We thank Dr. M. Rosenberg for her advice and help with anatomical sectioning. We also thank D. Eisler for his technical assistance. This research was supported by grants from FP7-PEOPLE-IRG-2008, Binational Agriculture research and Development (BARD; IS-4246-09), and Israel Science Foundation (ISF; 592/13).

1. Plant Growth
<ol>
	<li>Surface sterilize seeds.
	<ol>
		<li>Sterilize seeds (up to 30 mg) by wet or dry method. As an example, for dry sterilization method, add 1 ml glacial HCl 37% in 50 ml bleach, in a glass beaker in a closed desiccator for at least 4 hr and up to overnight. Perform this step in a chemical hood.</li>
	</ol>
	</li>
	<li>Spread the sterile seeds on plates with 0.8% plant agar in 0.5x Murashige &#38; Skoog (MS) medium. Wrap plates with Parafilm or porous tape and cover with aluminum foil.</li>
	<li>Store the plates at 4 &#176;C for 1 - 4 days to promote uniform germination.</li>
	<li>Transfer the plates to a growth chamber suitable for growing Arabidopsis seedlings. Place the plates vertically to maintain root growth on top of the agar medium and to prevent penetration.</li>
	<li>Transfer seedlings to fixative 7 days after germination (DAG).</li>
</ol>
2. Fixation
<ol>
	<li>Prepare 50 ml Richardson Solution by combining equal volumes of 1% Borax (buffer agent), 1% Methylene blue and 1% Azure (histological dyes) in double distilled water. Filter through a Syringe Driven 0.22 &#956;m PVDF Filter Unit before use.</li>
	<li>Prepare 50 ml of fixative: 1.25% Glutaraldehyde in 0.05 M Sodium Cacodylate (buffering agent). 
	Note: These materials are toxic and must be handled in a chemical hood.</li>
	<li>Add 100 &#181;l of Richardson solution (see 2.1) to the fixation solution (see 2.2) to obtain the final fixation solution.</li>
	<li>Mark each well of a 6-well plate with the corresponding sample name.</li>
	<li>Place 2 - 3 ml final fixation solution (see 2.3) into the wells, so that the seedlings, once transferred, will be fully covered in liquid.</li>
	<li>Carefully transfer, using forceps, up to 10 seedlings to each well. Seal the plates using Parafilm. Store at 4 &#176;C, in darkness, for at least one night, and up to one week.</li>
</ol>
3. Dehydration
<ol>
	<li>Dehydrate seedlings. Transfer them between wells of new 6-well plates, containing increasing concentrations of ethanol, as listed here: 10% ethanol for 15 min; 30% ethanol for 15 min; 50% ethanol for 15 min; 70% ethanol for 15 min; 85% ethanol for 15 min; 95% ethanol for 15 min; 95% ethanol at 4 &#176;C, overnight, at minimum. Handle seedlings by carefully grasping them by their cotyledons, preferable with plastic forceps.</li>
</ol>
4. Infiltration&#160;
<ol>
	<li>Prepare infiltration medium. Mix the contents of 1 bag of the historesin kit activator with 50 ml Basic resin liquid kit in a small glass bottle. Stir with magnet for 20 min. Infiltration medium may be kept at 4 &#176;C for a few weeks.</li>
	<li>Mark each well of a new 6-well plate with the corresponding sample name and fill with 2 - 3 ml of infiltration medium.</li>
	<li>Transfer the seedlings from the 95% ethanol solution to the infiltration medium using forceps. Make sure the seedlings are fully covered by the medium.</li>
	<li>Store at 4 &#176;C for at least 4 days, to ensure maximum penetration into the plant tissues.</li>
</ol>
5. Block Preparation
<ol>
	<li>Place small labels, pencil-marked with sample name, inside each well of embedding molds.</li>
	<li>Prepare embedding medium by adding 1 ml hardener to 15 ml infiltration medium (see 4.1). Do not attempt to prepare a volume smaller than 15 ml, to avoid polymerization issues.</li>
	<li>Fill half of each well in the mold with 200 &#181;l embedding medium and cover with an overhead film piece cut to the mold shape, but to a size 1 mm larger on each side. Incubate at room temperature for at least 2 hr, to allow polymerization. Do not exceed 5 hr.</li>
	<li>Prepare an additional batch of embedding medium (see 5.2) and keep on ice, to avoid polymerization while root tips are being arranged in the mold.</li>
	<li>Cut each root tip under a dissecting scope using scalpel and very carefully position it: vertically for transverse sections or horizontally for longitudinal section, at about 2 - 3 mm from the mold periphery. Fill the mold completely with blocking solution and cover with an overhead film. Note that the blocking solution will shrink slightly once it is polymerized.</li>
	<li>Keep at room temperature for 2 hr, at minimum.</li>
	<li>To dry the blocks, place in a box with dry silica gel and leave at room temperature overnight, at minimum.</li>
</ol>
6. Sectioning
<ol>
	<li>Prepare glass knives from glass rods with a knife maker. Use the glass knives for tissue sectioning. Cut the glass rod into a square and then cut the square diagonally to produce two knives, using a diamond scoring tool.</li>
	<li>Section blocks into 2 - 3 &#181;m width slices, using a sectioning machine. Place slices on a glass slide, on top of distilled water droplets and place the slide on a heat block set to low heat (50 - 60 &#176;C), until water evaporates.</li>
</ol>
7. Slide Preparation for Polarized Microscope
<ol>
	<li>Dilute Richardson solution (see 2.1) to 0.2% of the original solution and use 1 ml of it to cover the slices. Place on the heat block for 5 sec and then wash with distilled water.</li>
	<li>Dry the slide on a hot plate. Observe under a dissecting scope and mark the back of the slide with a marker, to indicate the position of the slices of interest, to facilitate their later identification under the microscope.</li>
	<li>Add 100 &#181;l of mounting medium and cover with a cover slip.</li>
</ol>
8. Image Acquisition
<ol>
	<li>Place the covered slide containing the root sections under a light microscope equipped with Polscope system suitable to obtain retardance information and a polarizer/interference optic filter (Figure 1 A-D).</li>
	<li>Select magnification that allows clear visualization of the entire sample (e.g., 40X in this study) and focus the microscope on an empty field that contains block section but no root section. Try to find a clean region with no blemishes. Use this region to set the background.</li>
	<li>Set the imaging system and software parameters: Set range to 17 nm. Apply &#34;Auto Exposure&#34; and obtain background image from the empty field (see 8.2). One background image is used per experiment.</li>
</ol>
9. Polscope Analysis of Cellulose Microfibril Orientation
<ol>
	<li>Analysis of longitudinal sections.
	<ol>
		<li>Place a slide containing longitudinal sections under the microscope and obtain a retardance image of it. Focus the microscope on a root section of interest. Provide a new file name.</li>
		<li>Capture an abrio image (image with retardation information) through the &#34;Live Abrio&#34; window. Be sure to capture a section that includes the cell wall, which can be identified by the stained cellulose running along the periphery of the cell, as shown in Figure 2C.</li>
	</ol>
	</li>
	<li>Image analysis.
	<ol>
		<li>Double-click on the relevant image in the image stack. Select &#34;Orientation Pseudo color&#34; in the &#34;Display Settings&#34; tab.</li>
		<li>To estimate the cellulose microfibril angle, match the corresponding shade of the cell wall to that in the color wheel. 
		NOTE: The color wheel in Figure 2C reflects the slow axis of light lying along the microfibril which indicates the direction along the long axis of the microfibrils. The angle of cellulose microfibrils is the angle indicated by the color and the long axis of the cell. For example, the elongating cell in the inset shows cyan color, tilted by about 90&#176; to the long axis of the cells.</li>
		<li>To quantify the average microfibril angle in the cell wall, use &#34;Retardance and Azimuth Stamp&#34; tool from the toolbar of the software. While selecting the tool, point and click on the region of interest in the obtained image. 
		NOTE: This will result in the display of the angles (azimuth) in degrees (Figure 2C).</li>
		<li>In an alternative open access polarized light image analysis software, lunch &#34;Pol-Analyzer&#34; plugin from the plugin menu. In the new opened window, select the &#34;Birefringence&#34; tab and open and then save the background file and the image file.</li>
		<li>In the image window, select &#34;Specs.&#34; and adjust the &#39;ROI pixel size&#39; (use 5 pixels for this study). In the &#34;Specs.&#34; window, select &#34;show values in Result table&#34;.</li>
		<li>Use the Polygon tool to mark the region of interest in the image window. Select &#34;Orientation Lines&#34; to obtain the orientation of the cellulose microfibril. 
		NOTE: This will result in the display of the angles (azimuth) in degrees. 
		NOTE: The average angle calculated by one of the two methods should be then normalized to the positioning of the root section along the slide. To correct for deviation from the horizontal positioning of the whole root section along the slide, quantify the average microfibril angle of the cell wall flanking the cell along its long axis (that is, perpendicular to the section and along the root growth axis, Figure 2C). This angle is 180&#176; when the root section is accurately positioned along the slide. For calculating the microfibril angle, subtract the average measured microfibril angle of the flanking cell wall from 180&#176;. Add the resulted difference to the average measured microfibril angle of the cell wall that lies flat on the slide.</li>
	</ol>
	</li>
</ol>
10. Polscope Analysis of Crystalline Cellulose Accumulation
<ol>
	<li>Use the transverse (cross) sections. Note that crystalline cellulose accumulation can only be compared between cells when their cellulose microfibrils are aligned at a similar angle (as determined in 9).</li>
	<li>Double-click on the relevant image in the image stack. Select &#34;Pseudo color&#34; in the &#34;Display Settings&#34; tab.</li>
	<li>To measure retardance. Zoom-in on the image. Select &#34;Region&#34; and mark all regions of interest. Move to the &#34;measurements&#34; tab and &#34;Copy to Clipboard&#34; all values.
	<ol>
		<li>Transfer the data to Excel and extract required information (mean region of retardance). Normalize the data. 
		NOTE: To account for differences in thickness between slices, express the values relative to a specific cell wall edge in the slide. To normalize, divide the average retardance corresponding to the cell wall of interest by the average retardance of a different cell wall in the same image. For example, the outer epidermal cell wall was normalized to the inner cortical cell wall in our study.</li>
	</ol>
	</li>
	<li>Repeat measurements for at least 3 sections per root, with a minimum of 3 roots per sample.</li>
</ol>

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The authors have no conflicts of interest.

Here, we present a method for determination of the accumulation of crystalline cellulose in the different tissues composing the Arabidopsis roots, while maintaining anatomical information. As such, it provides an additional step towards understanding growth processes in plants at a cellular resolution. This method can be also applied for the study of additional plant species and organs.
A number of points must be considered when applying the method. First, crystalline cellulose accumu...

The plant cell wall is a dynamic structure. Growing cells are surrounded by a primary cell wall, the organization of which allows cells to expand. Cells that cease to grow deposit a more rigid secondary wall that enhances the mechanical support of the plant. Both cell walls are composed of cellulose microfibrils embedded in a matrix of polysaccharides of different structures (e.g., hemicellulose and pectin) that vary across the different developmental stage and tissues1,2. Cellulose is synthesized as chains of (1,4)-&#946;-D-glucan that are tightly aligned to form the microfibril of a crystalline structure. Amorphous cellulose refers to the regions where the glucan chains are less ordered. The ratio between the crystalline and the amorphous domains is one parameter thought to affect the mechanical properties of the cell wall, by providing mechanical strength and viscoelastic characteristic, respectively3. Several methods have been developed to detect and quantify the two forms of cellulose arrangement, among them X-ray diffraction and cross polarization/magic angle spinning solid-state NMR4. X-ray diffraction can be used to determine the proportion of crystalline versus amorphous cellulose domains in the sample5. An alternative method uses fractionation of cell wall content into acid-insoluble and acid-soluble material, to distinguish between the crystalline and amorphous cellulose or other polymers, respectively. In this approach, incorporation of labeled glucose ([14C]Glucose) is used to quantify the cellulose6,7. These methods require large volumes of plant material for whole organ analyses, at best, and hence, are inadequately sensitive to tissue-specific variation in cell wall structure. Visualization of cellulose microfibrils at a cellular resolution can be achieved in live imaging studies combined with fluorescent dyes8,9, that can identify changes in the orientation of the cellulose microfibrils. However, these dyes are not used for quantification, they are not specific to crystalline cellulose and may interfere with the normal structure of the cell wall8. Polscope is an imaging technique that relies on the ability of crystalline cellulose to split light beams and retard part of the light10. Light retardation is strongest for microfibrils that lie perpendicularly to the direction of light propagation. For microfibrils with similar orientation, the higher the degree of crystallinity, the larger the light retardance11. Hence, polscope is used to study both the relative levels and orientation of the cellulose microfibrils.
Roots exhibit linear growth, during which cells originating at the stem cell niche, at the tip of the root, undergo a series of cell divisions, before they rapidly expand12. The cells comprising the root expand in a unidirectional (anisotropic) manner, as dictated by small molecule signaling hormones that impact the properties of the cell wall13. Differential responses to hormones, in time and space, provide a means of ensuring balanced organ growth14. Hence, high resolution analysis of cell wall structure can provide important information necessary to better understand the connection between cell type-specific responses to whole organ growth. Here, we report the implementation of polscope to study tissue-specific accumulation of crystalline cellulose in Arabidopsis roots, as observed in high quality anatomical sections. This method recently uncovered cell type-specific accumulation of crystalline cellulose in response to spatial perturbation of hormonal activity15.

<table><tbody><tr><td>Murashige &amp;amp; Skoog (MS)</td><td>Duchefa</td><td>M0221</td><td></td></tr><tr><td>Borax</td><td>LOBA CHEMIE&amp;nbsp;</td><td>6038</td><td></td></tr><tr><td>Methylene blue</td><td>Sigma</td><td>M-9140</td><td></td></tr><tr><td>Azure</td><td>Sigma</td><td>861065</td><td></td></tr><tr><td>Syringe Driven 0.22 mm PVDF Filter&amp;nbsp;</td><td>MILLEX-GV</td><td></td><td></td></tr><tr><td>Glutaraldehyde</td><td>EMS</td><td>16220</td><td></td></tr><tr><td>Sodium Cacodylate</td><td>Sigma</td><td>C-0250</td><td></td></tr><tr><td>Leica kit historesin&amp;nbsp;</td><td>Leica</td><td>7022 18 500</td><td></td></tr><tr><td>embedding molds</td><td>Agar Scientific</td><td>AGG3530</td><td></td></tr><tr><td>film 100 &amp;micro; P.P.C.&amp;nbsp;</td><td>www.Jolybar.com</td><td></td><td>overhead film</td></tr><tr><td>Knivemaker</td><td>LKB</td><td>7800</td><td></td></tr><tr><td>Ultratome III</td><td>LKB</td><td>8800</td><td>Ultratome</td></tr><tr><td>Shandon immumount&amp;nbsp;</td><td>Thermo</td><td>9990402</td><td>mounting medium</td></tr><tr><td>light microscope</td><td>Nikon</td><td>Eclipse 80i</td><td></td></tr><tr><td>Abrio imaging system</td><td>CRI</td><td>Abrio imaging system</td><td></td></tr><tr><td>Abrio V2.2 software</td><td>CRI</td><td>Abrio V2.2 software</td><td></td></tr><tr><td>Open access polarized light image analysis software &amp;nbsp;</td><td>OPS</td><td>OpenPolScope</td><td>&lt;a href=&quot;http://www.openpolscope.org/&quot;&gt;http://www.openpolscope.org/&lt;/a&gt;</td></tr></tbody></table>

high resolution quantification of crystalline cellulose accumulation in arabidopsis roots to monitor tissue-specific cell wall modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix containing polysaccharides that include cellulose microfibrils that form both crystalline structures and cellulose chains of amorphous organization. The orientation of the cellulose fibers and their concentrations dictate the mechanical properties of the cell. Several methods are used to determine the levels of crystalline cellulose, each bringing both advantages and limitations. Some can distinguish the proportion of crystalline regions within the total cellulose. However, they are limited to whole-organ analyses that are deficient in spatiotemporal information. Others relying on live imaging, are limited by the use of imprecise dyes. Here, we report a sensitive polarized light-based system for specific quantification of relative light retardance, representing crystalline cellulose accumulation in cross sections of Arabidopsis thaliana roots. In this method, the cellular resolution and anatomical data are maintained, enabling direct comparisons between the different tissues composing the growing root. This approach opens a new analytical dimension, shedding light on the link between cell wall composition, cellular behavior and whole-organ growth.

We study the impact of cell type-specific responses to brassinosteroids (BRs), using the Arabidopsis root as a model organ15-17. When the BR receptor BRI1 is targeted, in the background of bri1 mutant, to a subset of epidermal cells called non-hair cells (Figure 2A,B), it inhibits unidirectional cell expansion of neighboring cells, and whole-root growth15. Polscope analysis was performed to reveal the mechanism underlying this inhib...

Watch this Scientific Journal Video about High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications at JoVE.com

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Plant cells are surrounded by a cell wall, the composition of which determines their final size and shape. The cell wall is composed of a complex matrix ...

high-resolution-quantification-crystalline-cellulose-accumulation

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8.1K Views.  Technion-Israel Institute of Technology. Crystalline cellulose is an important constituent of the plant cell wall. However, its quantification at a cellular resolution is technically challenging. Here, we report the use of polarized light technology and root cross sections to obtain information of cell wall composition at a spatiotemporal resolution.

Video: High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part II: Carbohydrates

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st century. This has rekindled interest in the use of plant products as industrial raw materials for the production of liquid fuels for transportation2 and other products such as biocomposite materials6. Plant biomass remains one of the greatest untapped reserves on the planet4. It is mostly comprised of cell walls that are composed of energy rich polymers including cellulose, various hemicelluloses, and the polyphenol lignin5 and thus sometimes termed lignocellulosics. However, plant cell walls have evolved to be recalcitrant to degradation as walls contribute extensively to the strength and structural integrity of the entire plant. Despite its necessary rigidity, the cell wall is a highly dynamic entity that is metabolically active and plays crucial roles in numerous cell activities such as plant growth and differentiation5. Due to the various functions of walls, there is an immense structural diversity within the walls of different plant species and cell types within a single plant4. Hence, depending of what crop species, crop variety, or plant tissue is used for a biorefinery, the processing steps for depolymerisation by chemical/enzymatic processes and subsequent fermentation of the various sugars to liquid biofuels need to be adjusted and optimized. This fact underpins the need for a thorough characterization of plant biomass feedstocks. Here we describe a comprehensive analytical methodology that enables the determination of the composition of lignocellulosics and is amenable to a medium to high-throughput analysis (Figure 1). The method starts of with preparing destarched cell wall material. The resulting lignocellulosics are then split up to determine its monosaccharide composition of the hemicelluloses and other matrix polysaccharides1, and its content of crystalline cellulose7. The protocol for analyzing the lignin components in lignocellulosic biomass is discussed in Part I3.

Comprehensive Compositional Analysis of Plant Cell Walls Lignocellulosic biomass Part II: Carbohydrates

Plant biomass is a major carbon-neutral renewable resource that could be used for the production of biofuels. Plant biomass consists mainly of cell walls, a structurally complex composite material termed lignocellulosics. Here we describe a protocol for a comprehensive analysis of the content and composition of wall derived carbohydrates.

The need for renewable, carbon neutral, and sustainable raw materials for industry and society has become one of the most pressing issues for the 21st ...

Label-free in situ Imaging of Lignification in Plant Cell Walls

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find cost-effective and sustainable solutions has become a topical and critical task. Lignocellulosic biomass is poised to become the primary source of biomass for the conversion to liquid biofuels1-6. However, the recalcitrance of these plant cell wall materials to cost-effective and efficient degradation presents a major impediment for their use in the production of biofuels and chemicals4. In particular, lignin, a complex and irregular poly-phenylpropanoid heteropolymer, becomes problematic to the postharvest deconstruction of lignocellulosic biomass. For example in biomass conversion for biofuels, it inhibits saccharification in processes aimed at producing simple sugars for fermentation7. The effective use of plant biomass for industrial purposes is in fact largely dependent on the extent to which the plant cell wall is lignified. The removal of lignin is a costly and limiting factor8 and lignin has therefore become a key plant breeding and genetic engineering target in order to improve cell wall conversion.
Analytical tools that permit the accurate rapid characterization of lignification of plant cell walls become increasingly important for evaluating a large number of breeding populations. Extractive procedures for the isolation of native components such as lignin are inevitably destructive, bringing about significant chemical and structural modifications9-11. Analytical chemical in situ methods are thus invaluable tools for the compositional and structural characterization of lignocellulosic materials. Raman microscopy is a technique that relies on inelastic or Raman scattering of monochromatic light, like that from a laser, where the shift in energy of the laser photons is related to molecular vibrations and presents an intrinsic label-free molecular "fingerprint" of the sample. Raman microscopy can afford non-destructive and comparatively inexpensive measurements with minimal sample preparation, giving insights into chemical composition and molecular structure in a close to native state. Chemical imaging by confocal Raman microscopy has been previously used for the visualization of the spatial distribution of cellulose and lignin in wood cell walls12-14. Based on these earlier results, we have recently adopted this method to compare lignification in wild type and lignin-deficient transgenic Populus trichocarpa (black cottonwood) stem wood15. Analyzing the lignin Raman bands16,17 in the spectral region between 1,600 and 1,700 cm-1, lignin signal intensity and localization were mapped in situ. Our approach visualized differences in lignin content, localization, and chemical composition. Most recently, we demonstrated Raman imaging of cell wall polymers in Arabidopsis thaliana with lateral resolution that is sub-&mu;m18. Here, this method is presented affording visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species without staining or labeling of the tissues.

Label-free in situ Imaging of Lignification in Plant Cell Walls

A method based on confocal Raman microscopy is presented that affords label-free visualization of lignin in plant cell walls and comparison of lignification in different tissues, samples or species.

Meeting growing energy demands safely and efficiently is a pressing global challenge. Therefore, research into biofuels production that seeks to find ...

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through nanoindentations with an atomic force microscope (AFM) coupled with an optical inverted fluorescence microscope. The method consists of applying controlled forces to the sample while measuring its deformation, allowing quantifying parameters such as the apparent Young's modulus of cell walls at subcellular resolutions. It requires a careful mechanical immobilization of the sample and correct selection of indenters and indentation depths. Although it can be used only in external tissues, this method allows characterizing mechanical changes in plant cell walls during development and enables the correlation of these microscopic changes with the growth of an entire organ.

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

The atomic force microscopy indentation protocol offers the possibility to dissect the role of the physical properties of the cell wall of a particular cell of a tissue or organ during normal or constrained growth (i.e., under water deficit).

A method is described here to characterize the physical properties of the cell wall of epidermal cells of living Arabidopsis roots through ...

Developmental Biology

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 ...

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories can face is the development and attainment of these methods. Observation of phenotypes in a process of interest is a typical objective of research labs studying gene function and this is often achieved through image capture. A particular process that is amenable to observation using imaging approaches is the corrective growth of a seedling root that has been displaced from alignment with the gravity vector. Imaging platforms used to measure the root gravitropic response can be expensive, relatively low in throughput, and/or labor intensive. These issues have been addressed by developing a high-throughput image capture method using inexpensive, yet high-resolution, flatbed scanners. Using this method, images can be captured every few minutes at 4,800 dpi. The current setup enables collection of 216 individual responses per day. The image data collected is of ample quality for image analysis applications.

This protocol describes a process for rapid collection of images of Arabidopsis seedlings responding to a gravity stimulus using commercially-available flatbed scanners. The method allows for inexpensive, high-volume capture of high-resolution images amenable for downstream analysis algorithms.

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories ...

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) micro/nano-indentations, for a JPK&#160;AFM. Specifically, in this protocol we measure the apparent Young&#8217;s modulus of cell walls at subcellular resolutions across regions of up to 100 &#181;m&#160;x 100 &#181;m in floral meristems, hypocotyls, and roots. This requires careful preparation of the sample, the correct selection of micro-indenters and indentation depths. To account for cell wall properties only, measurements are performed in highly concentrated solutions of mannitol in order to plasmolyze the cells and thus remove the contribution of cell turgor pressure.
In contrast to other extant techniques, by using different indenters and indentation depths, this method allows simultaneous multiscale measurements, i.e. at subcellular resolutions and across hundreds of cells comprising a tissue. This means that it is now possible&#160;to spatially-temporally characterize the changes that take place in the mechanical properties of cell walls during development, enabling these changes to be correlated with growth and differentiation. This represents a key step to understand how coordinated microscopic cellular changes bring about macroscopic morphogenetic events.
However, several limitations remain: the method can only be used on fairly small samples (around 100 &#181;m in diameter) and only on external tissues; the method is sensitive to tissue topography; it measures only certain aspects of the tissue&#8217;s complex mechanical properties. The technique is being developed rapidly and it is likely that most of these limitations will be resolved in the near future.

We describe a method to map mechanical properties of plant tissues using an atomic force microscope (AFM). We focus on how to record mechanical changes that take place in cell walls during plant development at wide-field mesoscale, enabling these changes to be correlated with growth and morphogenesis.

We describe a recently developed method to measure mechanical properties of the surfaces of plant tissues using atomic force microscopy (AFM) ...

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Arabidopsis thaliana is a model organism commonly used to understand and manipulate various cellular processes in plants, and it has been used extensively in the study of secondary cell wall formation. Secondary cell wall deposition occurs after the primary cell wall is laid down, a process carried out exclusively by specialized cells such as those forming vessel and fiber tissues. Most secondary cell walls are composed of cellulose (40&#8211;50%), hemicellulose (25&#8211;30%), and lignin (20&#8211;30%). Several mutations affecting secondary cell wall biosynthesis have been isolated, and the corresponding mutants may or may not exhibit obvious biochemical composition changes or visual phenotypes since these mutations could be masked by compensatory responses. Staining procedures have historically been used to show differences on a cellular basis. These methods are exclusively visual means of analysis; nevertheless their role in rapid and critical analysis is of great importance. Congo red and calcofluor white are stains used to detect polysaccharides, whereas M&#228;ule and phloroglucinol are commonly used to determine differences in lignin, and toluidine blue O is used to differentially stain polysaccharides and lignin. The seemingly simple techniques of sectioning, staining, and imaging can be a challenge for beginners. Starting with sample preparation using the A. thaliana model, this study details the protocols of a variety of staining methodologies that can be easily implemented for observation of cell and tissue organization in secondary cell walls of plants.

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Plant cell wall composition varies between tissue types and can include lignin, cellulose, hemicelluloses, and pectin. Various staining techniques have been developed to visualize differences at the cell-type level. This paper is a compilation of commonly used cell wall staining techniques.

Arabidopsis thaliana is a model organism commonly used to understand and manipulate various cellular processes in plants, and it has been used extensively ...

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the adult plant. The female SMC (megaspore mother cell, MMC) differentiates in the ovule primordium and undergoes meiosis. The selected haploid megaspore then undergoes mitosis to form the multicellular female gametophyte, which will give rise to the gametes, the egg cell and central cell, together with accessory cells. The limited accessibility of the MMC, meiocyte and female gametophyte inside the ovule is technically challenging for cytological and cytogenetic analyses at single cell level. Particularly, direct or indirect immunodetection of cellular or nuclear epitopes is impaired by poor penetration of the reagents inside the plant cell and single-cell imaging is demised by the lack of optical clarity in whole-mount tissues.
Thus, we developed an efficient method to analyze the nuclear organization and chromatin modification at high resolution of single cell in whole-mount embedded Arabidopsis ovules. It is based on dissection and embedding of fixed ovules in a thin layer of acrylamide gel on a microscopic slide. The embedded ovules are subjected to chemical and enzymatic treatments aiming at improving tissue clarity and permeability to the immunostaining reagents. Those treatments preserve cellular and chromatin organization, DNA and protein epitopes. The samples can be used for different downstream cytological analyses, including chromatin immunostaining, fluorescence in situ hybridization (FISH), and DNA staining for heterochromatin analysis. Confocal laser scanning microscopy (CLSM) imaging, with high resolution, followed by 3D reconstruction allows for quantitative measurements at single-cell resolution.

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

We provide here an efficient and reliable protocol for immunostaining, Fluorescence in&#160;situ Hybridization, DNA staining followed by quantitative, high-resolution imaging in whole-mount Arabidopsis thaliana ovules. This method was successfully used to analyze chromatin modifications and nuclear architecture.

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the ...

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. Upon wounding of a leaf, an increase in cytosolic calcium ion concentration (Ca2+ signal) occurs at the wound site. This signal is rapidly transmitted to undamaged leaves, where defense responses are activated. Our recent research revealed that glutamate leaking from the wounded cells of the leaf into the apoplast around them serves as a wound signal. This glutamate activates glutamate receptor-like Ca2+ permeable channels, which then leads to long-distance Ca2+ signal propagation throughout the plant. The spatial and temporal characteristics of these events can be captured with real-time imaging of living plants expressing genetically encoded fluorescent biosensors. Here we introduce a plant-wide, real-time imaging method to monitor the dynamics of both the Ca2+ signals and changes in apoplastic glutamate that occur in response to wounding. This approach uses a wide-field fluorescence microscope and transgenic Arabidopsis plants expressing Green Fluorescent Protein (GFP)-based Ca2+ and glutamate biosensors. In addition, we present methodology to easily elicit wound-induced, glutamate-triggered rapid and long-distance Ca2+ signal propagation. This protocol can also be applied to studies on other plant stresses to help investigate how plant systemic signaling might be involved in their signaling and response networks.

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Extracellular glutamate-triggered systemic calcium signaling is critical for the induction of plant defense responses to mechanical wounding and herbivore attack in plants. This article describes a method to visualize the spatial and temporal dynamics of both these factors using Arabidopsis thaliana plants expressing calcium- and glutamate-sensitive fluorescent biosensors.

Plants respond to mechanical stresses such as wounding and herbivory by inducing defense responses both in the damaged and in the distal undamaged parts. ...

High-Resolution Imaging of Chlorophyll Biosynthesis in Arabidopsis Seedling

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly dynamic process of chlorophyll biosynthesis is triggered during the shift from the dark to the light in flowering plants. At the moment when etiolated seedlings are exposed to the first traces of sunlight, rapid (in order of seconds) conversion of protochlorophyllide into chlorophyllide is mediated by unique light-accepting protein complexes, leading via subsequent metabolic steps to the production of fully functional chlorophyll. Standard techniques for chlorophyll content analysis include pigment extraction from detached plant tissues, which does not apply to studying such fast processes. To investigate chlorophyll kinetics in vivo with high accuracy and spatiotemporal resolution in the first hours after light-induced de-etiolation, an instrument and protocol were developed. Here, we present a detailed procedure designed for statistically robust quantification of chlorophyll in the early stages of Arabidopsis de-etiolation.

Author Spotlight: Non-Invasive High-Resolution Measurement of Chlorophyll Synthesis During De-Etiolation

Here, we describe an advanced tool designed for chlorophyll biosynthesis monitoring during the early stages of Arabidopsis seedling de-etiolation. The novel methodology provides non-invasive real-time chlorophyll fluorescence imaging at high spatial and temporal resolution.

Chlorophyll biosynthesis is a hallmark of de-etiolation, one of the most dramatic stages in the plant life cycle. The tightly controlled and highly ...

High Resolution Quantification of Crystalline Cellulose Accumulation in Arabidopsis Roots to Monitor Tissue-specific Cell Wall Modifications

Summary

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Chapters in this video

Comprehensive Compositional Analysis of Plant Cell Walls (Lignocellulosic biomass) Part II: Carbohydrates

Label-free in situ Imaging of Lignification in Plant Cell Walls

Atomic Force Microscopy to Study the Physical Properties of Epidermal Cells of Live Arabidopsis Roots

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

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

AFM-based Mapping of the Elastic Properties of Cell Walls: at Tissue, Cellular, and Subcellular Resolutions

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

Wide-Field, Real-Time Imaging of Local and Systemic Wound Signals in Arabidopsis

Non-invasive Assay for Chlorophyll Biosynthesis Kinetics Determination during Early Stages of Arabidopsis De-etiolation