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Plant Physiol. Aug 2008; 147(4): 1469–1481.
PMCID: PMC2492624
Focus Issue on Membrane Trafficking

Advances in Fluorescent Protein-Based Imaging for the Analysis of Plant Endomembranes

An exciting new era for live cell imaging of plant endomembranes was ushered in just over 10 years ago when Jim Haseloff and colleagues reported on the removal of a cryptic intron in the sequence of wild-type GFP for successful expression in plant cells (Haseloff et al., 1997). Haseloff's success was quickly welcomed by labs around the globe; by targeting GFP to secretory organelles, scientists could now bring to light the secret life of plant endomembranes. The plant endomembranes comprise several organelles functionally interlinked for the production and transport of secretory compounds, such as proteins, lipids, and sugars. Most of the secretory compounds are synthesized in the endoplasmic reticulum (ER) and then transported to the Golgi apparatus to be sorted for transport to distal compartments such as the plasma membrane and vacuoles. A forward transport of secretory molecules from the ER is counterbalanced by a retrograde flow from distal compartments that allows membrane homeostasis and communication with the cell's surroundings. Fluorescent protein technology applied to live cell imaging of plant endomembranes has aided immensely in providing subcellular markers for the study of the complex spatial and temporal relationships among secretory organelles. Live cell imaging is now moving forward to novel challenges. With the integration of genomic, transcriptomic, and proteomic techniques, we begin to collect data on the putative functions of plant genes; we need to understand the intricate relationships among thousands of proteins. To complete the puzzle, we must understand how the pieces fit together; we must define the functions of gene products. Important questions include: When are our proteins synthesized? Where are they targeted? With what elements do they interact? Advances in the development of optical imaging at the cellular level now offer the exciting opportunity to answer these questions.

In this Update, we discuss several state-of-the-art applications of GFP imaging and how these techniques have been used to answer critical biological questions, with a focus on plant endomembrane trafficking.


Over the past 10 or so years, the visual marker GFP, coupled with optical microscopy (either conventional epifluorescence or confocal microscopy), has become a fundamental tool for plant cell biologists. The successful adoption of GFP and its fluorescent derivatives for live cell imaging of plant endomembranes has generated a strong demand for optical imaging instrumentation based on confocal laser scanning microscopy (CLSM), which eliminates out-of-focus blur while providing temporal resolution. This capability presents obvious advantages when organelles and proteins have to be visualized in living cells rather than in fixed sections. Furthermore, the advent of confocal microscopes with spectral imaging capabilities has provided the possibility of viewing separate overlapping spectra of multiple fluorochromes. Another advantage of CLSM is the ability to minimize interference of innate plant autofluorescence from such sources as cell walls, vacuolar contents, and chlorophylls.

Multiple Fluorochrome Imaging

In response to the increasing demand to visualize more fluorochromes in a shorter time period, sophisticated instruments have become available to perform analyses based on multiple labeling with two or more fluorochromes in live cells. The use of two fluorescent proteins together to establish faithfully the nature of the compartments to which a protein fusion is targeted is becoming increasingly more common; indeed, experiments based on simultaneous imaging of three fluorochromes are also being designed. For example, imaging using a line-sequential multitrack scanning mode of three fluorochromes makes possible the establishment of the dynamic relationship between different organelles in the plant endomembranes at a reasonable time resolution (Fig. 1). With a similar approach, Matheson et al. (2007) used triple labeling based on GFP5, yellow fluorescent protein (YFP), and the styryl-dye FM4-64 to determine the localization of the small GTPase ADP-ribosylation factor 1 (ARF1) with respect to putative endocytic compartments and the Golgi apparatus.

Figure 1.
Triple labeling aids the visualization of multiple endomembrane compartments. Time lapse sequence of a tobacco leaf epidermal cell coexpressing a GFP fusion to the small GTPase ARF1 (ARF1-GFP; A), the Golgi marker ST-mRFP (B), and the ER marker ssYFP-HDEL ...

Spinning-Disc Confocal Microscopy

Spinning-disc confocal microscope technology has improved imaging time resolution in comparison to CLSM, making possible the imaging of live cells in real-time with minimal photo-toxicity. Conventional confocal microscopy relies on laser scanning over a specimen driven by the mechanical movement of mirrors, whereas spinning-disc confocal microscopy is based on the utilization of a spinning disc with multiple pinholes. When coupled to an electron-multiplying CCD digital camera, scanning rates as fast as 1,000 frames/s can be achieved (for review, see Nakano, 2002). Not only does this technology facilitate the imaging of protein localization, it also enables the direct estimation of protein velocity, trajectory, distribution, and activity. Recent applications of this technology provided dynamic imaging of a cellulose synthase subunit (AtCesA6) in the plasma membrane and intracellular organelles as a citrine-YFP fusion (Paredez et al., 2006; Persson et al., 2007). Dual-channel spinning-disc confocal microscopy, using a cyan fluorescent protein (CFP)-tagged microtubule marker, helped elucidate the role of the cytoskeleton in proper deposition of cellulose in elongating tissues. Although technically challenged by particle density, field of view, and photobleaching, the authors also estimated the YFP:AtCesA6 lifetime by the persistence of individual, constantly moving fluorescent particles to be ≥15 min (Paredez et al., 2006). This observation agrees well with biochemical evidence for cotton CesA protein turnover (≤30 min; Jacob-Wilk et al., 2006).

Total Internal Reflection Fluorescence Microscopy

For applications that necessitate increased sensitivity to small changes in membrane fluorescence, total internal reflection fluorescence microscopy (TIRFM) may be an attractive choice (Goodin et al., 2007; Jaiswal and Simon, 2007). The principle of TIRFM is based on reflection rather than transmission. Briefly, incident light focused at the interface of two media having different refractive indices (e.g. glass and cytoplasm) becomes totally internally reflected if the angle of incidence is greater than a critical angle (Schneckenburger, 2005). At the point of reflection, an evanescent electromagnetic field (evanescent wave) is produced that penetrates a short distance into the second medium (e.g. the cytoplasm). The depth of penetration can range between 70 and 300 nm (Schneckenburger, 2005), and the fluorescence of fluorophores entering the evanescent wave can then be detected with high signal to noise ratios. Owing to the limitations of the penetration depth, however, TIRFM is most suitable for monitoring cell surface phenomena. On the other hand, because TIRFM uses a CCD camera for image acquisition, very fast events, which cannot be detected by CLSM, can now be captured. Additional attractive features, including reduced photobleaching, phototoxicity, and background noise, compared to CLSM make TIRFM an ideal technology that makes possible longer imaging periods of membrane surface-linked events. Although this technique has not been used much for studies of plant endomembranes, Goodin et al. (2007) have recorded very interesting images of the formation of ER tubules over time, leading them to suggest that ER tubules may fuse at distinct loci where ER membrane puncta form, at the termini or within tubules. This fine work would be a much more daunting task with conventional confocal microscopy.


Cell biology has undergone a renaissance of sorts since the cloning of GFP from the jellyfish Aequorea victoria (Prasher et al., 1992). Fluorescent proteins are widely accepted tools for studying protein localization, interactions, and dynamics. Although the first to be adopted and adapted as protein reporters in plant endomembranes were the blue-shifted GFP variants, mGFP4 and mGFP5 (Siemering et al., 1996; Haseloff et al., 1997), numerous fluorescent proteins have since been developed and cover nearly the entire visible spectrum. Several excellent guides are available to help in choosing suitable fluorescent proteins, keeping track of the properties of available fluorescent proteins, and verifying their suitability for particular applications (Chudakov et al., 2005, 2007; Shaner et al., 2005; Piston and Kremers, 2007). The following links may also be useful on-line resources for protein spectra and other properties, including references and distributors: http://www.aecom.yu.edu/facs/FluorescentProteins.html; http://nic.ucsf.edu/thornlab/gfps.htm; and http://thalamus.wustl.edu/nonetlab/ResearchF/AllFPs.htm.

The development of these fluorescent proteins has enabled researchers to visualize multiple proteins simultaneously to gain information regarding localization, dynamics, and interactions (Hanton and Brandizzi, 2006). Descendants of wild-type GFP include enhanced GFP, enhanced CFP (ECFP), and enhanced YFP (EYFP). ECFP and EYFP have also been further improved by modifications that have increased the brightness of these fluorochromes and reduced their ability to form dimers. Cerulean CFP, for instance, possesses improved brightness (e.g. quantum yield and extinction coefficient) and fluorescent life-time (Rizzo et al., 2004). Citrine (Griesbeck et al., 2001) and Venus YFP (Nagai et al., 2002) display improved stability at lower pH ranges and increased folding at 37°C. These proteins (Table I) have been used successfully in localization studies in plant endomembranes (for examples, see Paredez et al., 2006; Hanton et al., 2008).

Table I.
List of fluorescent proteins mentioned in this Update with indication of excitation and emission maxima and relevant references

For the red side of the spectrum, DsRed was cloned from the coral Discosoma and added to the fluorescent protein repertoire almost 10 years ago (Matz et al., 1999). Unfortunately, DsRed naturally assembles into tetramers and as such is not well suited for localization experiments due to mistargeting and aggregation. Mutational studies using DsRed have, however, produced the monomeric red fluorescent protein (mRFP1; Campbell et al., 2002). Currently, there are about 10 mRFP1 variants differing in spectral properties, which include mCherry (Shaner et al., 2004, 2005) and mPlum (Wang et al., 2004; Wang and Tsien, 2006). mRFP1 has been adopted successfully for the study of endomembranes. For example, ST-mRFP1 has been used as a marker for the plant Golgi apparatus (Runions et al., 2006). Other red and far-red fluorescent proteins are increasing in popularity in plant endomembrane studies. In our lab, we have successfully used mPlum to mark the Golgi apparatus and the small GTPase ARF1. With mPlum, we have been able to simultaneously covisualize organelles labeled with Venus YFP and mGFP5/CFP (F. Brandizzi and M. Rossi, unpublished data). mCherry has also been successfully used for the labeling of the tonoplast, ER lumen, plasma membrane, and Golgi (Nelson et al., 2007). For other fluorescent proteins in the red side of the spectrum, most recent has been the generation of TurboRFP and TagRFP from the sea anemone Entacmaea quadricolor and its derivatives Katushka and mKate (Merzlyak et al., 2007; Shcherbo et al., 2007). Because of their increase in brightness, these proteins appear to be promising alternatives to mCherry and mPlum. Although the possibility of using red fluorescent proteins is attractive for several applications, chlorophyll autofluorescence may challenge the detection of the fluorescent protein signal. Therefore, the autofluorescence of the plant tissues should be taken into consideration when working with fluorochromes that absorb/emit in the red region of the visible spectrum.

Another advance in fluorescent protein technology has been the development of “photoactivatable” fusion proteins. One of the first to be developed was a photoactivatable GFP (PA-GFP; Patterson and Lippincott-Schwartz, 2002). Activation of PA-GFP by a 413- or 405-nm laser light produces a significant increase in fluorescence emission when the protein is excited at 488 nm. At least two published studies have demonstrated the applicability of PA-GFP for the study of plant endomembranes. The PA-GFP developed by Patterson and Lippincott-Schwartz (2002) has recently been used to study the dynamic relationship between the ER and the Golgi apparatus in tobacco (Nicotiana tabacum) leaf epidermal cells (Runions et al., 2006). The fusion of PA-GFP to the cytosolic domain of calnexin, an integral type I ER membrane protein, made possible a comparison of the dynamics of the ER surface with the moving Golgi stacks labeled with ST-mRFP. In this way, Runions et al. (2006) determined that Golgi stacks move at the same rate and in the same direction as do photoactivated ER membrane proteins such as calnexin. It seems possible, then, that the Golgi apparatus in plants moves with, and not over, the surface of the ER, a possibility that supports the idea of continuity between Golgi bodies and discrete ER export sites on the ER membrane (daSilva et al., 2004; Hanton et al., 2007a). Another version of PA-GFP that is available for plant endomembrane studies was generated by introducing a T207H site mutation in the plant-codon optimized mGFP5 (Sutter et al., 2006). This PA-GFP variant was used to tag the Arabidopsis (Arabidopsis thaliana) KAT1 K+ channel to monitor its distribution and trafficking dynamics in tobacco leaves (Sutter et al., 2006). As mGFP5 may be used in coexpression analyses with YFP with a good spectral separation using the appropriate microscope settings (Brandizzi et al., 2002a, 2004), it would be interesting to determine whether the spectral properties of the photoactivated PA-GFP derived from mGFP5 could be used in combination with YFP for colocalization studies. If this were possible, then this PA-GFP could also be used for three fluorochrome colocalization analyses with a red fluorescent protein fusion for the simultaneous analysis of multiple organelles/proteins. Other photoswitchable proteins have been recently generated, including PS-CFP2 and Dendra2 (Chudakov et al., 2007). The excitation and absorption spectra of these proteins undergo a dramatic red shift upon photoactivatation. Although these proteins are potentially interesting for the study of protein dynamics in plant endomembranes, their applicability has yet to be tested.


Now that the sequences of the genomes of the monocotyledonous (rice [Oryza sativa]) and dicotyledonous (thale cress, Arabidopsis) reference plant species have become available, the next challenge is to assign function to the proteins encoded by these genomes. An important factor influencing functional assignments is protein localization, which can be assayed by any of several methods. With little doubt, one of the most rapid and conventional methods involves stable or transient expression of the products of genes of interest as fluorescent protein fusions. By coexpressing a gene of interest with known fluorescent markers of subcellular compartments, one can assign protein localization or relative distribution of fluorescent protein fusions. This approach is accomplished by simply selecting at least two different fluorescent fusions and appropriate microscope settings for resolved fluorochrome excitation and emission. The optical instrumentation and the multitude of fluorescent proteins that are now available make it possible to gain deeper insights into secretory organelles than ever imagined before. For example, using appropriate markers, researchers have shown that it is possible to distinguish the cis- from the trans-Golgi cisternae (Latijnhouwers et al., 2005; Saint-Jore-Dupas et al., 2006), and Matheson et al. (2008) have been able to separate the tonoplast from the plasma membrane, though these membranes often may appear to be juxtaposed. As plant cells have a variety of endomembrane compartments, major factors influencing the success of colocalization studies are the availability and diversity of specific organellar markers. A recent review by Jaideep Mathur provides an extensive list of proteins and derivatives that have previously been used as markers for endomembranes and many subcellular compartments (Mathur, 2007). This effort has been paralleled by the generation of a Web-based map of a plant cell with a description of organelle functions and available markers, a great on-line resource for teaching and research (http://www.illuminatedcell.com).

In an excellent effort to provide the plant community with a set of common organelle markers, Nelson et al. (2007) have generated a set of vectors containing a wide range of accepted markers tagged with various fusion proteins having either kanamycin or glufosinate resistance. Several of these markers can be used for colocalization studies or as markers for endomembrane studies. The markers are readily available from the Arabidopsis Biological Resource Center stock center (www.Arabidopsis.org) as either DNA stocks (kanamycin and glufosinate resistance) or as stable transformants (kanamycin) in Arabidopsis (Nelson et al., 2007).

Of course, the choice of the fluorochromes to use for colocalization analyses should be driven by the consideration of available instrumentation, such as microscopes with adequate combinations of laser lines, filters, and dichroics, and compatibility with excitation and emission spectra of the fluorescent proteins. In addition to these considerations, when choosing fluorochromes, researchers who aim to study protein localization and traffic in plant endomembranes must take into account factors that may affect the experimental setup. These factors include autofluorescence of the plant cell wall and chlorophylls and the low pH of certain organelles, such as the lytic vacuole, and of the apoplast, which may affect the detection of the fluorescence signal (Tamura et al., 2003; Zheng et al., 2004). It is also generally true that fluorescent proteins have been optimized for mammalian cells that express well at 37°C. As commonly used plant expression systems usually are grown at much lower temperatures, it is possible that the mammalian literature on protein-folding efficiency may not be directly applicable to plant studies.


Proteins involved in membrane trafficking can be identified by the coupled approach of forward genetics and confocal screening of ethane methyl sulfonate (EMS)-mutagenized lines of Arabidopsis stably expressing a fluorescent-marker protein of choice. Mutants showing abnormalities in marker trafficking can then be identified through screening by live cell imaging. Mutated genes responsible for the observed phenotypes can subsequently be identified by mapping. In our lab, for example, we have initiated a large analysis of Arabidopsis EMS mutants stably expressing fluorescent markers of endomembranes to find novel factors that regulate the activity and integrity of organelles of the early plant secretory pathway. Using EMS mutagenesis of stable Arabidopsis plants expressing a secretory GFP retained in the ER (ssGFP-HDEL; Batoko et al., 2000; Brandizzi et al., 2003), followed by confocal microscopy screening, we have been able to identify mutants with obvious alterations of the ER pattern (Fig. 2). With a similar approach, we have also been able to identify Arabidopsis mutants of the Golgi apparatus (Fig. 2) by screening Arabidopsis M2 mutants stably expressing the known Golgi marker ST-GFP (Boevink et al., 1998). This approach is very promising for the identification of novel genes that regulate the activity and morphology of the plant ER and Golgi apparatus.

Figure 2.
Live cell imaging for the identification of novel gene products in plant endomembranes. Confocal live cell images of leaf epidermal cells of Arabidopsis plants stably expressing the ER soluble marker ssGFP-HDEL show the network of tubules characteristic ...

A similar approach has also proven successful in identifying a large number of vacuolar mutants (Avila et al., 2003). In this case, EMS mutagenesis and confocal microscopy were used to identify Arabidopsis plants with aberrant distribution of a GFP fusion to a δ-tonoplast intrinsic protein. With the same approach, Chary et al. (2008) identified a mutant involved in ascribing cell shape and size, dubbed cell shape phenotype-1. Fine mapping identified the gene as a glycosyl-transferase family 20 member, trehalose-6-P synthase/phosphatase. With a similar strategy, a screen based on transgenic Arabidopsis EMS mutant lines expressing a soluble vacuolar marker, GFP-2sc, has been used to identify KAM1, a type II Golgi-localized protein (Tamura et al., 2005). kam1 mutants had a disrupted actin cytoskeleton and showed defects in endomembrane organization. These findings suggest the exciting possibility that KAM1 may be a key component in the Golgi-mediated organization of actin in plant cells (Tamura et al., 2005). Similarly, to identify genes involved in secretory trafficking, Teh and Moore (2007) mutagenized Arabidopsis stably expressing secreted GFP (sec-GFP), a secreted form of soluble GFP. By screening for intracellular accumulation of sec-GFP, they were able to identify two allelic mutants of a single guanine-nucleotide exchange factor for ARF-GTPases (ARF-GEF). This protein, dubbed GNOM-LIKE1, was further shown to be necessary for BFA-induced selective endocytosis from the plasma membrane (Teh and Moore, 2007). Therefore, this kind of screen is a powerful approach to successfully couple cell biology with the ability of forward genetics to uncover new proteins that regulate the function of endomembranes and the integrity of protein trafficking in the pathway and of the secretory organelles.


Colocalization studies can provide a wealth of information regarding not only location, but also protein dynamics. Very often, a protein travels through several compartments as it performs its biological activity (e.g. recycling receptor) and/or in response to various stimuli. Several optical imaging techniques are geared toward estimating the relative amounts of the protein pool and the rates at which proteins are interchanged. Because bioimaging allows noninvasive fluorophore-based analyses, these data can provide strong support for enzymatic mechanisms and molecular functions in a relatively unperturbed environment. Recent developments in bioimaging of endomembranes for protein interactions include fluorescence resonance energy transfer (FRET) and fluorescence complementation protocols. Very interesting and useful reviews with practical considerations for FRET and fluorescence complementation protocols specifically designed for plant cell biology have recently become available (Bhat et al., 2006; Ohad et al., 2007).


FRET is an elegant and powerful technique for testing protein-protein interaction events in living cells. The basic principle of FRET is that energy from an excited donor molecule is transferred to an acceptor fluorochrome when the donor fluorescence overlaps with the absorption spectrum of the acceptor. Such energy transfer can occur when donor and acceptor molecules are in close proximity and therefore are most likely interacting (Jares-Erijman and Jovin, 2003). Generation of a FRET signal depends not only on the overlap of the emission spectrum of the donor fluorophore and the excitation spectrum of the acceptor fluorophore but also on the orientation of the fluorophores toward each other in a distance range of 2 to 10 nm (Gadella et al., 1999). Energy transfer from the donor to the acceptor leads to a reduction in the donor's fluorescence intensity and a decreased lifetime in the excited state. If the acceptor is a fluorochrome, then the FRET process will lead to an increase in the fluorescence intensity emitted from the acceptor. The fluorescent protein pair most commonly used is CFP and YFP; these are usually spliced to the proteins being analyzed. For example, FRET can be measured upon YFP bleaching in a cell coexpressing a CFP and a YFP protein in the pair to be tested for interaction. If the two proteins are in close proximity, the CFP fluorescence emission intensity of the donor will rise concomitantly to a reduction in YFP fluorescence intensity value. The FRET signal can also be quantified by recording the decrease in the fluorescence lifetime of the donor by using fluorescence lifetime imaging microscopy (FLIM) and specialized hardware for this application (Bhat et al., 2006). There are not yet many reports of FRET and FRET/FLIM analyses for studies in plant endomembranes. However, the technique has been successfully used to probe microdomains in the plasma membrane of cowpea (Vigna unguiculata) protoplasts (Vermeer et al., 2004) and barley (Hordeum vulgare) leaf epidermal cells during pathogen attack (Bhat et al., 2005) and to determine the orientation of the VHA-subunits of the head and peripheral stalk of the V-ATPase complex subunits in Arabidopsis protoplasts (Seidel et al., 2005). FRET/FLIM has been applied recently for the study of protein transport and targeting in plant endomembranes. For example, Zelazny et al. (2007) have recently shown that members of two maize (Zea mays) membrane intrinsic proteins, which are important for modulating plasma membrane permeability, ZmPIP1 and ZmPIP2, have different subcellular distribution (Zelazny et al., 2007). ZmPIP1 fusion proteins were distributed in the ER, whereas ZmPIP2 fusions were found in the plasma membrane. In protoplasts coexpressing ZmPIP2, ZmPIP1 was redistributed to the plasma membrane. As shown with FRET/FLIM, this redistribution occurred as a result of the interaction between ZmPIP1 and ZmPIP2. These results suggest that the interaction of ZmPIP1-PIP2 is part of a regulatory mechanism for the regulation of plasma membrane permeability.

Precautions are advisable when attempting FRET analyses in the context of endomembranes. For example, in a crowded environment, such as a membrane, it is possible to observe the multimerization of certain fluorescent proteins (Snapp et al., 2003). This is because GFP and its variants can undergo weak dimerization both in solution and within cells (Zacharias et al., 2002). As the mutations that generate differences in the spectral properties of ECFP and EYFP fluorescent proteins are usually limited in number, there is the possibility that the CFP and YFP may form heterodimers. Therefore, in experimental setups that adopt fusions of CFP and YFP to proteins to be tested for interactions, a FRET signal may be generated by the multimerization of the fluorescent proteins rather than to specific interactions of the proteins that these are fused to. To overcome this possibility, monomeric fluorescent protein variants are particularly attractive for FRET applications to study protein-protein interactions. Lack of dimerization properties increases the confidence that a FRET signal is the result of a genuine interaction between proteins tested in an experiment rather than of dimerization of fluorescent proteins. Along the same lines, because a FRET signal is generated by proteins that are in close physical proximity and not necessarily interacting pairs, it is very important to optimize the FRET protocol with appropriate controls. Such controls may include FRET analyses using proteins that do not interact with the protein of interest or mutants of the proteins of interest that do not interact with each other and/or biochemical analyses (coimmunoprecipitation, for example). As a positive control, it is important to use either proteins that are known to interact or CFP/YFP on the same molecule and preferably in same environment. This control would allow measurement of the maximum possible FRET so as to provide a range by which to judge one's experimental FRET signal.

Fluorescence Complementation

Increasingly popular methods for analyzing protein-protein interactions in plants are the fluorescence complementation protocols such as the split-YFP (or bimolecular fluorescence complementation [BiFC]) and split-luciferase systems (Bracha-Drori et al., 2004; Walter et al., 2004; Fujikawa and Kato, 2007).

Split YFP

In the split-YFP system, the N terminus half of YFP is fused to a protein and the C terminus of YFP to a putative interacting partner. In theory, detection of the YFP signal in cells coexpressing the two fusions signifies protein-protein interaction events, as the YFP signal is detectable when the N- and C-terminal halves come in close proximity to form an intact YFP molecule. This system is attractive because it is easy to set up and does not require sophisticated imaging instruments, but it comes with a few caveats. The generation of an intact YFP molecule can be considered an irreversible process (Magliery et al., 2005); a stable interaction may be established between the two proteins fused to the YFP halves. Although this may not be a problem for proteins that dissociate with a very slow turnover, the formation of stably interacting protein couples may lead to physiological artifacts for those protein-protein events that are based on fast dissociation rates, for example, the interaction of a GTPase-like ARF1 with its effectors (Stefano et al., 2006; Matheson et al., 2007). Therefore, during the course of an experiment (i.e. from cell transformation to microscopy analyses), BiFC may introduce elements that lead to events that do not reflect normal cell physiology and these must be considered for correct interpretation of the results. Another point to consider is that in conditions of elevated availability of the YFP halves, YFP molecules may be generated simply because YFP halves tend to associate (Zamyatnin et al., 2006). It has also been reported that overexpression in mammalian cells of N- and C-terminal YFP peptides alone resulted in a low fluorescence emission signal; thus, the levels of expression of the split-YFP halves may be a factor to consider in BiFC experiments (Ozalp et al., 2005). Therefore, BiFC seems to be an exciting technique for studying protein-protein interactions, given appropriate controls. A good starting point may involve fusion of one half of YFP to proteins that are known not to interact with the proteins in analysis but that are localized in the same cell compartment and have the same orientation with respect to the membrane as the split-YFP fusions.

BiFC can also be used to determine protein orientation with respect to membranes. One daunting task in the study of proteins associated with endomembranes is to determine how the N and C termini of the proteins are oriented with respect to the membranes. This information may be very important when the position of functional amino acid domains needs to be established. As an alternative to the conventional biochemical assays based on proteinase protection, an in vivo protease protection assay has been developed in mammalian cells (Lorenz et al., 2006). This assay is based on monitoring the fluorescence signal from live cells expressing fluorescent protein fusions to the N or the C terminus of membrane proteins over time. In this experimental setup, cells are permeabilized and then treated with proteases. If the fluorescent signal fades with time due to proteolysis of the fluorescent protein, then the membrane protein is oriented with the fluorescent protein in the cytosol. Although this method is seemingly powerful, it may require careful controls to ensure that the disappearance of the fluorescence signal is not due to technical issues, such as photobleaching or excessive membrane permeabilization that may drastically alter the cell environment. Recently, Zamyatnin and colleagues demonstrated the usefulness of BiFC to determine membrane protein topology; model proteins used were the beet yellows virus-encoded p6 membrane-embedded movement protein, a protein with known topology, and the potato mop-top virus-encoded integral membrane TGBp2 protein with predicted topology (Zamyatnin et al., 2006). This application is based on the property of free split-YFP fragments to associate in plant cells as a result of their overexpression from strong promoters (Walter et al., 2004). In the topology study, YFP complementation was followed using various combinations of cytosolic and ER-targeted split YFPs in cells coexpressing the model proteins fused to complementary split YFPs. In this setup, the orientation of the N and C termini was deducted on the basis of fluorescence complementation with either the cytosolic or ER-luminal split YFP.

Split Luciferase

Another approach to analyze protein-protein interactions is the split-luciferase system in which the luciferase reporter gene is split into N-terminal and C-terminal halves. These halves are then used to tag two putative interacting proteins. If the two proteins interact, the luciferase halves will be in close proximity to permit bioluminescence upon addition of the luciferin substrate. The advantages of this system are many. First, an expensive, sophisticated microscope is not required, because imaging is performed by a CCD camera. As imaging is based on bioluminescence rather than excitation, background interference of autofluorescent compounds is not an issue. There is no photobleaching of the substrate, as lasers are not used to image samples. This system also allows whole organism imaging, in vivo. This system has been used recently to determine interaction between Arabidopsis SNARE pairs (Fujikawa and Kato, 2007). In this study, Arabidopsis protoplasts were transiently transformed in 96-well plates with the N- and C-terminal fragments of Renilla reniforms luciferase fused to bait and prey SNARE proteins SYP51 and SYP61. This technique should make it possible to screen for interactions among the large number of Arabidopsis SNARE proteins and therefore to provide further clues as to functional relationships among this class of proteins. As with BiFC, the split-luciferase technique will require appropriate controls to ensure that generation of a fluorescence or bioluminescence signal is the genuine consequence of protein-protein interaction rather than the result of overexpression of split-protein pairs.


Fluorescent proteins have been useful for studying plant endomembranes, not only for localization studies but also for quantitative analyses on membrane traffic. Here, we discuss several of these applications.

GFP has been used to assay movement of membrane and soluble protein cargo from the ER along the secretory pathway. For example, changes in the distribution of membrane GFP-Golgi markers have been followed in live cells to test the effect of overexpression of Sec12, the activating protein of the small GTPase Sar1, and the negative dominant mutants of regulatory small GTPases, such as Rab1, ARF1, and Sar1, on membrane transport from the Golgi and on Golgi integrity (Batoko et al., 2000; Takeuchi et al., 2000, 2002; Lee et al., 2002; daSilva et al., 2004; Stefano et al., 2006). Similarly, soluble secretory forms of GFP have been instrumental in analyzing protein traffic from the ER. Soluble secreted and vacuolar-targeted GFP forms expressed in nonsaturating conditions of ER export are usually not visible in these compartments due to low pH quenching and proteolysis events (Di Sansebastiano et al., 1998; Batoko et al., 2000; Tamura et al., 2003; Zheng et al., 2004). Therefore, these forms of GFP have been useful markers to determine factors that perturb their delivery to the final destination by visualization of accumulation of GFP fluorescence in the ER (i.e. Batoko et al., 2000).

GFP for Quantitative Analyses

Beyond qualitative results on the role of protein mutants that may cause accumulation of secretory fluorescent protein fusions in the ER, it is often useful to provide quantitative data. It has been shown that the fluorescence of a sec-GFP can be quantified from low-magnification confocal images of tobacco leaf epidermal cells transformed with Agrobacterium (Zheng et al., 2005). This approach has been used, for example, to estimate the accumulation of sec-GFP signal in the ER in the presence of dominant inhibitory mutants of Rab GTPases (Kotzer et al., 2004; Zheng et al., 2005). Although this method is powerful, interpretation of the results may be complicated by the stochastic nature of the transfection process that by nature leads to a large cell-to-cell variability of expression.

To overcome these limitations, Samalova et al. (2006) have developed a polyprotein-based system for ratiometric fluorescence imaging assays. In this approach, the authors have used the 20-residue, self-cleaving 2A peptide from the foot and mouth disease virus to generate polyproteins that express a fluorescent trafficked marker in fixed stoichiometry with a reference fluorescent protein in a different cellular compartment. With this approach, two fluorescent markers encoded as polyproteins have a significant correlation in their relative abundance. This feature presents a real advance for measuring the perturbations of the secretory activity. For example, the system can be used to follow the retention of soluble sec-GFP in the ER in relation to an ER-retained YFP in the presence of effectors in cell populations using low-magnification imaging and in individual cells at high magnification. Furthermore, if the effector protein (e.g. the dominant mutant of a GTPase) can be expressed as an active fluorescent protein fusion, the system presents an additional advantage in that the relative abundance of the effector can be determined directly in each cell and correlated with its effect on marker trafficking. The 2A-peptide system can also be used to express an untagged effector and a reference fluorescent protein encoded on the same vector, with the advantage of enabling expression of the untagged effector among cells by measuring the fluorescence of the reference marker. This approach appears to be more reliable in comparison to systems based on expression of two proteins transcribed divergently from the same cauliflower mosaic virus 35S enhancer elements. An analysis of the expression of two fluorescent ER luminal proteins, ssYFP-HDEL and ssGFP-HDEL, in these conditions showed a considerable variation in the relative expression of the two proteins (Samalova et al., 2006).

GFP and its spectral variants have also been useful as reporters for biochemical analyses on protein traffic and membrane protein orientation. As antibodies that recognize GFP and its variants are commercially available, the trafficking of GFP markers can be followed by cell fractionation analyses. For example, the arrival in the lytic vacuole of GFP-BP80, a prevacuolar marker, can be monitored by the detection of a characteristic GFP degradation product, termed the GFP-core (daSilva et al., 2005). Therefore, the perturbation of the BP-80 route to the vacuole can be monitored by following the relative abundance of the GFP-core on western-blot analyses. In addition, secretory GFP forms bearing a glycosylation peptide, glycosylated GFP, have been shown to be efficiently glycosylated in the plant ER (Batoko et al., 2000; Hanton et al., 2005). Therefore, they can be used as reliable markers to follow protein translocation into the ER and protein progression through the Golgi apparatus via endoglycosidase (ENDO-H) digestion (Hanton et al., 2007b). GFP has also been used as a tag in proteinase protection assays to determine the orientation of proteins with respect to the target membrane (Hanton et al., 2005; Renna et al., 2005).

Fluorescence Recovery after Photobleaching

GFP and its spectral variants have also been used in fluorescence recovery after photobleaching (FRAP) analyses to study protein traffic in plant endomembranes. FRAP is a powerful technique that enables the estimation of protein mobility within live cells. Exposing a subcellular population of fluorescently tagged fusion proteins to high-intensity laser light for a period of time causes irreversible fluorophore photobleaching within a defined area. Over time, if surrounding nonphotobleached fusion proteins are allowed to migrate into the photobleached region, a fluorescence recovery in the bleached area will occur. Measurement of the fluorescence recovery over time allows direct estimation of protein mobility. Recovery rates in FRAP experiments are influenced largely by temperature, viscosity of the medium, and particle radius (Lippincott-Schwartz et al., 2003). Membrane proteins, for example, exist in a more viscous medium compared to free-cytosolic proteins and therefore have lower expected diffusion rates. FRAP can facilitate estimates of other factors that influence protein mobility, such as active transport and compartmental confinement. In studies of plant endomembranes, FRAP has been used relatively recently in establishing the nature of the factors that influence protein traffic dynamics from the ER to the Golgi apparatus (Brandizzi et al., 2002b) and the dynamics of the proteins that regulate ER-to-Golgi protein transport (daSilva et al., 2004; Yang et al., 2005). With a similar approach, it has been recently shown that the small GTPase ARF1 binds to and is released from Golgi membranes at a rapid turnover rate (Stefano et al., 2006; Matheson et al., 2007). ARF1 is known to recruit coatomer to Golgi membranes. However, it has also been shown recently that ARF1 is capable of recruiting an additional effector, the golgin GDAP1, to the Golgi and post-Golgi organelles in plant cells (Matheson et al., 2007). FRAP analyses on fluorescent protein fusions of ARF1, coatomer, and GDAP1 have shown that the cycles of binding and release of ARF1 to and from the Golgi apparatus are similar to those of GDAP1 but faster than coatomer (Fig. 3). This observation has led to the suggestion that ARF1-GTP hydrolysis is not the sole factor influencing the association/dissociation rate of ARF1 effectors to membranes.

Figure 3.
Selective photobleaching enables measurement of protein dynamics. FRAP experiments on a cortical section of tobacco leaf epidermal cells expressing GDAP1-YFP (A), coexpressing YFP-GDAP1 and an untagged ARF1-GTP mutant (B; Stefano et al., 2006), expressing ...

A FRAP variant based on selective photobleaching of one fluorochrome in cells expressing two fluorescent proteins has also been developed and adopted for the study of plant endomembranes (daSilva et al., 2004). Specifically, for this technique, high-intensity laser light is used to selectively photobleach one fluorochrome (YFP) linked to a protein residing within the same subcellular space where a CFP fusion is localized; recovery of fluorescence in the YFP channel is then measured. This technique is particularly useful when a CFP fusion can be adopted as a reference point for moving organelles. For example, daSilva et al. (2004) adapted this technique to follow the recovery of fluorescence in Golgi bodies of cells coexpressing the H/KDEL receptor (ERD2) fused to YFP or to CFP. Fluorescent protein fusions to ERD2 are known to be localized in the ER and in the Golgi bodies. daSilva et al. (2004) bleached the YFP fluorescence and followed its recovery in Golgi bodies that were visible through the CFP channel. They were able to demonstrate that movement of cargo proteins in and out of the Golgi apparatus may occur during Golgi movement in plant cells. An alternative FRAP protocol may be based on the simultaneous photobleaching of two fluorochromes, such as CFP and YFP, fused to different proteins that are targeted to the same compartment. By following the recovery of the fluorescence of both proteins, it may possible to obtain a simultaneous comparison of the dynamics of two different proteins in the same confined environment.

Fluorescence Correlation Spectroscopy

An emerging technique for measuring endomembrane protein dynamics is fluorescence correlation spectroscopy (FCS). FCS makes possible the quantitation of fluorescence emission over time within a defined, small subcellular volume (1 fL), and therefore enables the direct estimation of protein concentrations and diffusion constants (Krichevsky and Bonnet, 2002). FCS provides information regarding protein dynamics, abundance, and microenvironment. This technique is often used for examining fluorophores in solutions. However, the development of more sophisticated commercially available microscopes is making possible the application of FCS to live cells. Recently, FCS was used in Arabidopsis protoplasts to show that the diffusion coefficients of YFP fusions to the AAA ATPase CDC48A examers, which may be involved in the ER-associated degradation pathway and in membrane fusion events, were variable depending on the cellular compartment. These fusions were also found to be much larger than expected; thus, they may form part of larger complexes (Aker et al., 2007).

It is obvious that plant endomembrane researchers are taking advantage of the latest advances in optical imaging science. We look forward to finding out in the next Update what novelties will have been adopted and what exciting results will have been produced to further our understanding of how secretory organelles communicate with each other.


Given the limitations on the length of this article and the breadth of endomembrane imaging in plants, we apologize to colleagues whose excellent work was not cited in this review. We are grateful to Professor Chris Hawes for providing nonmutagenized ssGFP-HDEL and ST-GFP Arabidopsis plants. We thank Karen Bird for editing the manuscript. The American Society of Plant Biologists is acknowledged as copyright holder of the content of Figure 3.


The author responsible for distribution of materials integral to the findings presented in this article in accordance with the policy described in the Instructions for Authors (www.plantphysiol.org) is: Federica Brandizzi (ude.usm@zzidnarb).



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