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. 2017 Nov 22;176(1):80–93. doi: 10.1104/pp.17.00962

Advances in Imaging Plant Cell Dynamics1,[OPEN]

George Komis 1, Dominik Novák 1, Miroslav Ovečka 1, Olga Šamajová 1, Jozef Šamaj 1,2
PMCID: PMC5761809  PMID: 29167354

Abstract

Advanced bioimaging uncovers insights into subcellular structures of plants.

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After the establishment of advanced fluorescence microscopy methods and the development of numerous fluorescent proteins, it is possible to follow the organization and dynamics of most organelles and subcellular compartments in cells of living plants. Nowadays, it is possible to address subcellular architecture at the nanoscale through the implementation of superresolution microscopy methods such as structured illumination microscopy, photoactivation localization microscopy, or stochastic optical reconstruction and stimulated emission depletion microscopy. In a developmental context, the dynamic cellular and subcellular changes can be monitored long term in whole plant organs by light-sheet fluorescence microscopy. This is a mesoscopic method offering high-speed imaging, very low phototoxicity, and bioimaging of vertically oriented plants. This Update aims to provide the principles, the current application range, and the expected potential of superresolution and light-sheet fluorescence microscopy methods as well as a brief description of the improvements of standard wide-field epifluorescence and confocal systems.

METHODS OF SUPERRESOLUTION MICROSCOPY

Superresolution microscopy is a term that collectively refers to various techniques that bend or overcome diffraction limitations that strictly define the resolution limit of classical far-field optical microscopy systems. Resolution in conventional far-field systems is proportional to the wavelength of excitation light and inversely proportional to the numerical aperture of the light-collecting lens.

One major category of superresolution microscopy includes techniques that illuminate the sample by means of patterned light, such as structured illumination microscopy (SIM; linear and nonlinear; Gustafsson, 2000; Rego et al., 2012) and stimulated emission depletion microscopy (STED; Dyba et al., 2003).

The second major category of superresolution techniques includes single-molecule localization strategies that interrogate the positioning of individual fluorophores with subdiffraction accuracy. Such techniques mainly include photoactivation localization microscopy (PALM; Betzig et al., 2006), stochastic optical reconstruction microscopy (STORM; Rust et al., 2006), and some related variants such as superresolution optical fluctuation microscopy (SOFI; Dertinger et al., 2010a, 2010b), Bayesian analysis of blinking and bleaching (3B), and superresolution radial fluctuations (Cox et al., 2011; Small and Parthasarathy, 2014; Gustafsson et al., 2016). These so-called pointillistic superresolution techniques exploit properties of specific fluorophores to undergo repeated transitions between on and off states under certain conditions (Dempsey et al., 2011; Vaughan and Zhuang, 2011).

SIM

SIM imposes a spatially modulated light pattern (generated by means of a grating with an effective spacing of λexc/2; Gustafsson, 2000) that combines with diffraction orders of the emitting sample to Moiré patterns. Such Moiré patterns are generated by rotations (either three or five rotations at 36° or 60° increments, respectively) and phase shifts (three or five at 2π/3 increments) of the grating. Individual images from each position of the grating are then combined, deconvolved, and used to reconstruct an image, which by definition is twice resolved compared with the Abbe limit (Gustafsson, 2000). Since linear SIM can rescue up to two diffraction orders, it is still diffraction limited. However, by means of linear SIM combined with high numerical aperture objectives (Komis et al., 2014), microtubules that were simply visualized using GFP-labeled tags (GFP-MBD [Marc et al., 1998] and GFP-TUA6 [Shaw et al., 2003]) could be practically resolved to the theoretical 100-nm threshold of linear SIM (Komis et al., 2014, 2015). This is twice as good as the theoretical optimum of 200 nm predicted by Abbe’s equation. The Z-resolution can be significantly better compared with confocal laser scanning microscopy (CLSM; Sahl et al., 2017, and refs. therein). The final spatial resolution achieved by SIM is dependent on the specifications of the platform used for image acquisition, since different commercial platforms employ different means to generate patterned light as well as different numbers of rotations and phase shifts (discussed by Komis et al. [2015]). The temporal resolution of SIM is limited by the mechanical rotations and phase shifts of the light pattern. Thus, reconstructed time series of highly dynamic structures can suffer from motion artifacts (Förster et al., 2016). However, the acquisition frame rate of any commercial SIM platform is sufficient to record moderately dynamic events such as end-wise length fluctuations of microtubules and tracking minor growth/shrinkage events with a length below 200 nm. On the basis of differences in the signal intensity, SIM allows one to discriminate the dynamics of very proximal or bundled microtubules even if these cannot be resolved optically (Komis et al., 2014). Although not as effective as STED (see below) or PALM/STORM in superresolving intracellular structures, linear SIM offers several advantages for cell imaging. It is fluorophore unlimited, moderately fast (with imaging speeds ranging from 100 ms for a single image with Deltavision OMX to 600 ms with Nikon NSIM or 1.2 s with Zeiss Elyra S.1), and well represented in the market (Komis et al., 2015).

Nonlinear SIM, whereby emission of the fluorophore is not linearly proportional to excitation illumination, is able to truly superresolve intracellular structures such as microtubules in a diffraction-unlimited manner (Rego et al., 2012). However, appropriate platforms are not yet widely accessible to the microscopy community.

SIM can be used to image living cells of variable sizes. Thus, live SIM studies have been conducted in bacteria (Bottomley et al., 2017), showing the subcellular distribution of RNA polymerase (Stracy et al., 2015) and the time-resolved assembly of the FtsZ ring during bacterial division (Strauss et al., 2012). 2D and 3D SIM can temporally resolve subcellular structures of small eukaryotic cells, such as the spindle pole body (Fig. 1, A–F) and mitotic spindle (Fig. 1, G and H), or the reorganization of the actin cytoskeleton during cell fusion in fission yeast (Dudin et al., 2015). In custom-built systems, 2D and 3D SIM has been reportedly used at video frame rates and documented the organization and dynamics of microtubules (Kner et al., 2009; Schvartz et al., 2017) and mitochondria (Shao et al., 2011) in mammalian cells.

Figure 1.

Figure 1.

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Comparative structured illumination and wide-field epifluorescence microscopy imaging of Schizosaccharomyces pombe cytoskeletal structures. A to C, Newly separated spindle pole bodies in a premitotic cell (A and B) and their clear discrimination based on normalized fluorescence intensity profiling (C) by means of SIM in S. pombe cells expressing the GFP-pcp1 spindle pole marker. D to F, Spindle bodies as visualized by wide-field epifluorescence (D and E) with less prominent resolution (F) compared with SIM. G and H, Mitotic spindle elongation visualized by SIM (G) and wide-field epifluorescence (H) in S. pombe cells expressing an mCherry-atb2 microtubule marker. Arrows indicate spindle pole bodies (A and D) and mitotic spindle poles (G and H). Bars = 2 µm (A and D), 1 µm (B and E), and 5 µm (G and H).

Depending on the optics and, especially, the objective numerical aperture, intracellular structures can be dynamically visualized at significantly better resolution than 100 nm (Li et al., 2015). In cases where the optical path can be split between two or more cameras, there is a possibility for dual or multiple-channel dynamic imaging of two or more subcellular compartments at the same time (for review, see Hirano et al., 2015; Li et al., 2015). Time-lapse 3D SIM imaging has been markedly improved by combining SIM and light-sheet fluorescence microscopy (LSFM) into a lattice light sheet with exceptional frame rates of acquisition (Chen et al., 2014).

In plants, 2D SIM can visualize various superresolved subcellular structures. These structures may include plasma membrane microdomains containing FLOTILIN1 (Flot1; Fig. 2A), actin filaments (Fig. 2, B and C), microtubules (Komis et al., 2014, 2015), and late endosomes (Fig. 2, E–H), providing quantitative information on the resolution improvement (Fig. 2D). Rare events such as vesicle fusion, which cannot be resolved by wide-field epifluorescence methods, can be captured by SIM (Fig. 2, I and J).

Figure 2.

Figure 2.

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The superresolution potential of structured illumination microscopy applied to subcellular live imaging of plants. A, Imaging of Flot1 clusters at the plasma membrane of an Arabidopsis cotyledon pavement cell expressing a GFP-Flot1 marker. The inset shows two linear clusters of Flot1 particles (arrows). B to D, Imaging of the actin cytoskeleton in a hypocotyl epidermal cell expressing a YFP-FABD2-YFP actin marker (B; the boxed area is magnified in C) with the capacity to resolve fine actin bundles at approximately 100 nm at full width at half-maximum (FWHM) of the intensity profile (D) shown in C. E to J, Visualization of late endosome fusion with SIM imaging in root epidermal cells or Arabidopsis expressing a GFP-FYVE late endosomal reporter (E and F) as compared with wide-field epifluorescence (G and H). The graphic depiction of the longitudinal (I) and the transverse (J) profiles, drawn as dotted lines in E, highlights the potential of SIM to identify the area of fusion as an area of decreased fluorescence intensity. Bars = 10 μm (A and B), 2 μm (C), and 1 μm (E, G, and inset in A).

2D and 3D SIM also can elucidate the structures of organelles such as plastids (Fig. 3, A and B), mitochondria (Fig. 3, C and D), nuclei (Fig. 3, E–H), and the endoplasmic reticulum (Fig. 3, I and J). Moreover, 3D SIM was used successfully to decipher the architecture of plasmodesmata (Fitzgibbon et al., 2010; Knox et al., 2015), address the structure of virus replication complexes (Linnik et al., 2013), and localize the positions of viral movement proteins in plasmodesmata (Tilsner et al., 2013).

Figure 3.

Figure 3.

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Structured illumination microscopy can deliver superresolved images of various plant organelles as compared with wide-field (WF) epifluorescence imaging. A and B, Plastids labeled by the marker ADP-sugar pyrophosphatase fused with GFP. C and D, Mitochondria labeled by the mitochondria-targeting sequence of the γ-subunit of the F1-ATPase fused with GFP. E and F, Nuclear structures of root epidermal cells expressing a nuclear localization signal-GFP marker. G and H, Nuclear structures of root epidermal cells expressing an HTB-mRFP marker. I and J, Structures of endoplasmic reticulum in root hairs stably expressing a GFP-HDEL marker. Bars = 2 μm (A–D, I, and J) and 10 μm (E–H).

Nevertheless, time-lapse imaging using either 2D or 3D SIM is still very scarce in plant science. In the 2D mode, time-lapse SIM revealed superresolved cortical microtubule dynamics (Komis et al., 2014) and the dynamics of Flot1 under resting conditions or after flg22 treatment (Yu et al., 2017). In both cases, the subject of interest was confined at the cell periphery, being positioned close to the coverslip and, thus, facilitating image acquisition. In the same manner, other cortical structures, including the endoplasmic reticulum (Fig. 4, A and B) and the actin cytoskeleton (Fig. 4, C and D), can be tracked over time for relatively long periods.

Figure 4.

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The potential for time-resolved structured illumination microscopy imaging. A and B, Overview (A) and time-lapse imaging (B) of cortical endoplasmic reticulum in a hypocotyl epidermal cell expressing a GFP-HDEL marker. The dynamic reorganization of the endoplasmic reticulum tubular structure is depicted by arrowheads (B). C and D, Overview (C) and time-lapse imaging (D) of actin filaments in a hypocotyl epidermal cell expressing a GFP-FABD2 marker. The dynamic reorganization of actin filaments is depicted by arrowheads (D). Bars = 5 μm (A and B) and 10 μm (C and D).

SIM also can be used for more advanced quantitative studies, which may include the colocalization of different subcellular structures. Colocalization studies using SIM combined with single-particle averaging were used to map the positions of individual components of complex subcellular organelles such as the spindle pole body of fission yeast (Burns et al., 2015; Bestul et al., 2017). Finally, it was shown that fluorescence recovery after photobleaching (FRAP) analysis can be adopted by SIM to provide quantitative information on protein kinetics (Schneider et al., 2013).

SIM is prone to spherical aberrations due to refractive index mismatches. This may hamper the visualization of the subcellular architecture of plant samples not adequately coated by the observation medium. It can be avoided by mounting plants in an oxygen-saturated perfluorocarbon (Littlejohn et al., 2010, 2014; Littlejohn and Love, 2012; Chi and Ambrose, 2016). SIM also is prone to optical artifacts that may be caused by inaccurate grid movements (Langhorst et al., 2009), and resolving capacity depends on the labeling quality and the signal-to-noise ratio (SNR) of the sample. If the object in focus is dimly labeled and obscured by out-of-focus light or background fluorescence, the contrast of the grating pattern will be deteriorated and will not allow the calculation of the final image at the expected high resolution (Langhorst et al., 2009). Moreover, SIM is heavily dependent on postacquisition image reconstruction, while settings of the deconvolution algorithms may lead to artifacts in image reconstruction (for review, see Komis et al., 2015; Demmerle et al., 2017).

The principle of structured illumination can be combined with other microscopy methods, such as spinning disk to deliver superresolution images or wide-field epifluorescence to reduce out-of-focus readouts and provide optical sectioning of the sample. In the first case, the speed of imaging using the combination of SIM and spinning disk can achieve temporal resolutions between 30 and 100 frames per second (Hayashi and Okada, 2015). To our knowledge, a hybrid system of SIM-spinning disk is offered as an add-on for wide-field epifluorescence microscopes provided by Andor (DSD system; http://www.andor.com/learning-academy/revolution-dsd-principles-of-operation). The Zeiss Apotome system is a structured illumination add-on for wide-field microscopes that markedly improves image quality and allows for optical sectioning minimizing out-of-focus blur (https://applications.zeiss.com/C125792900358A3F/0/05BA0ED63B5E4CA2C1257E85004AB36A/$FILE/EN_41_011_081_ApoTome2.pdf).

Single-Molecule Localization Microscopy

Single-molecule localization microscopy (SMLM) includes all techniques that improve the precision of localization of single fluorophores based on their blinking (transition between emitting and dark states) under redox conditions (STORM; Rust et al., 2006; Dempsey et al., 2011) or on the localization of photoactivatable, photoswitchable, or photoconvertible fluorescent proteins (PALM; Nienhaus and Nienhaus, 2016). The precision of localization itself depends on the number of photons collected from a single molecule, the pixel size of the detector, the background signal, and the sd of the point-spread function. Therefore, it is important to acquire images at high SNR using cameras with low noise. Thus, PALM/STORM acquisitions are preferably performed using a total internal reflection microscopy (TIRF) illumination mode and electron microscopy CCD cameras.

The resolution of SMLM techniques can be optimally 10 times better than Abbe’s limit (i.e. approximately 20 nm), but these techniques suffer from extreme limitations related to sample stability and fluorophore performance during acquisition. Since they investigate fluorophore localization at tens of nanometer precision, it is essential that the sample should be very firmly immobilized, and the examined structures must exhibit negligible dynamics during the thousands of time frames needed for image acquisition (Halpern et al., 2015). The two most important fluorophore properties affecting the performance of SMLM are the number of photons detected for every switching event (transition between on/off states) and the duty cycle of the fluorophore, which represents the fraction of time that the fluorophore spends in the on state. For optimal SMLM, a high photon yield and a low duty cycle of the fluorophore are desirable (Dempsey et al., 2011). Therefore, SMLM imaging is fluorophore limited, especially regarding genetically encoded fluorescent proteins. Although plant codon-optimized photoactivatable/photoswitchable proteins (Lummer et al., 2011, 2013) and photoconvertible proteins (Hosy et al., 2015) are available, these have been used very rarely for SMLM imaging of plant samples so far. One such study employed PALM to investigate the perinuclear organization of the actin cytoskeleton in living tobacco (Nicotiana tabacum) BY-2 suspension cells labeled with the tetrameric variant of photoswitchable EosFP (IrisFP) and achieved resolution of approximately 50 nm (Durst et al., 2014). Another study utilized fusions of the monomeric photoconvertible protein mEos3.2 with plasma membrane or tonoplast proteins to decipher the kinetics of their motility by means of single-particle tracking PALM (Hosy et al., 2015). Our provided example of PALM imaging shows the nuclear distribution of DRONPA-tagged plant-specific END BINDING1c (EB1c), a member of the microtubule plus end-binding protein family of Arabidopsis (Arabidopsis thaliana), in comparison with the diffuse nuclear signal observed by wide-field microscopy (Fig. 5, A–C). Moreover, PALM localization is more effective then SIM in deciphering EB1c nuclear distribution (Fig. 5, D and E).

Figure 5.

Figure 5.

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Photoactivation localization microscopy of EB1c protein in root epidermal cells of Arabidopsis expressing an EB1c-DRONPA marker and STED of actin filaments in Arabidopsis cotyledon epidermal cells expressing a GFP-FABD2 actin marker. A to C, Wide-field (WF) epifluorescence (A) and PALM (B) imaging and quantitative assessment (C) of the precision of localization of B at approximately 40 nm. D and E, Comparison of SIM (D) and PALM (E) localization of EB1c. F and G, Comparison of CLSM (F) and STED (G) imaging of actin filaments in Arabidopsis cotyledon epidermal cells. H and I, Volumetric CLSM (H) and STED (I) imaging of actin microfilament organization in Arabidopsis cotyledon epidermal cells. Bars = 2 μm (A, B, D, and E) and 10 μm (F–I).

STORM is heavily dependent on the photochemical properties of the fluorophores. This method is used preferentially for fixed and immunolabeled probes. Moreover, the blinking of fluorophores suitable for STORM requires special buffer conditions, but with the appropriate fluorophore-buffer combination, the resolution can reach 5 nm (Dempsey et al., 2011). Efficient blinking of commonly used STORM fluorophores can be achieved in commercial mounting media, especially in Vectashield (Olivier et al., 2013). The introduction of Escherichia coli dihydrofolate reductase (eDHFR) as a protein tag of intracellular structures allows the labeling of eDHFR fusion proteins with fluorophore-conjugated trimethoprim primers used for live STORM applications (Wombacher et al., 2010).

STORM studies are very limited in plants and restricted to fixed probes. Such studies include the mapping of RNA polymerase II in interphase nuclei of Arabidopsis in fixed samples with Cy and AlexaFluor fluorophores (Schubert and Weisshart, 2015) and the visualization of cortical microtubules in fixed root cells immunolabeled with antibodies conjugated to the photoswitchable fluorophore AlexaFluor 647 at a resolution of approximately 50 nm (Dong et al., 2015).

STED

STED is a conceptually simple technique that projects two laser beams on the sample. The central Gaussian beam excites the sample while being surrounded by a doughnut-shaped eraser pulsed laser beam (Dyba and Hell, 2003). Superresolution in STED is achieved by the return in the ground state of nearly all excited fluorophores, except for those in the center of the excitation focus. Current STED beams include lasers at 592 nm, 660 nm (which corresponds to the second excitation maximum of chlorophyll), and 775 nm (https://www.leica-microsystems.com/fileadmin/downloads/Leica%20TCS%20SP8%20STED%203X/Brochures/Leica%20TCS%20SP8%20STED%203X-Brochure_EN.pdf). If compared with all other superresolution techniques, STED is based on scanning the sample with the combination of laser beams; in this way, it bears resemblance to CLSM. Since its development (Dyba and Hell, 2003; Dyba et al., 2003), STED has not found fertile ground in plant cell biology. There is only one study available (Kleine-Vehn et al., 2011). This might relate to the extreme output of the laser lines used, rendering STED quite phototoxic and unsuitable for mid- to long-term observations of living plant samples (for review, see Turkowyd et al., 2016). The original commercial STED designs used continuous-wave (CW) lasers (CW-STED), increasing the photon load and phototoxicity. An additional problem with CW lasers is that, for a short time, excited fluorophores outside the zero intensity center of the STED beam still emit photons before emission is suppressed. Such residual photons contribute to blurring of the resulting image when they are detected. This problem can be managed by so-called gated STED, detecting photons with a certain delay, in order to exclude the possibility that photons arising outside the center of the STED beam contribute to the image (Moffitt et al., 2011; Neupane et al., 2015; Castello et al., 2016). Another approach to improve the resolution of STED is the use of pulsed lasers. In this case, both lasers can be delivered as synchronous or asynchronous picosecond pulses, as in the original development of the method (Dyba and Hell, 2003). Moreover, image acquisition can be improved by resonant scanning, allowing very fast acquisition rates and high-rate volumetric imaging of actin filaments in living cotyledon epidermal cells at better resolution compared with CLSM (Fig. 5, F–I). A positive feature of STED is that, being a confocal microscopy-based technology, it can be used for superresolution in combination with quantitative imaging methods such as fluorescence correlation spectroscopy (FCS) either at the cell surface (Andrade et al., 2015) or in deeper parts of the cell (Lanzanò et al., 2017). Unlike SIM and PALM/STORM, which require postacquisition image processing to deliver the superresolved result, STED provides superresolution during acquisition.

Software-Based Superresolution for Live Cell Imaging

Software-based superresolution relies on optical fluctuations of fluorophores occurring under standard imaging conditions and the ability to record these fluctuations via single-photon-sensitive cameras. Such methods are described briefly here because they require a relatively cheap microscopic setup and represent an attractive option for the plant cell biologist.

One such example includes the acquisition of oversampled Z-stacks (i.e. smaller sample section thickness than what is defined by the Nyquist sampling rate; see https://svi.nl/NyquistRate) using a spinning disk microscope and subsequent 3D deconvolution based on commercially available software (Okamoto et al., 2012; Nakano, 2013). This approach was used in the study of cellulose synthase trafficking (Fujimoto et al., 2015).

Another software-based approach that can deliver superresolution results from any basic wide-field epifluorescence setup is called 3B (Cox et al., 2011). This method can be implemented as an ImageJ (https://imagej.nih.gov/ij/) plugin or a stand-alone platform (Cox et al., 2011; Rosten et al., 2013; both available from http://www.coxphysics.com/3b/). It is useful for time-lapse data sets of samples labeled with conventional fluorescent proteins. A big drawback of 3B is that it is extremely time consuming, which makes the processing of large data sets troublesome. However, cloud computing at affordable prices can overcome this drawback (Hu et al., 2013). A recently developed method called superresolution radial fluctuations (SRRF; Gustafsson et al., 2016; https://bitbucket.org/rhenriqueslab/nanoj-srrf/wiki/Home) is implemented as a plugin for Fiji image-analysis software (Schindelin et al., 2012; https://fiji.sc/) and can deliver superresolution information from both wide-field and confocal sources at a good temporal frame rate (1 s). In this respect, Andor has commercialized a much accelerated SRRF stream live superresolution workflow (http://www.andor.com/srrf-stream).

SOFI is another postacquisition software-processing procedure (Dertinger et al., 2009). Like 3B and SRRF, SOFI processes data sets acquired by standard wide-field epifluorescence platforms with no specification on the fluorophore but with the requirement that the fluorophore must exhibit at least two detectable fluorescent states (a dark one and a bright one; Dertinger et al., 2009). SOFI can deliver superresolution images at a lower sample SNR compared with default PALM/STORM approaches (Geissbuehler et al., 2011). Importantly, SOFI postacquisition processing can be combined with spinning disk imaging (Hosny et al., 2013).

LSFM

LSFM is a fast mesoscopic imaging method combining advantages of wide-field fluorescence microscopy and 4D volumetric imaging. It is based on sample illumination by a thin sheet of light (with controllable thickness in the range of micrometers). The light sheet can be generated in either a unilateral or, ideally, a bilateral fashion. Such an adjustable sheet of light enables optical sectioning of the sample and excitation of the fluorophores only within the thin illuminated volume. This illuminated sample volume is well adjusted to the focal plane of the detection objective, thus eliminating out-of-focus fluorophore excitation and light emission. Thus, LSFM prevents the photobleaching of fluorophores outside of the light sheet volume. Because the detection path is oriented orthogonally to the illumination, the system is capable of spherical aberration-free acquisition (Weber and Huisken, 2011). To allow optical sectioning, it is possible to either move the light sheet through the sample (Kumar et al., 2014) or, alternatively, to move the sample with respect to a stationary light sheet (Huisken et al., 2004; Chen et al., 2014; Reynaud et al., 2015; Kumar et al., 2016). With the high speed of sample movement through the light sheet, the fluorophore photobleaching as well as phototoxicity in the sample are reduced considerably. The above setup is supported by the currently existing LSFM platforms, as it allows very fast optical sectioning and rapid image acquisition. Therefore, the total effective excitation energy absorbed by the sample in LSFM can be several orders of magnitude lower in comparison with the epifluorescence wide-field or confocal laser scanning systems (Ichikawa et al., 2014; Stelzer, 2015). The sample mounted on a rotor can be rotated during multiangular acquisition, allowing precise postacquisition rendering for 4D (x, y, z, and t) image reconstruction. Although LSFM is a mesoscopic imaging method not even reaching resolution close to the diffraction limit, it became the most advanced approach for long-term developmental live cell imaging at the subcellular, cellular, tissue, organ, and whole-organism levels.

A great potential of LSFM in developmental studies is provided by its ability to maintain biological samples under almost natural conditions. The standard approach of animal sample preparation is full embedding and immobilization of the sample in transparent gel-like agarose (Kaufmann et al., 2012). Immobilization of plants in agarose ensures their stabilization in the vertical orientation. However, embedding of plant aerial parts is not compatible with proper physiological conditions, which are necessary for long-term imaging. Recently developed open systems for plant preparation and maintenance during LSFM long-term imaging (Ovečka et al., 2015; von Wangenheim et al., 2017) ensure that plants are maintained in the microscope vertically oriented (according to the gravity vector). Simultaneously, continuous root growth is supported by appropriate culture medium, and the aerial part grows freely in the air. Controllable illumination devices maintain desirable photoperiod protocols; thus, imaging experiments can be performed over a period of several days. Altogether, LSFM is a forefront method for long-term developmental imaging of plants in a natural vertical orientation maintaining their undisturbed growth in near-environmental conditions.

The most common type of LSFM devices is based on a classical selective-plane illumination microscopy (SPIM) principle with a horizontal setup for both illumination and detection objectives, where the sample is introduced to the field of view vertically from above. Microscopy modalities based on this hardware arrangement are either commercially available (Lightsheet Z.1 from Carl Zeiss) or can be constructed in place directly according to the OpenSPIM concept (http://openspim.org). The OpenSPIM concept was developed to adapt modular SPIM systems to the particular requirements of end users (Gualda et al., 2013; Pitrone et al., 2013). The concept is well supported by the community and provides a broad range of biological applications (Girstmair et al., 2016). By means of the vertical placement of the sample, both commercially available and custom-made LSFM/SPIM systems are suitable and already have been used successfully for long-term developmental imaging of plants. An alternative LSFM system based on the inclined optical arrangement, like dual-view inverted SPIM, might be designed for the imaging of animal living samples at subcellular resolution (Kumar et al., 2014, 2016). Such a system is optimal for the horizontal placement of rather thin and transparent samples mounted in a standard glass slide format (Kumar et al., 2014); therefore, it has no practical application on living plants so far.

Applications of LFSM in plant biology are dynamically evolving, while studies on growing Arabidopsis roots provided long-term observations of cell division progression in cells carrying the microtubule markers GFP-MBD (Maizel et al., 2011) and GFP-TUA5 (Ovečka et al., 2015; Fig. 6A). However, the spatial and temporal resolutions of LSFM also allow characterizing the redistributions of actin filaments in the phragmoplast during cytokinesis of Arabidopsis root cells carrying the actin marker YFP-FABD2-YFP (Fig. 6B) or tobacco BY-2 suspension cells carrying another actin marker, GFP-Lifeact (Fig. 6C). Owing to the high speed of imaging, LSFM can track the dynamics of more than one subcellular structure (Fig. 6D). Thus, Ovečka et al. (2015) showed live coimaging of actin filaments and microtubules in the cotyledon epidermal cells, while von Wangenheim et al. (2016) tracked cell fate by simultaneous imaging of plasma membrane and nuclear markers during lateral root formation. LSFM, unlike other live cell-imaging techniques, was able to successfully localize signaling protein such as mitogen-activated protein kinase 6 (MPK6) to the preprophase band of Arabidopsis root meristematic cells (Smékalová et al., 2014). One possible target of MPK6 is plant-specific EB1c. This protein is localized in the nucleus of nondividing cells. Long-term developmental imaging by LSFM revealed the correlation between nuclear size and EB1c content in diverse root tissues (epidermis, cortex, and endodermis) and in different developmental zones (meristematic, transition, and elongation zones) of the Arabidopsis root (Fig. 6, E–G; Novák et al., 2016). LSFM also allowed the precise characterization of cell division patterns during the formation and growth of Arabidopsis lateral roots (von Wangenheim et al., 2016). Another important advantage of LSFM for dynamic developmental studies of plants is deep imaging of nuclei (Novák et al., 2016) and endosomes (Berson et al., 2014) in inner root tissues and the undisturbed development of root hairs (Fig. 6, E–I). The good spatial and excellent temporal resolutions of LSFM also facilitate the simultaneous monitoring of mitotic microtubules in several tissue layers of the root meristem of the crop species Medicago sativa that is significantly thicker in comparison with Arabidopsis (Vyplelová et al., 2017). Recently, it was demonstrated that developmental imaging can be scaled up in existing commercial platforms by customized sample imaging chambers, allowing high-throughput imaging of developmental processes such as root growth (de Luis Balaguer et al., 2016).

Figure 6.

Figure 6.

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Advances of LSFM imaging of dynamic cellular and developmental processes in plants. A, Imaging of microtubules (MTs) in a time-course recording of a dividing Arabidopsis root epidermal cell as visualized using a GFP-TUA5 marker. B, Time-course imaging of actin in phragmoplast of a dividing Arabidopsis root epidermal cell (arrows) as visualized using a YFP-FABD2-YFP marker. C, Time-course imaging of actin in phragmoplast of a dividing BY-2 suspension cell as visualized using a GFP-Lifeact marker. D, Covisualization of actin using GFP-FABD2 and microtubules using mCherry-TUA5 in cotyledon stomata and pavement cells of Arabidopsis. E to G, Localization of EB1c in nuclei of Arabidopsis root as visualized using an EB1c-GFP marker. The high potential of LSFM for deep tissue imaging provides insight into EB1c-GFP nuclear localization in cells of different developmental zones (E), different root cell layers (F), as well as in individual cell files (G). H, Localization of the small Rab GTPase RabA1d in early endosomal-trans-Golgi network (EE-TGN) compartments accumulating at the tip of growing root hairs as visualized using a GFP-RabA1d marker. I, Imaging of late endosomes in Arabidopsis cortical and epidermal root cells as visualized using a YFP-RabF2a marker. Bars = 5 µm (A, B, and G–I) and 10 µm (C–F).

LSFM also can track physiological processes such as calcium fluxes by using ratiometric calcium indicators in the intact and freely moving nematode Caenorhabditis elegans (Ardiel et al., 2017) or during the neuronal activity of zebrafish larvae (Barykina et al., 2017). In the first case, fast exposure (2.5–4.5 ms) was translated to a 5-Hz volumetric acquisition frame rate, allowing imaging of the entire animal. Along with advancing volumetric image acquisition by paralleled illumination with more than one light sheet (Dean et al., 2017), such results exemplify the potential of LSFM to monitor fast physiological events with subcellular resolution in whole organisms, organs, or tissues.

The growth of sensitive cells such as root hairs may be compromised under standard conditions of sample preparation and imaging with conventional microscopes (Ovečka et al., 2005). By using LSFM, it is possible to monitor the growth of root hairs from their initiation to full differentiation but also to track fast-moving organelles such as endosomes and correlate their dynamics to the pattern of root hair elongation (Fig. 6, H and I; Berson et al., 2014). Moreover, LSFM was combined recently with Förster resonance energy transfer (FRET) to address the respiration rates of root hairs using a FRET sensor for MgATP production (De Col et al., 2017). A similar FRET-based sensor was used to follow cytosolic calcium oscillations during root (Costa et al., 2013) and root hair (Candeo et al., 2017) growth.

Of considerable importance for the implementation of LSFM in any laboratory is the fact that LSFM data sets are very big and routinely exceed terabyte sizes. This makes the postacquisition storage, processing (e.g. multiview registration and fusion), and analysis of such data sets quite demanding. At present, there are software solutions that efficiently address the problem of data access and postacquisition processing, including BigDataViewer (Pietzsch et al., 2015) and TeraFly (Bria et al., 2016).

Obviously, LSFM is not able to compete with superresolution methods providing nanotracking of structures and organelles at the subcellular level. However, LSFM significantly advanced the developmental imaging of plants, providing near-environmental conditions. In the near future, the functional combination of LSFM and superresolution concepts (Chen et al., 2014) might provide a new platform for very powerful spatiotemporal imaging of living plants.

SHORT UPDATE ON WIDE-FIELD AND CONFOCAL MICROSCOPY SYSTEMS

Research-grade wide-field epifluorescence microscopes and single-photon CLSM devices are in the standard setup of any cell biology research unit, and there have been numerous studies employing such microscopes. Especially CLSM has been in the forefront of qualitative and quantitative studies, such as the assessment of protein-protein interactions through FRET, recording of protein dynamics with fluorescence lifetime imaging (FLIM), and interactions with FLIM/FRET (Hoffmann et al., 2014; Garagounis et al., 2017; Lv et al., 2017), or the recording and the quantification of diffusion/mobility of single molecular species through FCS (Li et al., 2016). Especially in CLSM, speed, sensitivity, and the spectral range of image acquisition have been improved by resonant scanning approaches and the implementation of gallium arsenide phosphide alloy-based multispectral or hybrid detectors using sensitive avalanche photodiodes combined with the high dynamic range of PMTs (for review, see Jonkman and Brown, 2015).

For more specialized applications, platforms are expanding to include TIRF, particularly suitable for in vitro microscopic assays, variable-angle emission microscopy, with increased depth of excitation compared with TIRF, and finally, spinning disk microscopy, which is ideal for fast 3D time-lapse imaging of demanding events such as mitosis (Komis et al., 2017) but with reduced resolution capacity compared with CLSM (Wang et al., 2005). Detailed descriptions of these modalities are beyond the scope of this Update, and the reader is referred to relevant reviews (Henty-Ridilla et al., 2013; Shaw and Ehrhardt, 2013; Li et al., 2015).

Airyscan CLSM

A recent advancement of CLSM with improved resolution is Airyscan CLSM. The Airyscan detector has 32 subAiry (0.25 AU each) gallium arsenide phosphide alloy-based elements that are geometrically fitted to acquire the entire Airy pattern of emitted light and assign it to the correct position by software reconstruction (Weisshart, 2014; Huff et al., 2015). The averaged image from the detector elements can have improved xy resolution by a factor of 1.7 (resulting in a theoretical value of 140 nm). This corresponds to a CLSM device with the pinhole adjusted to approximately 0.2 AU but with much improved SNR (Weisshart, 2014). At the fast imaging mode, Airyscan frame rates approximate resonant scanning, being able to reach 27 frames per second but with compromise in resolution. Airyscan CLSM is a very recent advancement of microscopy technologies, and its applications have been limited. By taking advantage of high-speed acquisition at improved resolution, it was possible to capture microtubule-buckling events during the contraction of cardiomyocytes at high resolution (Robison et al., 2016). Very recently, Airyscan CLSM was used to capture mitotic spindle and phragmoplast dynamics in Arabidopsis ktn1-2 mutants devoid of the KATANIN1 protein (Komis et al., 2017).

CONCLUSION AND OUTLOOK

A survey of the available literature can show the importance of microscopy in plant live imaging. All the existing fluorescence-based microscopy methods have been applied to visualize different organelles and subcellular compartments. For quantitative approaches such as FRAP, FRET, FLIM, and FCS, the CLSM apparatus remains the workhorse, although other methods may be used instead (i.e. SIM can be combined with FRAP [Conduit et al., 2015] and STED can be used for FCS [Lanzanò et al., 2017]). With no doubt, recently developed advanced superresolution methods (such as SIM, PALM, STORM, and STED, software-based superresolution approaches, and Airyscan CLSM) and methods for long-term developmental imaging (such as LSFM and SPIM) have provided new dimensions and perspectives for better imaging of plant cells, especially in terms of significantly improved spatial and temporal resolution.

The field of microscopy is vigorously advancing toward hybrid methods, including Bessel or Airy beam illumination and lattice light-sheet microscopy. These were developed to compensate for speed, spatial resolution, and physiological imaging in just one microscope (Fahrbach and Rohrbach, 2012; Chen et al., 2014; Vettenburg et al., 2014). In the near future, they will possibly be applicable to the cumbersome plant specimens (Meinert et al., 2016). Such platforms will allow subcellular tracking of complex structures like the mitotic spindle in plant cells with unprecedented spatiotemporal resolution and with developmental connotations, similar to already published lattice light-sheet imaging of animal cells (Yamashita et al., 2015).

Integration of the precision of localization-based superresolution methods in plant cell imaging (including PALM and STORM) is expected to expand and help us uncover details of the molecular architecture and intermolecular associations within complex subcellular structures in the near future.

Acknowledgments

We thank Jinxing Lin (Key Laboratory of Photosynthesis and Molecular Environmental Physiology, Institute of Botany, Chinese Academy of Sciences) for Arabidopsis seeds carrying the proFlot1::FLOT1:GFP marker, Dr. Juraj Gregan (Department of Genetics, Faculty of Natural Sciences, Comenius University) for Schizosaccharomyces pombe strains carrying GFP-pcp1 and mCherry-atb2 spindle pole body and microtubule markers, Dr. Yoshihisa Oda (Department of Biological Sciences, Graduate School of Science, University of Tokyo) for the HTB-mRFP chromatin marker, Dr. Javier Pozueta (Instituto de Agrobiotecnología, Universidad Pública de Navarra) for Arabidopsis seeds carrying ADP-sugar pyrophosphatase fused with GFP, Dr. Yasuo Niwa (Laboratory of Plant Cell Technology, Graduate School of Nutritional and Environmental Sciences, University of Shizuoka) for Arabidopsis seeds carrying GFP-tagged mitochondria-targeting presequence of the N terminus of the γ-subunit of F1-ATPase, Dr. Jim Haseloff (Division of Cell Biology, Medical Research Council Laboratory of Molecular Biology, United Kingdom) for Arabidopsis seeds carrying GFP fused to the HDEL endoplasmic reticulum retention sequence mGFP5, Dr. Jiří Macas (Institute of Plant Molecular Biology, Czech Republic) for Arabidopsis seeds carrying GFP fused to a nuclear localization signal and to the complete coding region of GUS, Dr. Arun Sampathkumar (Max Planck Institute of Molecular Plant Physiology) for Arabidopsis seeds carrying the double markers GFP-FABD2 and mCherry-TUA5, and Dr. Andrei Smertenko (Institute of Biological Chemistry, Washington State University) for BY-2 cultures expressing GFP-Lifeact.

Footnotes

1

This work was supported by the Czech Science Foundation (GAČR), projects 15-19284 and 16-24313S.

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