1. Plant growth
<ol>
	<li>Sow Arabidopsis seeds expressing microtubule binding domain fused with green fluorescent protein (MBD-GFP)10 on soil and keep in long day (16 h day /8 h night), 20 &#176;C/6 &#176;C conditions for 1 week for germination.</li>
	<li>After germination, transfer seedlings to new pots with sufficient growth space to allow robust vegetative growth. Keep plants in short day (8 h day /16 h night), 20 &#176;C/16 &#176;C conditions for 3-5 weeks.</li>
	<li>Transfer plants to long day (16 h day /8 h night), 20 &#176;C/16 &#176;C conditions and keep until plants bolt (2-3 weeks). Allow the inflorescence to grow up to 2-5 cm long.</li>
</ol>
2. Medium preparation
<ol>
	<li>Prepare the dissecting dishes by filling small Petri dishes (approximately 5.5 cm wide, 1.5 cm deep) to approximately half their depth with 2% agarose. The dissecting dishes could also be used for single time point imaging.</li>
	<li>Prepare the growth medium.
	<ol>
		<li>Prepare 50 mL of a 1,000x vitamin stock solution with 5 g of myo-inositol, 0.05 g of nicotinic acid, 0.05 g of pyridoxine hydrochloride, 0.5 g of thiamine hydrochloride, and 0.1 g of glycine.</li>
		<li>Prepare the growth medium, composed of &#189; Murashige and Skoog medium, 1% sucrose, 1.6% agar, 1x vitamins, pH = 5.8. Autoclave and allow it to cool.</li>
		<li>On a sterile, clean bench, sterilize hinged plastic boxes (approximately 5.2 cm x 5.2 cm x 3 cm) by immersing in 70% ethanol for 15 min.</li>
		<li>On a sterile, clean bench, once the medium is at bearable warmth, add N6-benzyladenine to a final concentration of 200 nM, and 1/1,000 PPM (plant preservative mixture) and mix well. Fill the sterile boxes to approximately half their depth with the medium.</li>
	</ol>
	</li>
</ol>
3. Dissection of the SAM
<ol>
	<li>Cut the inflorescence and remove the older flower buds with sharp forceps by peeling the flowers at the base of the peduncles until it is difficult to see them with the naked eye.</li>
	<li>Create a slit in the agarose in the dissecting dish with forceps and plant the inflorescence base into the thick agar. This gives good solid support for further fine dissection of younger flowers to expose the SAM.</li>
	<li>Remove the remaining flower buds by pushing them down with the forceps starting with the oldest and sequentially progressing to the younger stages under a dissecting microscope until the SAM is visible. The SAM is usually exposed when older flowers up to stage 6 to 7 are removed. Once the SAM is exposed, avoid dehydration of the sample by moving quickly to the next step.</li>
</ol>
4. Transfer and growth of cultured SAMs
<ol>
	<li>Plant the freshly dissected sample as detailed in section 1 into the growth medium in the rectangular plastic hinged culture box with the SAM just exposed above the medium surface. Add a few drops of sterile deionized water to the edges of the culture boxes and close the lid to maintain the humidity inside the box. Ensure that the added water does not cover the SAM.</li>
	<li>Close the lid and wrap the box with micropore tape. Place the growth box in long day or continuous day conditions at 22 &#176;C and grow for 12-24 h to allow the SAM to recover from the dissection procedure and adapt to the culture conditions.</li>
</ol>
5. Imaging of the SAM
<ol>
	<li>Fill the culture box containing the SAM with sterile deionized water to cover the sample. Check under the dissecting microscope and remove any air bubbles by forcefully spraying water directed at the sample with a 1 mL pipette.</li>
	<li>Place the culture box on an upright confocal microscope stage. Take care that the culture box does not contact the microscope objectives. Use a long distance 40x or 60x water dipping lens of numerical aperture 0.8-1 that is optimal for imaging without a cover glass.</li>
	<li>Lower the objective into the water and check for air bubbles formed on the objective&#39;s front lens. Remove any bubbles by lowering the stage and gently wiping the lens with an optical tissue and adding a small amount of water to the front lens of the objective with a Pasteur pipette before reimmersion into the water.</li>
	<li>Using the GFP filter and epi-illumination module of the confocal microscope, adjust the XY controller to locate the sample. Adjusting the position of the oculars, put the SAM directly under the light source and focus along the Z axis until the apex is located. 
	NOTE: Do not look directly at the ultraviolet light.</li>
	<li>Once the sample is found, illuminate it using a laser capable of exciting GFP (i.e., a 488 nm or 496 nm laser source). Adjust the optical zoom of the microscope so that the entire SAM and the stage 1 floral primordia are in the field of view. Further adjust the power of the laser output and gain settings to ensure optimal signal-to-noise ratio. 
	NOTE: The high intensity of the laser will result in photo bleaching of the sample. The under- and overexposure palette help ensure better adjustment of these settings.</li>
	<li>Allow the sample to settle down for 2-5 min. Acquire confocal Z stacks of the sample at 0.25 &#181;m-0.5 &#181;m Z slice intervals at a resolution of approximately 0.3 &#181;m pixel size. Ensure that the total imaging time required for acquiring Z stacks for one sample does not exceed 10 min from the time the sample is immersed into water until the completion of the acquisition.</li>
	<li>Immediately remove the water and transfer the culture box back to the growth chamber. Keeping the samples for a long time under water will influence the osmotic status of the cells.</li>
</ol>
6. Micromechanical perturbation of the SAM
<ol>
	<li>Set up the imaging conditions as described in section 5 and acquire preablation image stacks of the cortical microtubule organization.</li>
	<li>Decant the water in the culture dish. Perform the ablation with a clean syringe needle (0.4 mm x 20 mm).
	<ol>
		<li>Using a dissecting microscope, carefully hold the needle and slowly approach the SAM. 
		NOTE: Breath holding and handling the needle with a relaxed grip helps avoid shaking.</li>
		<li>Briefly contact the SAM dome with the needle tip to confirm that the ablation is done. Preferably perform the ablation on the periphery of the SAM, which allows visualization of the behavior and transitioning of unorganized cortical microtubules in the central region of the dome.</li>
	</ol>
	</li>
	<li>Refill the culture dish with sterile deionized water and add propidium idodide (10 &#181;g/mL). In addition to monitoring the GFP cortical microtubule signal, visualizing the propidium iodide using a separate channel can clearly mark regions of ablated cells. Propidium iodide is illuminated using the 561 nm or any other suitable laser with emission recorded between 600-650 nm.</li>
	<li>Acquire image stacks right after ablation (see section 6) and repeat the acquisition process every 2 h for a period of 6 h. Return the culture dish along with the sample into the incubation chamber after every time point. Ensure there is some water left on the culture media and wrap the culture dish with micropore tape to prevent desiccation.</li>
</ol>
7. Data visualization and quantification
<ol>
	<li>Generate surface projections using freely available software such as MORPHOGRAPHX11, FIJI12, or macro SurfCut13.</li>
	<li>Perform extraction of cortical microtubule anisotropy using FibrilTool14 macro in FIJI. 
	NOTE: Detailed protocols for software use can be obtained from the respective citations.</li>
</ol>

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The authors have nothing to disclose.

The assessment of mechanical signal transduction events is crucial to identify molecular regulators involved in the mechano-perception and transduction pathways. The protocol described here provides a quantitative view of such events by using the cortical microtubule response as a readout for such a process in Arabidopsis SAMs. The procedure described here is routinely used in several studies in various tissue types16,17,18</...

Plant cells are surrounded by an extracellular matrix of polysaccharides and glycoproteins that mechanically resembles a fiber reinforced composite material capable of dynamically changing its mechanical properties1. Growth in plant cells is driven by the uptake of water into the cell, which results in a concomitant buildup of tensile forces on the cell wall. In response to such forces, modifications to the physical state of the cell wall allows for cell expansion. Cells with primary walls are capable of undergoing rapid growth compared to secondary cell wall containing cells mainly due to differences in the chemical composition of the polysaccharides within. Primary wall cells are composed of cellulose, hemicellulose, and pectin in addition to glycoproteins, and lack lignin, a component that is present in the secondary cell wall2. Cellulose, a glucose polymer linked via &#946;-1,4 bonds, is the major component of the cell walls. It is organized into fibrillar structures that are capable of withstanding high tensile forces experienced during cell growth3. In addition to withstanding tensile forces, mechanical reinforcement along a preferential direction results in turgor-driven expansion along an axis perpendicular to the net orientation of the cellulose microfibril. The organization of the cellulose microfibrils is influenced by the cortical microtubule cytoskeleton, as they guide the directional movement of the cellulose-synthesizing complexes located at the plasma membrane4. Therefore, monitoring cortical microtubule organization using a microtubule-associated protein or tubulin fused with a fluorescent molecule serves as a proxy for the observation of overlying patterns of cellulose in plant cells.
The patterning of the cortical microtubule cytoskeleton is under the control of cell and tissue morphology derived mechanical forces. Cortical microtubule organization does not have any preferential organization over time in cells located at the apex of the SAM, whereas cells in the periphery and the boundary between the SAM and the emerging organ have a stable, highly organized supracellular array of cortical microtubules5. Several approaches have been developed to physically perturb the mechanical status of the cells. Changes to osmotic status, as well as treatment with pharmacological and enzymatic compounds that influence the stiffness of the cell wall can result in subsequent changes in the tensile forces experienced by the cell6,7. The use of contraptions that allow for the gradual increase in compressive forces experienced by tissues is another alternative8. Application of centrifugal forces has also been shown to influence the mechanical forces without any physical contact with the cells9. However, the most widely used means of changing directional forces in a group of cells take advantage of the fact that all epidermal cells are under tension and physical ablation of cells will eliminate turgor pressure locally as well as disrupting cell-to-cell adhesion, thereby modifying the tensile forces experienced by the neighboring cells. This is performed either by targeting a high-powered pulsed ultraviolet laser or by means of a fine needle.
Here we elaborate on the process of imaging and assessing cortical microtubule behavior for mechanical perturbation at the SAM.

<table><tbody><tr><td>FibrilTool</td><td></td><td></td><td>Boudaoud, A. &lt;em&gt;et al&lt;/em&gt;., Nat Protoc. 2014</td></tr><tr><td>FIJI</td><td></td><td></td><td>Schindelin, J. &lt;em&gt;et al&lt;/em&gt;., Nat Methods. 2012</td></tr><tr><td>glycine</td><td>Merck</td><td>1.04201.1000</td><td></td></tr><tr><td>Leica SP8 confocal microscope</td><td>Leica</td><td>DM6000 CS</td><td></td></tr><tr><td>MAP4-GFP</td><td></td><td></td><td>Marc, J. &lt;em&gt;et al&lt;/em&gt;., Plant Cell 1998</td></tr><tr><td>micropore tape</td><td>Leukopor</td><td>02482-00</td><td></td></tr><tr><td>MorphographX</td><td></td><td></td><td>Strauss, S. &lt;em&gt;et al&lt;/em&gt;., Methods Mol Biol. 2019</td></tr><tr><td>myo-inositol</td><td>Sigma</td><td>I5125</td><td></td></tr><tr><td>N6-benzyladenine</td><td>Sigma</td><td>B3408</td><td></td></tr><tr><td>nicotinic acid</td><td>Sigma</td><td>N4126</td><td></td></tr><tr><td>plastic hinged box</td><td>Electron microscopy sciences</td><td>64312</td><td></td></tr><tr><td>PPM (Plant Preservative Mixture)</td><td>Plant Cell Technology</td><td>PPM</td><td></td></tr><tr><td>Propidium iodide</td><td>Sigma</td><td>P4864</td><td></td></tr><tr><td>pyridoxine hydrochloride</td><td>Sigma</td><td>P9755</td><td></td></tr><tr><td>SURFCUT</td><td></td><td></td><td>Erguvan, O. &lt;em&gt;et al&lt;/em&gt;., BMC Biol. 2019</td></tr><tr><td>thiamine hydrochloride</td><td>Sigma</td><td>T4625</td><td></td></tr></tbody></table>

live cell imaging of microtubule cytoskeleton and micromechanical manipulation of the arabidopsis shoot apical meristem

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in molecular and imaging technologies allowed us to gain insights into how biochemical signals influence morphogenetic events. However, it is increasingly evident that apart from biochemical signals, mechanical cues also impact several aspects of cell and tissue growth. The Arabidopsis shoot apical meristem (SAM) is a dome-shaped structure responsible for the generation of all aboveground organs. The organization of the cortical microtubule cytoskeleton that mediates apoplastic cellulose deposition in plant cells is spatially distinct. Visualization and quantitative assessment of patterns of cortical microtubules are necessary for understanding the biophysical nature of cells at the SAM, as cellulose is the stiffest component of the plant cell wall. The stereotypical form of cortical microtubule organization is also a consequence of tissue-wide physical forces existing at the SAM. Perturbation of these physical forces and subsequent monitoring of cortical microtubule organization allows for the identification of candidate proteins involved in mediating mechano-perception and transduction. Here we describe a protocol that helps investigate such processes.

Figure 1 shows typical projection images obtained from MBD-GFP lines with cells at the center of the dome containing disorganized cortical microtubules, and cells at the periphery having a circumferential distribution (Figure 1A,B), whereas the boundary domain cells contain cortical microtubules aligned parallel to the cell&#39;s long axis. These observations show differences in the spatial distribution of cortic...

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Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Here we describe a protocol for live cell imaging of the cortical microtubule cytoskeleton at the shoot apical meristem and monitoring its response to changes in physical forces.

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in ...

live-cell-imaging-microtubule-cytoskeleton-micromechanical

Research

JoVE Journal

Developmental Biology

5.2K Views.  Max Planck Institute of Molecular Plant Physiology. Here we describe a protocol for live cell imaging of the cortical microtubule cytoskeleton at the shoot apical meristem and monitoring its response to changes in physical forces.

Video: Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Live Confocal Imaging of Developing Arabidopsis Flowers

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. Recent advances in confocal microscopy, combined with the development of numerous fluorophores, have overcome these issues and opened the possibility to study the expression of several genes simultaneously, with a good cellular resolution, in live samples. Live confocal imaging provides plant biologists with a powerful tool to study development, and has been extensively used to study root growth and the formation of lateral organs on the flanks of the shoot apical meristem. However, it has not been widely applied to the study of flower development, in part due to challenges that are specific to imaging flowers, such as the sepals that grow over the flower meristem, and filter out the fluorescence from underlying tissues. Here, we present a detailed protocol to perform live confocal imaging on live, developing Arabidopsis flower buds, using either an upright or an inverted microscope.

Live confocal imaging provides biologists with a powerful tool to study development. Here, we present a detailed protocol for the live confocal imaging of developing Arabidopsis flowers.

The study of plant growth and development has long relied on experimental techniques using dead, fixed tissues and lacking proper cellular resolution. ...

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the flexible and efficient GreenGate cloning system with the proven and benchmarked LhGR system (here termed GR-LhG4) for the inducible expression, we have generated a set of transgenic Arabidopsis lines that can drive the expression of an effector cassette in a range of specific cell types in the three main plant meristems. To this end, we chose the previously developed GR-LhG4 system based on a chimeric transcription factor and a cognate pOp-type promoter ensuring tight control over a wide range of expression levels. In addition, to visualize the expression domain where the synthetic transcription factor is active, an ER-localized mTurquoise2 fluorescent reporter under control of the pOp4 or pOp6 promoter is encoded in driver lines. Here, we describe the steps necessary to generate a driver or effector line and demonstrate how cell type specific expression can be induced and followed in the shoot apical meristem, the root apical meristem and the cambium of Arabidopsis. By using several or all driver lines, the context specific effect of expressing one or multiple factors (effectors) under control of the synthetic pOp promoter can be assessed rapidly, for example in F1 plants of a cross between one effector and multiple driver lines. This approach is exemplified by the ectopic expression of VND7, a NAC transcription factor capable of inducing ectopic secondary cell wall deposition in a cell autonomous manner.

Inducible, Cell Type-Specific Expression in Arabidopsis thaliana Through LhGR-Mediated Trans-Activation

Here, we describe the generation and application of a set of transgenic Arabidopsis thaliana lines enabling inducible, tissue-specific expression in the three main meristems, the shoot apical meristem, the root apical meristem, and the cambium.

Inducible, tissue-specific expression is an important and powerful tool to study the spatio-temporal dynamics of genetic perturbation. Combining the ...

Genetics

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; for example, how the zygote polarizes to establish the body axis and how the embryo specifies various cell fates during organ formation. This manuscript describes an in vitro ovule culture method to perform live-cell imaging of developing zygotes and embryos of Arabidopsis thaliana. The optimized cultivation medium allows zygotes or early embryos to grow into fertile plants. By combining it with a poly(dimethylsiloxane) (PDMS) micropillar array device, the ovule is held in the liquid medium in the same position. This fixation is crucial to observe the same ovule under a microscope for several days from the zygotic division to the late embryo stage. The resulting live-cell imaging can be used to monitor the real-time dynamics of zygote polarization, such as nuclear migration and cytoskeleton rearrangement, and also the cell division timing and cell fate specification during embryo patterning. Furthermore, this ovule cultivation system can be combined with inhibitor treatments to analyze the effects of various factors on embryo development, and with optical manipulations such as laser disruption to examine the role of cell-cell communication.

In Vitro Ovule Cultivation for Live-cell Imaging of Zygote Polarization and Embryo Patterning in Arabidopsis thaliana

This manuscript describes an in vitro ovule cultivation method that enables live-cell imaging of Arabidopsis zygotes and embryos. This method is utilized to visualize the intracellular dynamics during zygote polarization and the cell fate specification in developing embryos.

In most flowering plants, the zygote and embryo are hidden deep in the mother tissue, and thus it has long been a mystery of how they develop dynamically; ...

Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic development. The activity and morphology of SAMs determine important agronomic traits, such as shoot architecture, size and number of reproductive organs, and most importantly, grain yield. Here, we provide a detailed protocol for analyzing both the surface morphology and the internal cellular structure of the living SAMs from different species through laser scanning confocal microscope. The whole procedure from the sample preparation to the acquisition of high resolution three-dimensional (3D) images can be accomplished within as short as 20 minutes. We demonstrate that this protocol is highly efficient for studying not only the inflorescence SAMs of the model species but also the vegetative meristems from different crops, providing a simple but powerful tool to study the organization and development of meristems across different plant species.

This protocol presents how to live image and analyze the shoot apical meristems from different plant species using laser scanning confocal microscopy.

The shoot apical meristem (SAM) functions as a conserved stem cell reservoir and it generates almost all aboveground tissues during the postembryonic ...

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

Bioengineering

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In general, for observations over a long period, the technique requires transformation of a fluorescent protein; however, for most systems, genetic transformation is either time-consuming or technically unavailable. This manuscript presents a protocol for 3-D time-lapse imaging of cell wall dynamics over a 3 day period using calcofluor dye (which stains cellulose in the plant cell wall), developed in the moss Physcomitrium patens. The calcofluor dye signal from the cell wall is stable and can last for 1 week without obvious decay. Using this method, it has been shown that the detachment of cells in ggb mutants (in which the protein geranylgeranyltransferase-I beta subunit is knocked out) is caused by unregulated cell expansion and cell wall integrity defects. Moreover, the patterns of calcofluor staining change over time; less intensely stained regions correlate with the future cell expansion/branching sites in the wild type. This method can be applied to many other systems that contain cell walls and that can be stained by calcofluor.

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

This manuscript presents a detailed protocol to image the 3-D cell wall dynamics of living moss tissue, allowing the visualization of the detachment of cell walls in ggb mutants and thickening cell wall patterns in the wild type during development over a long period.

Time-lapse imaging with fluorescence microscopy allows observation of the dynamic changes of growth and development at cellular and subcellular levels. In ...

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue patterning. During animal development, key cell proliferation and patterning events occur very quickly. For instance, in Caenorhabditis elegans all cell divisions required for the larval body plan are completed within six hours after fertilization, with seven mitotic cycles1; the sixteen or more mitoses of Drosophila embryogenesis occur in less than 24 hr2. In contrast, cell divisions during plant development are slow, typically on the order of a day 3,4,5 . This imposes a unique challenge and a need for long-term live imaging for documenting dynamic behaviors of cell division and differentiation events during plant organogenesis. Arabidopsis epidermis is an excellent model system for investigating signaling, cell fate, and development in plants. In the cotyledon, this tissue consists of air- and water-resistant pavement cells interspersed with evenly distributed stomata, valves that open and close to control gas exchange and water loss. Proper spacing of these stomata is critical to their function, and their development follows a sequence of asymmetric division and cell differentiation steps to produce the organized epidermis (Fig. 1).
This protocol allows observation of cells and proteins in the epidermis over several days of development. This time frame enables precise documentation of stem-cell divisions and differentiation of epidermal cells, including stomata and epidermal pavement cells. Fluorescent proteins can be fused to proteins of interest to assess their dynamics during cell division and differentiation processes. This technique allows us to understand the localization of a novel protein, POLAR6, during the proliferation stage of stomatal-lineage cells in the Arabidopsis cotyledon epidermis, where it is expressed in cells preceding asymmetric division events and moves to a characteristic area of the cell cortex shortly before division occurs. Images can be registered and streamlined video easily produced using public domain software to visualize dynamic protein localization and cell types as they change over time.

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

We describe a protocol using chamber slides and media to immobilize plant cotyledons for confocal imaging of the epidermis over several days of development, documenting stomatal differentiation. Fluorophore-tagged proteins can be tracked dynamically by expression and subcellular localization, increasing understanding of their possible roles during cell division and cell-type differentiation.

Imaging in vivo dynamics of cellular behavior throughout a developmental sequence can be a powerful technique for understanding the mechanics of tissue ...

Biology

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ binding proteins that initiate Ca2+ signaling cascades. However, we still know little about how stimulus specific Ca2+ signals are generated. The specificity of a Ca2+ signal may be attributed to the sophisticated regulation of the activities of Ca2+ channels and/or transporters in response to a given stimulus. To identify these cellular components and understand their functions, it is crucial to use systems that allow a sensitive and robust recording of Ca2+ signals at both the tissue and cellular levels. Genetically encoded Ca2+ indicators that are targeted to different cellular compartments have provided a platform for live cell confocal imaging of cellular Ca2+ signals. Here we describe instructions for the use of two Ca2+ detection systems: aequorin based FAS (film adhesive seedlings) luminescence Ca2+ imaging and case12 based live cell confocal fluorescence Ca2+ imaging. Luminescence imaging using the FAS system provides a simple, robust and sensitive detection of spatial and temporal Ca2+ signals at the tissue level, while live cell confocal imaging using Case12 provides simultaneous detection of cytosolic and nuclear Ca2+ signals at a high resolution.

Measuring Spatial and Temporal Ca2+ Signals in Arabidopsis Plants

Ca2+ signaling regulates diverse biological processes in plants. Here we present approaches for monitoring abiotic stress induced spatial and temporal Ca2+ signals in Arabidopsis cells and tissues using the genetically encoded Ca2+ indicators Aequorin or Case12.

Developmental and environmental cues induce Ca2+ fluctuations in plant cells. Stimulus-specific spatial-temporal Ca2+ patterns are sensed by cellular Ca2+ ...

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and cytoskeletal rearrangement. Real-time and high-resolution image analysis of such intracellular events will increase the understanding of plant cell biology at the molecular level. Variable angle epifluorescence microscopy (VAEM) is an emerging technique that provides high-quality, time-lapse images of fluorescently-labeled proteins on the plant cell surface. In this article, practical procedures are described for VAEM specimen preparation, adjustment of the VAEM optical system, movie capturing and image analysis. As an example of VAEM observation, representative results are presented on the dynamics of PATROL1. This is a protein essential for stomatal movement, thought to be involved in proton pump delivery to plasma membranes in the stomatal complex of Arabidopsis thaliana. VAEM real-time observation of guard cells and subsidiary cells in A. thaliana cotyledons showed that fluorescently-tagged PATROL1 appeared as dot-like structures on plasma membranes for several seconds and then disappeared. Kymograph analysis of VAEM movie data determined the time distribution of the presence (termed &#8216;residence time&#8217;) of the dot-like structures. The use of VAEM is discussed in the context of this example.

The goal of this protocol is to demonstrate how to monitor fluorescently-tagged protein dynamics on plant cell surfaces with variable-angle epifluorescence microscopy, showing blinking dots of GFP-tagged PATROL1, a membrane trafficking protein, in the cell cortex of the stomatal complex in Arabidopsis thaliana.

A plant&#8217;s cell surface is its interface for perceiving environmental cues; it responds with cell biological changes such as membrane trafficking and ...

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

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Confocal Live Imaging of Shoot Apical Meristems from Different Plant Species

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

3-D Time-Lapse Imaging of Cell Wall Dynamics Using Calcofluor in the Moss Physcomitrium patens

Long-term, High-resolution Confocal Time Lapse Imaging of Arabidopsis Cotyledon Epidermis during Germination

Measuring Spatial and Temporal Ca²⁺ Signals in Arabidopsis Plants

Real-time Imaging of Plant Cell Surface Dynamics with Variable-angle Epifluorescence Microscopy