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. 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 (C). Besides a cytosolic pattern, ARF1-GFP is also distributed to punctate structures that colocalize with the Golgi apparatus (arrow) and to others that do not colocalize with it (arrowhead). The ER network is visible thanks to the ssYFP-HDEL labeling (open arrow). The time lapse imaging makes possible the visualization of the dynamic relationship between the organelles labeled with the GFP/YFP/RFP fluorochromes. High temporal resolution in acquisition of the signal from each fluorochrome was achieved with the line-sequential multitrack scanning mode configuration of the microscope. Images were acquired using an inverted Zeiss LSM510 META with a 40× oil immersion lens. The fluorescence of ST-mRFP, ssYFP-HDEL, and ARF1-GFP was detected using 594-, 514-, and 405-nm laser lines for excitation of RFP, YFP, and GFP5, respectively. The main dichroic used was HFT 405/514/594, with emission filters 520 to 555 IR and 465 to 510 IR nm (for YFP and GFP, respectively), and the spectral dye separation of the META for the RFP imaging was 609 to 689 nm. Time lapse scanning was acquired with the microscope's imaging system software. Time frame of acquisition is indicated above the segments. D, Merged images of A, B, and C. Insets are magnified sections of merged images (D). Scale bar = 5 μm. 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). 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, and to CLSM TIRFM an technology that makes possible imaging of membrane Although this has not been used for of plant et have very images of the of ER over to that ER may at ER membrane at the or This be a with conventional confocal has a of the of GFP from the et al., Fluorescent proteins are for protein localization, and Although the to be and as protein in plant endomembranes were the GFP and et al., Haseloff et al., fluorescent proteins have been and the visible are to in suitable fluorescent proteins, of the of fluorescent proteins, and for applications et al., 2007; et al., and 2007). 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