of sophisticated instrumentation with newfluorescent probes has transformed light microscopy into apowerful analytical tool that has allowed researchers to gainnew insights on basic plant cellular structure and function. Asnew approaches emerge and current technologies undergo re-finement, we are constantly presented with a new suite of toolsto tackle previously intractable questions in plant cell biology.Due to space limitations, this review will highlight just someof the developments in optical microscopy and imaging tech-nology outlined above and how they have revolutionized ourview of plant cell function. We will also suggest some of the

1548 [Vol. 87AMERICAN JOURNAL OF BOTANY

directions these new tools could lead us as we take plant cellbiology into the new millennium.

ADVANCED OPTICAL MICROSCOPYINSTRUMENTATION AND TECHNIQUES

For more than three centuries, the light microscope has provided biologistswith a powerful visual and analytical tool to study cells and tissues. With theintroduction of the electron microscope in the 1930s, researchers were ableto take cell biology several steps further into understanding cell function byresolving structures not previously visible with the light microscope. Althoughelectron microscopy continues to play an essential role in characterizing cel-lular function it can only yield static images of cells and tissues. Yet it hasalways been clear to microscopists that the plant cell possesses an intricatelypatterned, highly dynamic, three-dimensional structure, which is thought toform the basis of cellular function. Conventional light microscopy in com-bination with other contrast-enhancing approaches such as differential inter-ference contrast (DIC) microscopy and phase contrast have provided tools tomonitor the structural dynamics of the cell (for a full discussion of thesetechniques, see Ruzin, 1999). These microscopy approaches have continuedto be instrumental in cataloging the structural basis of plant cellular processesfrom Robert Hooke’s discovery in 1665 that cells formed a functional unit ofcork to modern high-resolution DIC video-enhanced microscopy that allowsindividual plant microtubules to be imaged in solution (Moore et al., 1997).However, cell biologists had to await significant technological advances to beable to capture and, more recently, manipulate the dynamic, three-dimensionalnature of specific molecular components of the plant cell in vivo.

Confocal microscopy—Confocal microscopy is proving to be one of themost exciting advances in optical microscopy of the last century. Althoughconventional wide-field epifluorescence microscopy has been a powerful toolfor locating specific molecular components of the cell, it suffers from theproblem of out-of-focus fluorescence interfering with the contrast and reso-lution of the final image (Webb, 1999). With the commercialization of con-focal microscopes in 1988, biologists began seeing cells in three dimensionsand with much more clarity than previously possible. The impact that confocalmicroscopy has had on cell biology is evident in the number of reviews andbooks written on the subject (e.g., Pawley, 1995; Sheppard and Shotton, 1997;Webb, 1999). The number of scientific papers employing confocal microscopyin plant biology has also grown dramatically since it was first introduced andis 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 focusedbeam of laser light. The laser illuminates the sample, and the emitted fluo-rescent light is collected by the microscope objective. Light emitted from thefocal plane of the microscope passes through a pinhole aperture positioned infront of a photomultiplier detector. The photomultiplier then passes the de-tected light signal to a computer, which will use it to generate an image. Thecritical feature of the confocal approach is that light emitted from points aboveor below the plane of focus is blocked by the pinhole and so never reachesthe detector. Thus only an image of the fluorescence from the focal plane isobserved (i.e., the microscope has generated an ‘‘optical section’’). A singletwo-dimensional view of the optical section is then obtained by scanning thelaser from point to point over the sample until the entire focal plane of thesample is imaged (Lichtman, 1994). Because only the light at the focal planeof the sample passes through the pinhole onto the detector, the resulting imageis free from out- of-focus fluorescence. By collecting a series of optical sec-tions at different focal planes, a full three-dimensional image of the samplecan be reconstructed (Webb, 1999). As sectioning is performed using opticsrather than the physical sectioning of the sample, living cells can be analyzed.

A series of optical sections through a root cell of Medicago truncatulacolonized with a symbiotic fungus (Glomus versiforme) demonstrates thequality of images that can be obtained using confocal microscopy (Figs. 1and 2). A root was stained with wheat germ agglutinin (WGA) linked to thefluorescent dye Texas-red. WGA selectively binds to the N-acetoglucosamineacid residues on the hyphal surface. Fluorescence from the Texas red labeling

the surface of individual hyphae can then be captured at different planeswithin the cell using the confocal microscope. Figure 1 shows that the con-focal imaging approach not only produces blur-free images, but also capturesthe three-dimensional architecture of the cell. Thus each plane in the colonizedcell has a unique pattern of hyphae. A projection of sequential optical sectionstaken throughout the depth of the cell shows the complex pattern of hyphaeas they ramify throughout the single colonized cell (Fig. 2). Such an appre-ciation of the three-dimensional details and complexity that forms the basisof cellular structure is now routinely possible in large part because of wide-spread availability of confocal microscopes.

Confocal microscopy has been applied to answer questions as varied asdetermining cellulose microfibril orientation (Verbelen and Stickens, 1995),in situ localization of RNA and DNA probes (Wymer et al., 1999), determin-ing the spatial and temporal characteristics of cell wall pH during root growth(Taylor, Slatter, and, Leopold, 1996; Bibikova et al., 1998) and for monitoringfluorescent dye movement to study cell-to-cell communication in roots andshoots (Zhu, Lucas, and Rost, 1998; Gisel et al., 1999). Three-dimensionalmorphological examination of surface structures used to be the realm of scan-ning electron microscopy (SEM), but confocal microscopy is proving to bean effective tool in these type of studies as well (Lemon and Posluszny, 1998;Melville et al., 1998), with the important advantage of being applicable toliving cells. However, it is important to note that the confocal microscope hasonly reached its place as an indispensable tool for the plant biologist because,in parallel, researchers have been developing an ever increasing range offluorescent reagents to visualize defined cellular structures and processes (seebelow).

Multiphoton fluorescence microscopy—A new approach to live cell im-aging that is likely to have as dramatic an impact as confocal microscopy ismultiphoton microscopy. This technique uses a laser to deliver a burst of low-energy, long-wavelength photons in a short pulse. The photon flux density inthe pulses is high enough that some of the photons hit a target fluorochromesimultaneously. When the target absorbs two photons at essentially the sametime, they produce a similar effect as a single short-wavelength photon withtwice the energy (i.e., half the wavelength). For example, two photons ofinfrared radiation (e.g., 700 nm) when absorbed simultaneously can excite aUV 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 willarrive at the fluorochrome simultaneously drops off dramatically away fromthe focal plane of the microscope. Thus fluorescence is only effectively ex-cited at the focal plane, and an optical section analogous to a confocal opticalsection is generated. Unlike the confocal approach, however, the optical sec-tion is produced by the excitation of the fluorochrome in a single focal plane.In addition, the multiphoton microscope imposes much less demand on thedetector optics because, unlike the confocal microscope, the emitted light doesnot have to be accurately focused through a pinhole to achieve the opticalsectioning effect. This helps improve the efficiency of detection and the abilityto image deep into tissues.

Multiphoton microscopy has the potential to circumvent some of the prob-lems inherent in confocal imaging. The high-intensity, short-wavelength ir-radiation used to excite the samples in many confocal imaging situations canbe damaging when imaged over long periods. Also, the shorter wavelengthsof light tend to penetrate less far into a sample (Gilroy, 1997). Multiphotonmicroscopy, on the other hand, limits excitation to the focal plane of thesample, and, therefore, photobleaching of the fluorochrome outside the focalplane is minimized. This feature, in addition to the fact that the longer wave-length photons used in multiphoton microscopy have less energy, cuts downon phototoxicity to the sample (Denk, Piston, and Webb, 1995). Longer wave-length photons are also less prone to scattering, making it possible to imagedeeper into the sample (Gilroy, 1997; Fricker and Oparka, 1999). Multiphotonmicroscopy promises to generate confocal-like images while making longerobservation times possible and should provide an important complement tothe confocal microscope (Sako et al., 1995).

November 2000] 1549BLANCAFLOR AND GILROY—PLANT CELL BIOLOGY IN NEW MILLENNIUM

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 opticalsections (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 germagglutinin, which binds to the surface of the hyphae. Single optical sections are free from out-of-focus fluorescence, a typical problem encountered withconventional fluorescence microscopy. 2. Projection of 29 optical sections taken at 0. 2-mm intervals from the same cell depicted in Fig. 1. Confocal microscopyallows the projection of several optical sections to reveal the intricate three-dimensional structure of a cell. Note the highly complex distribution of hypha in acolonized 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 unparalleledopportunities to image the plant cell. However, the insightsthey have revealed have resulted in large part because of theexplosion in the availability of optical probes that can be usedwith these techniques. Chemists are continuously developingnew fluorescent labels to allow imaging of proteins, nucleicacids, lipids, and just about any specific cell component. Inaddition, targeted probes for organelles and even specific ionsare now commercially available (Haugland, 1999). Many ofthese reagents have yet to be successfully applied to plantcells, and we can expect some plant-specific problems withthe use of some probes. For example, although fluorescentprobes to image membrane potential are widely used in animalcells (e.g., Montana, Farkas, and Loew, 1989; Haugland,1999), plant biologists have had little success with equivalentapproaches. Similarly, the ubiquitous ‘‘ester-loading’’ methodemployed to load fluorescent dyes into the animal cell cyto-plasm has had very limited success in plants where dyes maycompletely fail to penetrate the cell (Gilroy, 1997; Fricker etal., 1999) or accumulate in the vacuole rather than the cyto-plasm (Swanson and Jones, 1996). However, there is a vastarray of untried fluorescent probes available to the plant bi-ologist, and their potential to reveal specific insights into howthe plant cell functions is enormous. This is especially truewhen they are combined with image analysis approaches, suchas ratio imaging, which allows fluorescence images to yieldquantitative information about the dynamics of the cell.

Fluorescence ratio imaging—The fluorescence of manyfluorochromes is sensitive to their molecular environment. Forexample, the fluorescence of perhaps the most widely usedfluorochrome, fluorescein, increases as the pH rises. This hasled to the design of a whole spectrum of dyes that are selectiveindicators of specific cell parameters or molecules. Thus thereare dyes whose fluorescence reports parameters ranging fromlipid mobility to specific ion concentrations (e.g., Ca21, Mg21,

H1, K1, Na1, Cl-, Zn21; Haugland, 1999). These indicatorshave driven much of the advances in our understanding ofcellular control as they provide an image of the dynamics ofa well-defined cell component. Their usage is perhaps mostprevalent in the study of ionic signals such as Ca21 and pH.For example, Ca2 1-selective dyes such as Fluo-3 and CalciumGreen, whose fluorescence increases in response to Ca21 lev-els, have been instrumental in demonstrating a previously un-suspected role for Ca21 release from intracellular stores andintracellular waves of Ca21 in pollen tube growth (reviewedin Franklin-Tong, 1999). However, there have always beenlimitations 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 concen-tration, photobleaching, and differences in optics between re-gions of a tissue or even subregions of a cell. This limitationwas elegantly overcome by applying the ratio analysis ap-proach (Tanasugarn et al., 1984; Tsien and Poenie, 1986). Inthis approach a spectral shift in fluorescence is monitored rath-er than simply monitoring fluorescence intensity. A wave-length of fluorescence that increases with ion concentration iscompared with one that is invariant or decreases with ion level.The ratio of these intensities is independent of dye concentra-tion and many of the other optical artifacts that make the signalfrom single-wavelength dyes hard to quantify accurately.

The ratio analysis approach has allowed reliable, quantita-tive, spatial, and temporal analysis of images from fluorescentprobes. Pollen tube growth and signaling in guard cells serveto highlight the power of this technology and the biologicalsurprises it has revealed. Ratio imaging has shown a tip-fo-cused gradient in Ca21 centered on the growing tip of the pol-len tube (Fig. 3). The gradient is always associated with thegrowing tip and altering the gradient redirects growth (Malhoand Trewavas, 1996; Bibikova, Zhigilei, and Gilroy, 1997).When tip growth ceases, the gradient is lost. Such data led tothe idea that localized Ca21 influx across the plasma membranegenerated a tip-focused Ca21 gradient that promoted secretion


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 ofTrandescantia. The pollen tube was microinjected with Indo-1 and Ca21 levels determined from the emission ratio 400–430 nm/460–480 nm using a ZeissLSM 410 UV confocal microscope. Ca21 levels were pseudocolored according to the inset scale. Fluorescence ratio imaging reveals the highly localized calciumgradient 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 enhancedGFP (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, andN. L. Paiva, unpublished data). Scale bar 5 20 mm. 5. A leaf of Nicotiana benthamiana inoculated with tobacco mosaic virus that was genetically modified toexpress GFP. The leaf shows green fluorescence associated with the veins when photographed under UV light. The advent of GFP technology has allowed thevisualization of the pathways by which viruses move in living plants (N. Cheng and R. S. Nelson, unpublished data). Bar 5 1 cm. 6. Reorientation ofTrandescantia pollen tube growth after local photoactivation of caged Ca21 ionophore (A23187). Uncaging of the ionophore was accomplished by UV irradiatingthe 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 cellat 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 fluorescenceratio imaging of Ca21 levels in Trandescantia pollen tubes during localized photoactivation of caged Ca21 ionophore (Bibikova et al., 1997). The pollen tubewas microinjected with calcium green/rhodamine and Ca21 levels pseudocolored according to the scale in Fig. 3. Note that Ca21 is elevated at the site ofionophore uncaging. Scale bar 5 10 mm.


and therefore drives growth. The link between the Ca21 gra-dient and growth was even more strongly made when it wasnoted by several researchers that pollen tubes grow in pulsesand that the gradient also oscillated (reviewed in Franklin-Tong, 1999). However, a closer examination revealed that thestory is not so simple. The increase in the Ca21 gradient co-incides with growth but comparing this to measurements madeusing the self-referencing (vibrating) microelectrode showedCa21 influx into the tip lagged the elevation of the cytosolicgradient by 10 s (Pierson et al., 1996; Holdaway-Clarke et al.,1997; Messerli and Robinson, 1997). We are still trying tounderstand how these observations fit into a model of the reg-ulation of tip growth, but Ca21 incorporation into newly syn-thesized wall polymers, stretch-activated Ca21 channels, andCa21 release from internal stores all seem plausible featuresthat could be interacting to give these ion dynamics. Recentimaging data suggest the Ca21 gradient is controlled in partby the Rop class of monomeric GTPases (Li et al., 1999) andpossibly influenced by the microtubule cytoskeleton (Bibiko-va, Blancaflor, and Gilroy, 1999). Thus the ratio imaging datahave not only indicated that Ca21 is a critical player in thegrowth machinery but has also shown us that we have onlyjust begun to characterize its action.

In guard cells, the subtlety of potential signaling systems isdramatically revealed by the ratio analysis approach. Guardcells respond to multiple environmental signals including redand blue light, CO2, humidity, and the stress hormone ABA(abscisic acid; Assmann and Shimazaki, 1999). This ability tointegrate numerous signals has led to intensive studies on sig-naling in these cells. Studies with inhibitors of Ca21 signalinghinted at a Ca21-dependent system of signaling. However, onlyafter ratio imaging allowed the spatial and temporal dynamicsof the system to be seen and manipulated was the full extentof the complex signaling network in this single cell revealed(reviewed in Hetherington et al., 1998). These studies showedthe operation of Ca21-dependent and -independent signalingsystems in response to ABA that is influenced by the environ-mental history of the plant (Allan et al., 1994). Evidence hasalso accumulated for information being encoded in the spatialpatterns, amplitude, and frequency of transient elevations inCa21 levels occurring within the guard cell cytoplasm (Gilroy,Read, and Trewavas, 1990; Grabov and Blatt, 1998; Staxen etal., 1999). For example, Staxen et al. (1999) presented evi-dence that the frequency of Ca21 transients might encode theambient ABA concentration. Recent work with calmodulin-dependent protein kinase II from mammalian cells shows asingle enzyme can decode Ca21 transients into specific fre-quency-related intermediate enzymatic activities (Dekoninckand Schulman, 1998), providing a model of how plant cellsmight use such frequency-related information. It is clear thatmost cells respond to myriad signals with a remarkable abilityto integrate and respond appropriately. We can anticipate thatthe guard cell story of a complex and dynamic regulatory net-work will be reprised in many other systems as more plantcells receive equivalent, intensive analysis by cell physiolo-gists.

Aequorin—No review of imaging and analysis of signalingsystems in plants would be complete without mentioning theadvances made in the use of aequorin as a luminescent Ca21

indicator. Aequorin, like GFP, is a protein originally from thejellyfish A. victoria that has proven of immense use to cellbiologists. Aequorin is a luminescent protein consisting of an

apoprotein (apoaequorin) and coelentrazine, its small cofactor.Light emission from aequorin is triggered by Ca21 with therate of luminescence being proportional to the Ca21 concen-tration. The gene for apoaequorin has been cloned, and whencells expressing this protein are treated with coelentrazine,functional aequorin is reconstituted. The luminescence rate canbe calibrated to an absolute Ca21 concentration making it pos-sible to use transgenically expressed aequorin as a Ca21 sensor.Plants such as tobacco and Arabidopsis have been stably trans-formed with aequorin and their Ca21 dynamics monitored ei-ther by putting whole plants in a luminometer, or by visual-izing light emission using a highly sensitive camera.

Aequorin has revealed some surprising insights into plantsignaling and regulation. It has been used to show that thereare proliferating waves of Ca21 and organ-specific responseswhen challenged by a host of environmental signals such assalinity, drought, cold, heat shock, blue light, elicitors, touch,or anoxia (reviewed in Trewavas, 1999). With aequorin it wasalso shown that there are oscillations in Ca21 associated withthe circadian rhythms of the plant (Johnson et al., 1995). Ae-quorin has been targeted to various organelles and has shown,for example, that not only is chloroplast Ca21 regulated inde-pendently of the cytosol (Johnson et al., 1995), but also, thatdespite 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, aequorinswith various spectral properties, and even a ratioable aequorinare available (Knight et al., 1993). However, imaging the rapiddynamics of Ca21 at the cellular level using aequorin is verydifficult, and imaging with subcellular resolution has provenimpossible (unless the protein is specifically targeted to a sub-cellular site) due to the inherently low signal generated by thisluminescent protein. These applications are more suited toanalysis by fluorescent dyes and GFP-based sensors such asthe cameleon Ca21 probe (see below). However, aequorin hasa far superior dynamic range than these fluorescent sensorsand can therefore easily detect rapid Ca21 transients in wholeplants. Aequorin’s high signal-to-background also makes it us-able in plant cells where autofluorescence makes fluorescentsensors inapplicable. These advantages and disadvantages ofaequorin highlight that although there is a remarkable array ofprobes and technologies available to the plant biologist tomonitor cell behavior, each is appropriate to answer a specifickind of question and it is in their combination and comple-mentary usage that the real power of these new approacheslies.

GREEN FLUORESCENT PROTEINS: NOVEL INSIGHTSON CELLULAR STRUCTURE, DYNAMICS, AND

SIGNALING IN LIVING PLANT CELLS

The discovery of GFP and several of its spectral variantsfrom Aequorea victoria (Ellenberg, Lippincott-Schwartz, andPresley, 1999; Palm and Wlodawer, 1999; Haseloff, 1999) andmore recently a red fluorescent protein from another anthozoanspecies (Matz et al., 1999) has provided a unique complementto the battery of imaging devices that have been developedduring the latter part of the 20th century. GFP is a 27-kDaprotein that fluoresces bright green when excited by UV orblue light (Fig. 4). GFP fluorescence depends on an internalchromophore and so, unlike other biological fluorochromes, itdoes not need the addition of a cofactor to fluoresce (Cubbit,


Wollenweber, and Heim, 1999; Prendegast, 1999). In addition,mutated forms of GFP have been generated with altered spec-tral characteristics (e.g., Blue-FP, Cyan-FP, and Yellow-FP), aswell as altered stabilities and solubilities (Heim and Tsien,1996). The gene for GFP can be fused to other target genesand the resulting construct genetically engineered into a cell.The behavior of the resulting GFP-fusion protein can then befollowed in vivo. The entire GFP protein appears necessary togenerate the fluorochrome (Cubbit, Wollenweber, and Heim,1999) and so the entire 27-kDa GFP coding region must befused to the gene of interest. Remarkably, attaching the 27-kDa GFP to many proteins does not alter their activity, al-though this cannot be taken for granted and must be rigorouslyproven in vitro and in vivo before results from the GFP taggedprotein can be easily interpreted. However, GFP fusions havebeen applied with great success to follow the dynamics of theplant cytoskeleton, endomembrane trafficking, organelle dy-namics, and macromolecular transport. GFP has also beenused to mark specific cell types and to follow cell signalingevents. These topics are discussed below.

The plant cytoskeleton—Through the use of highly specificantibodies against cytoskeletal proteins, researchers were ableto infer from fixed, static images how the plant cytoskeletonreorganizes during various developmental stages and in re-sponse to a variety of environmental stimuli (Lloyd, 1987; Cyr,1994; Hush and Overall, 1996; Nick, 1998). A more dynamicelement was added to this snapshot view of the cell throughthe use of fluorescent analog cytochemistry, whereby a fluo-rescently labeled protein (e.g., rhodamine-tubulin) is intro-duced into the cell by microinjection. The labeled proteincould then be incorporated into the pool of normal cellularproteins and their behavior monitored by fluorescence or con-focal microscopy. Using this technique, the highly dynamicnature of both the cortical and spindle microtubule cytoskel-eton 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 cyto-skeleton. By microinjecting small amounts of fluorescently la-beled phalloidin, a low molecular weight fungal metabolitethat binds preferentially to F-actin, it was possible to observethe actin cytoskeleton in living plant cells undergoing division(Schmit and Lambert, 1990; Cleary, 1995), in pollen tubes(Miller, Lancelle, and Hepler, 1996), and in Transdescantiastamen hairs (Ren et al., 1997).

The procedures to load fluorescently labeled proteins intocells, however, can be a technically demanding and slow pro-cess, especially for walled plant cells. Microinjection, beingan invasive technique, can also perturb cellular function. Fur-thermore, the technique suffers from limited observation timesand is only applicable to isolated or exposed single cells (Kost,Mathur, and Chua, 1999). These limitations have been partiallyovercome with the use of GFP. By fusing the microtubule-binding domain of the microtubule-associated protein 4(MAP4) with GFP and transiently expressing the recombinantprotein in plant cells, Marc et al. (1998) were able to labelcortical microtubules in living epidermal cells. A time courseanalysis of the labeled microtubules revealed intricate dynam-ics including localized reorientations, lengthening, shortening,and translocation of microtubules.

Kost, Spielhofer, and Chua (1998) have taken a similar ap-proach for actin by transiently expressing GFP fused to the

actin binding domain of talin, a mammalian actin-binding pro-tein (designated GFP-mTn). GFP-mTn constructs decoratedactin filaments in tobacco pollen tubes and cultured BY2 cells,which allowed the in vivo observation of actin dynamics inthese cells. More recently, stable transformation of Arabidop-sis with the GFP-mTn and GFP-MAP4 fusions was used tomonitor actin filament and microtubule dynamics during tri-chome development (Mathur et al., 1999; Mathur and Chua,2000).

Although these are the only reports to date on visualizingthe cytoskeleton in living plant cells using GFP fusions, wecan anticipate that studies on the plant cytoskeleton will ad-vance rapidly in the next few years because of this technol-ogy. Additional advances will come by applying multiphotonmicroscopy to intact plants expressing the GFP-cytoskeletonbinding protein fusions to image cytoskeletal dynamics incells located deeper in an intact plant organ (Potter et al.,1996; Schwille et al., 1999). Studies with fluorescent analogcytochemistry already suggest that microtubules on differentfaces of a cell may have vastly different dynamics and or-ganization (Yuan et al., 1992), and studies on fixed materialhave shown that the cytoskeleton in the innermost cells ofroots reorganizes rapidly when exposed to a variety of hor-mones and other stimuli (Shibaoka, 1994; Blancaflor andHasenstein, 1995a, b; Blancaflor, Jones, and Gilroy, 1998;Nick, 1998; Hasenstein, Blancaflor, and Lee, 1999). Imagingthe cytoskeleton deep into living tissues using multiphotonand GFP technology should reveal the dynamic nature ofthese responses. Furthermore, the availability of several spec-tral variants of GFP will be of enormous value in allowingmultiple tagging experiments to probe the interaction be-tween different components of the plant cytoskeleton in liv-ing cells. For example, studies of the dynamics of the inter-action between microtubules and actin should be high on thepriority list of plant cytoskeletal researchers (Tominaga et al.,1997; Collings et al., 1998).

Visualizing intracellular organelle dynamics andendomembrane trafficking—The use of GFP has also led tonew insights on the dynamics and functions of various cellularorganelles and the endomembrane system of plants. The cre-ation of GFP fusions that contain specific localization sequenc-es results in the retention of GFP in specific organelles andmembrane systems and allows their direct observation in vivo(Kohler, 1998). Although membrane-permeable fluorescentdyes have been available to observe the dynamics of somecellular components such as the endoplasmic reticulum (ER)and mitochondria in living plant cells, questions as to theirspecificity have been a major concern (Hepler and Gunning,1998). Furthermore, rapid photobleaching and the possibletoxic effects of these fluorescent dyes have made long-termobservation of these organelles in living plant cells difficult(Scott et al., 1999).

So far, the most common and readily available GFP-basedsubcellular markers are constructs that have the ER peptideretention signal, KDEL. By fusing the KDEL sequence to thecoding sequence of GFP and expressing this construct inplants, the dynamic cortical network of membranous tubulesrepresenting the ER could be visualized (Boevink et al., 1996,1998). This new vital ER marker has already provided newinformation on structure/function of the endomembrane sys-tem as well as vesicle trafficking in plants (Hawes, Brandizzi,and Andreeva, 1999). For example, GFP fused to the ER re-


tention signal has not only demonstrated the highly dynamicnature of the plant endomembrane system, but has also re-vealed the existence of rapidly moving organelles (Haseloffand Siemering, 1998). Although these organelles have beensuggested to be proplastids, an alternative explanation as tothe 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 ratsialyl transferase) in combination with a second construct con-sisting of GFP fused to the Arabidopsis H/KDEL homologueof the yeast HDEL receptor, aERD2, the simultaneous visu-alization of the Golgi and ER was possible. This provided thefirst evidence that the ER and Golgi in plants are closely as-sociated (Boevink et al., 1998).

In addition to the ER and Golgi, GFP has also been targetedto the mitochondria (Kohler et al., 1997b), vacuole (Sanse-bastiano et al., 1998), plastids (Kohler et al., 1997a; Tirlapuret al., 1999), and the nucleus (Chytilova, Macas, and Gal-braith, 1999). Among other findings, this work has allowedthe monitoring of the mobility of thin tubular projections orig-inating 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 gen-erating a protein fusion between GFP and a spliceosomal pro-tein, the dynamics of coiled bodies, nuclear organelles in-volved in RNA processing, have been observed in higherplants for the first time (Boudonck, Dolan, and Shaw, 1999).

By creating GFP fusions to known cellular proteins or theirputative targeting peptides, it has also been possible to deter-mine the dynamics and localization patterns of these proteinsin living plant cells. For example, GFP linked to phragmo-plastin, a protein known to be associated with cell plate for-mation, has allowed the dynamics of early cell plate devel-opment to be observed in vivo (Gu and Verma, 1997). Aunique family of calmodulin-dependent Ca21-ATPases were lo-calized to the ER and nuclear envelope of Arabidopsis (Honget al., 1999) and a calmodulin-binding transporter protein wasshown to be associated with the plasma membrane of barleyaleurone protoplasts (Schuurink et al., 1998). Various proteins(e.g., CRY2, COP1, phyB) implicated in light signal trans-duction were linked to GFP and shown to translocate to thenucleus (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 GFPwas demonstrated to localize exclusively to the mitochondria(Galvez et al., 1998). GFP fusions to transit peptides ofphage-type organellar RNA polymerases also allowed local-ization of this protein to the mitochondria and plastids. Thesedata provided additional support for the involvement of thesepolymerases in the transcriptional machinery of plant mito-chondria and plastids (Hedtke et al., 1999). Therefore, withthe availability of several spectral variants of GFP and theability to target GFP to almost any cellular organelle desired,we predict that the direct interactions between these differentorganelles and/or protein fusions will be an area of intenseresearch.

Macromolecular transport and virus movements inplants—Virus movement in plants is a complex process thatis generally considered to involve both cell- to-cell movementvia the plasmodesmata (Reichel, Mas, and Beachy, 1999) andsystemic transport via the vascular system (Nelson and vanBel, 1998). Despite being intensively studied, the pathways

and mechanisms by which viruses moved in plants were poor-ly understood primarily because of an inability to observe theinfection process in live tissue over time (Nelson and van Bel,1998; Santa Cruz, 1999). However, through the observation ofGFP fluorescence from chimeric viruses, researchers can ef-fectively monitor the vein classes by which viruses move afterinfection 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 fromcell to cell are also beginning to come to light. Although theassociation of viral components with some of the host cellcomponents has been reported previously, these associationswere demonstrated mainly through electron and light micros-copy of fixed plant tissues (Nelson and van Bel, 1998). Withmodified viruses expressing GFP fusions to viral proteins, spe-cific localization of viral proteins to plasmodesmata (Itaya etal., 1997; Oparka et al., 1997) and other subcellular compo-nents of the plant host have been elegantly demonstrated. Forexample, by linking the tobacco mosaic virus movement pro-tein (MP) to GFP, colocalization of MP-GFP to cellular actinwas observed and MP was demonstrated to bind actin in vitro(McLean, Zupan, and Zambryski, 1995). MP-GFP has alsobeen shown to localize to fluorescent aggregates and filamen-tous structures in infected BY2 cells. Treatment of infectedcells with microtubule-depolymerizing compounds disruptedthe filamentous strands providing evidence for MP-microtu-bule association (Heinlein et al., 1995, 1998). These studiesall indicate that the local spread of viruses could be facilitatedby interactions between a virus and these components of thehost plant (Reichel, Mas, and Beachy, 1999).

The progress made from studies on plant virus movementhas had a significant impact on plant cell biology and plantphysiology. The ability of viruses to penetrate the physicalbarriers presented by the plant has opened new ideas on thefundamental mechanisms by which macromolecules are trans-ported throughout the plant. These transport stories have re-vealed 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 setof results pointing to a remarkable informational macromolec-ular exchange between plant cells that are likely to coordinatemany aspects of plant development and function (Lucas andWolf, 1999; Oparka et al., 1999). The reader is referred to theseveral excellent reviews published recently that have thor-oughly addressed these issues (Lazarowitz, 1999; Lazarowitzand Beachy, 1999; Santa Cruz, 1999; Thompson and Schulz,1999).

Plant cellular signaling—Another exciting use of GFP fu-sion proteins is to allow the visualization and quantification ofprotein–protein interactions. This technique is called fluores-cence resonance energy transfer (FRET) and relies on the phe-nomenon whereby a donor fluorescent molecule, when excitedwith the appropriate wavelength of light, transfers some of itsemission energy to an adjacent acceptor chromophore. SinceFRET is dependent on the proximity of the donor to the ac-ceptor, measurement of the efficiency of FRET can be used toestimate the distance between donor and acceptor fluoro-phores. Therefore if two different proteins or molecules aretagged with either a donor or acceptor fluorophore, FRET candetermine whether these proteins are in close association (Gor-don et al., 1998).


The concept of FRET in combination with GFP technologyoffers new methods for elucidating signal transduction path-ways in plants. For example, spectral variants of GFP wererecently used to construct a novel intracellular Ca21 sensorreferred to as ‘‘cameleon.’’ This probe was designed so thattwo spectral variants of GFP with overlapping emission andexcitation spectra were fused at opposite ends of calmodulin(a Ca21-binding protein) and the M13 peptide, (the calmodu-lin-binding domain from myosin light chain kinase). The twovarieties of cameleon constructs are the yellow-cameleon,which consists of cyan fluorescent protein (CFP) and the yel-low fluorescent protein (YFP), and the blue cameleon, whichconsists of blue fluorescent protein (BFP) and GFP (Miyawakiet al., 1997).

Cameleons report Ca21 in the following manner. Binding ofCa21 to the calmodulin within the cameleon induces a confor-mational change causing it to bind to the M13 peptide. Thecameleon molecule is folded in the process such that the donor(e.g., CFP) and acceptor (e.g., YFP) are brought in to closerproximity to each other, thereby increasing FRET efficiency.The donor:acceptor fluorescence ratio can then be calibratedto the cytoplasmic Ca21 concentration (Miyawaki et al., 1997,1999). Although the BFP constructs suffer from low signalstrength in plants, plant-expressed CFP constructs are bright(Blancaflor, Dowd, and Gilroy, unpublished data) making theCFP-YFP cameleon the appropriate choice for use with planttissues.

Cameleons were first used to measure intracellular calciumin mammalian cells (Miyawaki et al., 1997; Emmanouilidouet al., 1999), but the yellow cameleon construct has now beensuccessfully engineered into plant cells (Allen et al., 1999;Fricker and Oparka, 1999; Gadella, van der Krogt, and Bis-seling, 1999). Allen et al. (1999) constitutively expressed thisyellow cameleon in Arabidopsis and used this to investigatecytoplasmic Ca21 changes in guard cells. Extracellular Ca21 orABA application induced cytoplasmic Ca21 transients in guardcells, which were similar to the changes observed using fluo-rescent dyes. These results indicate that GFP-based Ca21 in-dicators are suitable for plants and therefore will be of enor-mous value in understanding the complex role of Ca21 in plantcell signal transduction.

In addition to Ca21, pH has also been shown to tightly reg-ulate numerous cellular signaling events and physiologicalprocesses in plant cells (Guern et al., 1991). Like previousstudies with Ca21, measurement of cytoplasmic pH has reliedon fluorescent ratiometric dyes (Bibikova et al., 1998; Frickeret al., 1999; Scott and Allen, 1999). GFP-based pH indicatorshave also been recently developed for monitoring cytoplasmicand organelle pH in mammalian cells (Kneen et al., 1998;Llopis et al., 1998). Various laboratories are attempting to en-gineer these GFP-based pH indicators into plants, and it willbe exciting to see whether these pH sensors function in livingplant cells.

The use of GFP-based ion sensors to study plant signaltransduction is still in its infancy, but is likely to generateinformation not previously obtainable with the traditional fluo-rescent dyes. This is due to the fact that these GFP-based ionsensors can be easily targeted to organelles (see discussionabove); therefore the subtle changes in ions that appear to bepart of the signaling cascades of some plant processes such asgravitropism can now be determined (Legue et al., 1997; Ro-sen, Chen, and Masson, 1999; Scott and Allen, 1999). Fur-thermore, the cameleon indicators can now be imaged with

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 develop-mental biology, labeling of cells to follow cell fate has reliedmainly on the microinjection of dyes into specific cell typesor immunocytochemical detection of specific cell types aftertissue fixation (Brand, 1999). With GFP, it is now possible tomark specific cells or tissues in living Arabidopsis plants andfollow their fates directly during development (Fricker andOparka, 1999; Haseloff, 1999). This was accomplished by us-ing the targeted gene expression approach that uses the GAL4yeast transcription activator. In this ‘‘enhancer-trap’’ strategy,the GAL4 gene is randomly integrated into the Arabidopsisgenome, bringing it under the control of different genomicenhancer sequences that can direct a wide range of expressionpatterns in the plant. However, in order to obtain efficient ex-pression, it was necessary to alter codon usage by employinga derivative sequence (GAL4-VP19). With GAL4-VP19linked to mGFP5, which codes for ER-targeted GFP, the gen-erated expression patterns of GFP in Arabidopsis roots couldbe readily visualized with a confocal microscope (Haseloff,1999).

Using the enhancer trap approach, root-cap-specific pro-moters were recently identified and used to genetically ablate(i.e., selectively kill through the localized expression of a cy-totoxic protein) root caps of Arabidopsis to study the conse-quences that such ablations have on root development (Hase-loff, 1999). With the availability of specifically marked celllines, researchers were also able to follow cellular fate andinfer positional information that contributes to a specific typeof patterning in roots and shoots (Berger et al., 1998a, b; Sa-batini et al., 1999). Cell-specific marking using GFP has alsobeen useful in the identification of endodermal protoplasts forpatch-clamp experiments (Maathuis et al., 1998) and compan-ion cells for single-cell detection of gene transcripts (Brandtet al., 1999).

One of the more exciting prospects of the enhancer trapapproach is the ability to activate other genes of interest in theGFP-marked cell lines. This can be accomplished by subclon-ing the desired gene behind a tandem array of five optimizedGAL4 binding sites (the Upstream Activation Sequences) andtransforming plants with this construct. In the absence of theGAL4 transcriptional activator, however, the desired gene isnot expressed. By crossing this plant with a line with a knownpattern of GAL4 expression, known because of the GFP ex-pression patterns, the silenced gene is activated only in thecells where GAL4 is expressed (Haseloff, 1999). Using thisapproach, it should be possible, for example, to express theGFP-based cameleon or pH indicators in specific cells or tis-sues in the plant and monitor signaling-related ion fluxes inthese cells. Since the fluorescence signal will be coming ex-clusively from specific cells, the problem of interfering signalsfrom other tissues is eliminated. Furthermore, it should alsobe possible to express particular GFP-protein fusions such asthe GFP:MAP4 or GFP:talin fusions described above in spe-cific cells and follow the dynamics of the cytoskeleton as thesecells progress developmentally.

OPTICAL TECHNIQUES FOR MICROMANIPULATIONOF PLANT CELLS

In addition to the improved ability to visualize dynamicstructures and quantify various physiological processes in


Fig. 7. Displacement of mineral-containing vesicles (statoliths) using optical tweezers. The arrow indicates the initial trapping of statoliths on one side ofthe rhizoid. Similar to the graviresponse, rhizoids bend toward the side of statolith accumulation (E. B. Blancaflor, and S. Gilroy, unpublished data). Scale bar5 20 mm.

cells, optical microscopy is now being used as a precise toolto manipulate both the physical and chemical properties of thecell. The fields of chemistry and physics have contributed sub-stantially to the development of these new optical approachesand a representative from each field is discussed below.

Caged probe technology—Caged or photoactivatableprobes provide the opportunity to manipulate cell biochemistryor signaling activities in situ using light. All caged probesoperate on a similar theme. A biologically active molecule isderivatized by a caging group, which then sterically blocks theactivity of the molecule. The caging group is chosen carefullyto be photoactivatable. Thus, when the caging group absorbslight (usually only UV light is energetic enough for these ap-plications), it photolyzes, releasing the biologically active mol-ecule. Some photolysis by-products are also released, and theirpotential biological activity has to be carefully checked. How-ever, the caged molecules used in plant cells to date have re-vealed no indication of biological activity or cytotoxicity ofthese by products.

When the caged molecule is loaded into the cell (e.g., bymicroinjection), then illumination using a standard fluores-cence microscope is all it takes to release the molecule in thecell. More sophisticated setups use a flash UV illuminator orlocalized UV laser irradiation to produce highly controlledtemporal or spatial uncaging. In addition, the more UV lightthat is used, the more probe is activated. Thus the cell biologistcan now control the spatial, temporal, and amplitude aspectsof a molecule’s activity in the living cell. As with fluorescentprobes, the range of caged probes that are available has growndramatically in the last few years. There are now caged hor-mones, amino acids, peptides, drugs (such as caged taxol),nucleotides, ionophores, fluorochromes, and a range of caged-chelators and even caged-nitric oxide (Haugland, 1999). Mostare commercially available, although some of the more spe-cialized reagents must be obtained from the laboratory wherethey were synthesized.

Not surprisingly this technology has been applied to manip-ulate a wide range of control features of animal cells (Adamsand Tsien, 1993). However, though used in several plant stud-ies, its potential has yet to be fully explored by plant cellbiologists. For example, caged Ca21, Ca21 chelators, and ion-ophores have been used to show a requirement for Ca21 in theresponse to GA and ABA in aleurone cells (Gilroy, 1996),phytochrome action (Shacklock, Read, and Trewavas, 1992;

Fallon, Shacklock, and Trewavas, 1993), the signaling path-way of stomatal closure (Gilroy, Read, and Trewavas, 1990)and in the localization of growth in pollen tubes (Fig. 6; Malhoand 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 Trewa-vas, 1990) and pollen tube growth (Franklin-Tong, 1999) andcaged ABA has been used to localize one site of ABA per-ception to an intracellular site in the guard cell (Allan et al.,1994). Given the ability to quantitatively regulate the spatialand temporal release of a caged molecule with subcellular pre-cision, we can predict an increasing application of such ap-proaches to probe the spatial and temporal requirements ofregulation in the plant cell.

Laser micromanipulation—The use of lasers is not onlylimited to providing a coherent light source for microscopicimaging of cells, but has also been used for the micromanip-ulation of whole cells and organelles. Laser micromanipulationcan be divided into two types. The first type, called laser twee-zers or optical trapping, makes use of a continuous-wave, low-power infrared laser beam. The laser beam is focused on anobject at the focal plane of the microscope, which results inthe refraction of the incident laser beam. The refraction of thelaser produces forces that pull the object toward the laserbeam, which can be used to hold the object in place. Whenapplied to cells, moving the laser beam can pull organelles orwhole cells from one place to another (Berns, 1998; Greulichand Pilarczyk, 1998).

One example of the use of laser tweezers in biology hasbeen to address the problem of gravity sensing in the greenalga Chara. Chara is anchored to its substratum via single-cell, root-like structures called rhizoids. The tips of these rhi-zoids contain numerous vesicles that are filled with heavy min-eral deposits (Fig. 7). Upon tilting the rhizoids by 908, thesevesicles sediment rapidly to the new lower flank of the cell.Curvature commences shortly after sedimentation, and it isbelieved that the sedimentation of these vesicles is responsiblefor the initial perception of gravity leading to the redirectionof rhizoid growth (Sievers, Buchen and Hodick, 1996). Lasertweezers have been used to mimic the gravitational displace-ment of these vesicles and have been shown to alter the di-rection of rhizoid growth (Fig. 7; Leitz, Schnepf, and Greulich,1995). By combining laser trapping experiments and imaging


techniques such as fluorescence ratio analysis, it is now pos-sible to link displacement of the mineral containing vesiclesto other cell regulatory elements (e.g., changes in Ca21 distri-bution) that have been proposed to drive growth reorientationduring rhizoid gravitropism (Sinclair and Trewavas, 1997).

Optical trapping techniques have also been applied to studythe physical properties of the actin cytoskeleton in living plantcells. In this case, light from an argon-ion laser is focused ontoa transvacuolar strand and the force needed to displace strand-associated vesicles is measured to give an indication of theviscoelastic properties of actin (Schindler, 1995). Using thistechnique, it was demonstrated that various factors includinglipids, Ca21, pH, aluminum, and the plant hormones auxin andcytokinin alter the tension of the actin cytoskeleton in soybeansuspension cells (Grabski et al., 1994; Grabski and Schindler,1995, 1996). Furthermore, Ca21-regulated kinases and phos-phatases have been identified as possible signaling moleculesthat modulate the changes in the viscoelastic properties of theactin cytoskeleton (Grabski et al., 1998). However, whetherthe transvacuolar strands from which tension measurementswere taken actually represent the actin cytoskeleton remainsquestionable. With the availability of GFP to directly visualizethe cytoskeleton in living plant cells (see above; Kost, Spiel-hofer, and Chua, 1998; Marc et al., 1998), it would be inter-esting to combine optical trapping techniques and fluorescenceimaging to probe further into the physical properties of thecytoskeleton.

The second type of laser micromanipulation is called opticalscissors or laser ablation. This technique relies on the precisedelivery of a high-powered laser beam to a target cell or or-ganelle without affecting the areas surrounding the target (Po-nelies et al., 1994; Berns, 1998; Greulich and Pilarczyk, 1998).With this technique, whole plant cells have been ablated toprecisely define the gravity-sensing cells in root caps of Ara-bidopsis (Blancaflor, Fasano, and Gilroy, 1998, 1999). Signalsthat affect cell fate determination during root developmenthave also been studied with the use of laser ablation (van denBerg et al., 1997; Berger, 1998; Sabatini et al., 1999).

Other applications of laser ablation include exposing theplasma membrane of plant cells by ablating out a small regionof the cell wall to facilitate genetic transformation (Guo,Liang, and Berns, 1995) or to allow easy access of micropi-pettes for patch-clamp experiments (Henriksen and Assmann,1997; Bouget, Berger, and Brownlee, 1998). Laser ablationwas also used to cut a hole in the wall of a single pollen grainto isolate and eventually amplify its genomic DNA (Matsun-aga et al., 1999). It is also now possible to combine lasertweezers and laser ablation to obtain an even more subtle ma-nipulation of plant cells. Buer et al. (1998) demonstrated thatcells of Agrobacterium rhizogenes could be inserted into calluscells of Gingko biloba. They accomplished this by ablating a2–4 mm hole through the cell wall with a pulsed laser beamand moving one or two bacterial cells through the hole byoptical tweezers. Using optimum osmotic conditions, theyachieved a 95% cell survival rate. Therefore this techniquepromises to be an important tool to address questions in plantbiology ranging from plant–microbe interactions to cellular or-ganelle dynamics (Buer et al., 1998). For example, it wouldbe interesting to insert dense particles into statolith-free Chararhizoids (Kiss, 1994) to probe further into the gravity-sensingmechanisms of these cells (Sievers, Buchen, and Hodick,1996).

CONCLUSIONS AND PROSPECTS

The rapid advances in optical probes and imaging technol-ogy make these exciting times for plant cell biology. The threelines of technological advances that have brought us to thisunprecedented view of the dynamic plant cell show no signof slowing. First we can predict a continuing impact of ever-increasing computing power. Image analysis is inherently acomputer-based science and as computing power increases wecan predict that techniques to improve resolution and extracteven more complex information from raw fluorescence im-ages, such as real-time image deconvolution, will becomecommonplace. The second thread of advance will be new andexciting imaging technologies. We have highlighted multipho-ton microscopy as a novel imaging approach with enormouspromise, but this is but one of the many technologies, such asfluorescence lifetime imaging and correlation imaging, on thehorizon. The third line of advance will be in fluorescent andluminescent probes. GFP technology is clearly an importantstep towards the ultimate goal of noninvasively monitoring thedynamics of the components of the plant cell, but there willbe an increasing need for fluorescent probes to monitor notonly levels and distribution of regulators but also their activity.For example, recently, calmodulin activity has been imagedusing novel fluorescent dyes (Hahn, Waggoner, and Taylor,1990; Torok et al., 1998) and a GFP-based probe (Persechiniand Cronk, 1999). These successes suggest that developingprobes to image the activities of cellular regulators is an at-tainable goal.

The most significant advances will be made by combiningthese three areas of technology. For example, fluorescence cor-relation microscopy uses powerful computational analysis offluorescence detected by very sensitive imaging sensors(Haupts et al., 1998; Schwille et al., 1999). It has been usedto look at the changes in parameters such as molecular dif-fusional constants of GFP fusion proteins to image their in-teractions with other cell components (Brock et al., 1999; Trieret al., 1999). Computation, imaging, and fluorescent probeshave come together to reveal the dynamics of the environmentaround a single molecule and have provided a very powerfultool to monitor the activities of the living cell. The applicationof such technologies to the vast array of unanswered questionsin plant development, function, and regulation will be a majorgoal for plant cell biology researchers in the new millennium.

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