We are thankful to Sabin Russell for editing assistance. This work was part of the DOE Joint BioEnergy Institute (http://www.jbei.org) supported by the U. S. Department of Energy, Office of Science, Office of Biological and Environmental Research, through contract DE-AC02-05CH11231 between Lawrence Berkeley National Laboratory and the U.S. Department of Energy.

1. Stem Embedding
<ol>
	<li>Make a 7% agarose solution in water (7 g of electrophoresis-grade agarose in 100 ml distilled water). Dissolve the agarose by autoclaving for 20 min or by microwaving for 20 min at lowest intensity (e.g., 10% intensity of a 1,250 watt&#160;microwave).</li>
	<li>Prepare a homemade mold for embedding the stems using plastic vials.
	<ol>
		<li>Using a razor blade, cut off the conical bottom of a 2 ml screw-cap microcentrifuge tube (Part A). With a syringe needle, puncture a hole in the tube cap slightly larger than the diameter of the stem to be embedded. Next, cut 0.5 cm off the bottom of a 0.6 ml microcentrifuge tube (Part B).</li>
		<li>Add the cut-off 0.5 cm portion of the 0.6 ml microcentrifuge tube into the precut 2 ml microcentrifuge tube. Seal the two parts using Parafilm (Figure 1). NOTE: The bottom of the tube is cut to facilitate the removal of the embedded sample after the agarose has solidified. The hole in the cap will aid in holding the stem steady when the stem is embedded in agarose. The mold will serve to hold the agarose and hold the stem straight when embedding. The Parafilm will hold the two parts, A and B, until the agarose is set and the stem is embedded.</li>
		<li>Using a razor blade, cut a stem section from the area of interest, (e.g. middle of the first stem internode). NOTE: Ideally, the stem should be cut just before embedding to avoid destruction by dehydration. Alternatively, the stem can be temporarily stored in a plate containing a filter paper wetted with water. This plate can be placed on ice until ready to be embedded to prevent tissue dehydration and deformation. Storing the stem for periods longer than 30 min without high moisture will cause the stem to dry out.</li>
	</ol>
	</li>
	<li>Melt the agarose in the microwave at low intensity. CAUTION: The bottle containing agarose may be hot. Use a mitt to pick up the bottle. Let the melted agarose stand at room temperature (between 20 &#176;C and 25 &#176;C) until it cools to about 50 &#176;C.</li>
	<li>Slowly pipette 5 ml of the agarose into the premade plastic mold to fill the vial. NOTE: Make sure there are no air bubbles in the molds. Air bubbles will cause gaps in the agarose mold and will not completely cover the stem sample, which will eventually lead to uneven cutting of the sample sections.
	<ol>
		<li>Let the agarose cool down for 30 to 60 sec to become semisolid. Test with a small toothpick, making sure that it can penetrate and stand up, but not float, in the agarose. NOTE: If the stem is placed when the agarose is still hot it will damage the sample by shriveling the stem, destroying the cell morphology.</li>
		<li>Place the stem in this agarose-filled vial. Make sure that the stem stays straight. Place the perforated cap in such a way that the tip of the stem is held by the cap. Do not turn the screw-on cap. Sometime more than one stem (of a single kind) can be embedded in a single vial; but in that particular case, do not use the cap. NOTE: If the screw-on cap is turned, the stem will get twisted.</li>
	</ol>
	</li>
	<li>Leave the vial at room temperature or at 4 &#176;C for 10-30 min, until it solidifies.</li>
	<li>Remove the mold by gently sliding it out of the tube. Open the Parafilm. Push the bottom of Part B of the mold gently with thumb to release the solidified agarose from Part A onto a glass microscope slide.</li>
	<li>Cut the agarose embedded stem into three approximately equal pieces of 1.2 cm in length. Cut out the part of the stem that was not in the agarose and discard it.
	<ol>
		<li>Store these pieces of agarose with the embedded stems in airtight 2 ml microcentrifuge tubes in a dark box at 4 &#176;C until ready to section. Alternatively, store the tubes at 4 &#176;C for a maximum of one week. NOTE: Storage is possible, but the experimenter needs to be aware that the embedded stems are still alive and, depending of the experiment, artifacts could potentially be introduced.</li>
	</ol>
	</li>
</ol>
2. Stem Sectioning
<ol>
	<li>CAUTION: Read the vibratome manual carefully and follow all safety instructions.</li>
	<li>Make sure the specimen disc is clean and completely dry (free of any solid or liquid). Clean the specimen disc with a razor blade to remove any residual glue from prior experiments and wash with water. Then dry with a paper towel. NOTE: This will ensure that the adhesive works well and binds to the agarose specimen. If this step is not done properly, it is possible that the specimen may not adhere to the disc and will later cause the specimen to fall off the disc during sectioning.
	<ol>
		<li>Use soft paper wipes to remove any moisture from the agarose block containing the specimens.</li>
		<li>Place a small drop of tissue adhesive on the specimen disc. Streak out the adhesive to cover the area in the middle of the plate using the tip of the adhesive bottle. Quickly place the agarose block so that the specimens are either perpendicular to or in parallel with the plate for transversal or longitudinal cross-sections, respectively. Allow the adhesive to fix the sample to the specimen disc at room temperature or at 4 &#176;C for 10-30 min. NOTE: This is a time-sensitive step.</li>
	</ol>
	</li>
	<li>Fix the specimen disc in place on the buffer tray or trough. The block/s of agarose with the sections should be in parallel with the razor blade. Fill the buffer tray with distilled water at room temperature until the samples are completely submerged.</li>
	<li>Cut a razor blade in half and then trim the ends with a sturdy scissors so that the blade stays completely flat. Attach one-half of the precut razor blade onto the knife holder.
	<ol>
		<li>Set the angle on the knife holder to 84&#176;, the speed to 0.90 mm/sec (position 8 on the vibratome) and frequency to 50 Hz (position 5 on the vibratome). Cut sections of 100 &#181;m thickness. Use continuous mode. NOTE: This thickness was selected to assure that the tissue can withstand the various acid and base treatments used in some of the staining protocols. Set the window for sectioning and allow 15-20 min on a continuous mode for the vibratome to section an agarose block of approximately 1.2 cm.</li>
	</ol>
	</li>
	<li>Collect the sections using a disposable plastic pipette during sectioning in the buffer tray. Alternatively, the sections can be transferred from the buffer tray into a glass beaker and then to a clear Petri plate. Transfer few sections to the 2.0 ml microcentrifuge tubes. Note: The specimens are collected from a Petri plate because it is easier to see the sections on a clear background, and because the buffer tray can be primed for the next specimen.</li>
	<li>The sections can be stored in a 50 ml tube or collected and stored in 1.5 to 2 ml microcentrifuge tubes at 4 &#176;C for 30 to 60 min. NOTE: Storing the sections for longer periods will result in poor image quality.</li>
</ol>
3. Microscope and Camera Setting for Imaging
<ol>
	<li>Adjust the Koehler illumination on the microscope. Choose an objective. Focus on the specimen first. Bring the light intensity to one-half or lower. Close the field diaphragm (FD) and aperture (AP) or bring it to the lowest possible setting on that particular objective.
	<ol>
		<li>The higher the magnification, the larger the ring will be. Use the condenser focusing knob to focus the field iris as sharply as possible.</li>
		<li>Slowly bring the image of the field iris (hexagon-shaped small ring) to the center using the condenser-centering knobs (pin-shaped screws surrounding the condenser). Slowly increase the diaphragm (FD) such that the field iris reaches the edge. Stop increasing the FD just after the field iris has passed the edge.</li>
		<li>Slowly increase the aperture (AP) to adjust contrast. Adjust the light intensity as required. NOTE: Koehler illumination needs to be adjusted for every objective, every experimental day. Note the numbers obtained after adjustment for the day. These numbers will be reused each time that objectives are switched back and forth for that particular day.</li>
	</ol>
	</li>
	<li>Adjust the aperture, intensity, exposure time, gain, and filters as described in Table 1. This information should be used only as a guide.</li>
	<li>Use a high-resolution digital camera with a large format interline CCD chip with no mechanical shutter to capture fluorescence images; and a high-resolution, high-speed camera for bright-field images.</li>
	<li>Process the slides using standard software. Load the software. Click &#8220;Acquire&#8221; and &#8220;Set camera/board,&#8221; then select the camera to be used. Click &#8220;OK.&#8221; Click &#8220;Acquire&#8221; followed by &#8220;Color tab.&#8221; Under &#8220;Color tab,&#8221; choose &#8220;Bayer method&#8221; followed by &#8220;Average four.&#8221; Add the suggested gain (red, green, blue) from Table 1. Select the &#8220;Set save&#8221; tab and click on the directory where images are to be stored. Set &#8220;Exposure time&#8221; from the suggestions in Table 1. Set &#8220;Binning&#8221; at &#8220;1&#8221; and &#8220;Live binning&#8221; at &#8220;1.&#8221; Focus on the region of interest and then click on &#8220;Save live.&#8221;</li>
	<li>Output images on the camera software are in the &#8220;TIF&#8221; format. Analyze the images for histochemistry of the cell types. Use standard computer software for downstream uses of presentation and publication.</li>
</ol>
4. Ultraviolet and Bright-field Imaging
<ol>
	<li>Use a 1 ml pipette with the pipette tip cut so that the sections can be pipetted out easily. Gently pipette sections into the tip and on to a microscope slide. Cover the sections with a cover slip. Observe the sections under UV light or bright-field lighting.</li>
</ol>
5. Phloroglucinol-HCl (Wiesner) Staining12
<ol>
	<li>Dissolve 0.3 g of phloroglucinol in 10 ml absolute ethanol to prepare a 3% phloroglucinol solution. Mix one volume of concentrated HCl (37 N) to two volumes of 3% phloroglucinol in ethanol; this solution is phloroglucinol-HCl (Ph-HCl) or Wiesner stain. NOTE: This solution is to be made fresh on the day of the staining and cannot be stored, because the solution will degrade over time and the staining will become ineffective. CAUTION: HCl is highly corrosive, so use extreme caution while handling the Ph-HCl stain.</li>
	<li>Transfer stem sections to a 2.0 ml microcentrifuge tube. Add 1 ml of the Ph-HCl solution to the tube containing the sections and cap it immediately because the HCl in the Ph-HCl stain is highly corrosive. Gently move the tube to assure that all the sections are stained. An alternative to this step is to keep the tube cap open so that the Ph-HCl solution can be pipetted up and down, preferably without disturbing the sections. NOTE: Be careful not to pipette the sections into the pipette tip, as this may damage those sections.
	<ol>
		<li>Use a 1 ml pipette with the pipette tip cut in such a way that the sections can be pipetted out easily without damage. Gently pipette sections into the tip and onto a microscope slide. Cover the sections with cover slip. Observe the sections under bright-field lighting. NOTE: The Ph-HCl solution dries up in 5-10 min, causing a deterioration of the specimens. Therefore, the imaging has to be completed within that period of time.</li>
	</ol>
	</li>
</ol>
6. M&#228;ule Staining13,14
<ol>
	<li>Dissolve 0.2 g of potassium permanganate in 40 ml distilled water to prepare a 0.5% potassium permanganate solution. Store in a dark bottle at room temperature for a maximum of 7&#160;days.
	<ol>
		<li>Prepare a 3.7% HCl solution by adding 1 ml concentrated HCl (37 N) to 9 ml distilled&#160;water (1:10). Prepare a fresh 3.7% HCl solution on the day of the experiment. Make sure that the 37 N HCl stock solution being used is not too old. Concentrated ammonium hydroxide solution (14.8 M) should be stored at 4 &#176;C to prevent the molarity of the saturated solution from decreasing.</li>
	</ol>
	</li>
	<li>Transfer stem sections to a 2.0 ml microcentrifuge tube. Add 1 ml of the 0.5% potassium permanganate solution to the tube containing the sections.
	<ol>
		<li>Pipette the 0.5% potassium permanganate solution up and down gently, preferably without disturbing the sections. Incubate for 2 min and repeat the pipetting. Let the microcentrifuge tube stand until all the sections settle down. NOTE: Be careful not to pipette the sections into the pipette tip as the sections may get damaged. It may be hard to see the stem sections as the solution is dark, so keep the tube with the sections on test tube rack and allow them to settle for 2 min.</li>
		<li>Using a 1 ml pipette, draw out 700 &#181;l of 0.5% potassium permanganate solution. Add 700 &#181;l of distilled water to rinse out the potassium permanganate solution. Repeat 3-4x or until the water solution stays clear.</li>
		<li>Discard the water. Quickly add 1 ml of 3% HCl until the deep brown color is discharged from the sections. This may happen within 3-5 min or may require 2&#160;washes of 5 min each. Pipette out all the 3% HCl solution and immediately proceed to the next step.</li>
		<li>Add 1 ml of concentrated ammonium hydroxide solution. Use a 1 ml pipette with the tip cut in such a way that the sections can be pipetted out easily without damage. Gently pipette sections into the tip and onto a microscope slide. Cover the sections with a coverslip. Observe the sections under bright-field lighting. CAUTION: Because ammonium hydroxide is extremely corrosive, it requires extra care and attention to avoid causing corrosion to the stage of the microscope or lens. NOTE: The ammonium hydroxide fumes can fog the coverslip, making it difficult to observe the sample. Therefore be extra careful before adding the coverslip. The solution dries up in 5-10 min, so the imaging has to be completed within that period of time.</li>
	</ol>
	</li>
</ol>
7. Congo Red Staining11,15
<ol>
	<li>Dissolve 0.5 g congo red in 100 ml of distilled water to prepare a 0.5% congo red solution.</li>
	<li>Transfer stem sections to a 2.0 ml microcentrifuge tube. Add 1 ml of the 0.5% congo red solution to the tube. Gently pipette the 0.5% congo red solution up and down without disturbing the sections. NOTE: Be careful not to pipette the sections into the pipette tip as this may damage the sections.
	<ol>
		<li>Incubate at room temperature for 10 min and repeat the pipetting. Follow with another 3 min of incubation at room temperature.</li>
		<li>Use a 1 ml pipette to draw out 700&#160;&#181;l of 0.5% congo red solution. Add 700 &#181;l of distilled water to rinse out the congo red solution. Repeat the wash 3-4x until the washing solution is clear.</li>
		<li>Use a 1 ml pipette with its tip cut in such a way that the sections can be pipetted out easily without damage. Gently pipette sections into the tip and onto a microscope slide, and cover them with coverslip. Observe the samples on the microscope slide under blue-light excitation using a filter with a bandpass of 560/40. NOTE: Do not store the sections in water for long prior to the microscopic analysis, as the water will wash out the congo red in 10-30 min.</li>
	</ol>
	</li>
</ol>
8. Calcofluor White Staining9
<ol>
	<li>Dissolve 0.02 g of fluorescent brightener 28 (calcofluor white M2R) in 10 ml distilled water to prepare a 0.2% stock solution. The 0.2% solution can be stored for six months in a dark bottle at 4 &#176;C. Prepare a working solution of 0.02% by diluting the 0.2% stock solution 10x&#160;with distilled water.</li>
	<li>Transfer stem sections to a 2.0 ml microcentrifuge tube. Add to the tube 1 ml of the 0.02% calcofluor white solution. Gently pipette the 0.02% calcofluor white solution up and down. NOTE: Be careful not to pipette the sections into the pipette tip as this may damage the sections.
	<ol>
		<li>Incubate at room temperature for 5 min and repeat the pipetting. Incubate the section for another 3 min at room temperature. NOTE: Allow the sections to settle onto the bottom of the tube to make it easier to pipette out the solution without damaging the sections.</li>
		<li>Use a 1 ml pipette to draw out 700 &#181;l of 0.02% calcofluor white solution and add 700 &#181;l of distilled water; wait for 5 min to rinse out the calcofluor white solution. Repeat the wash 3-4x. Use a 1 ml pipette with the pipette tip cut in such as way that the sections can be pipetted out easily without damage.</li>
		<li>Gently pipette sections into the tip and onto a microscope slide and cover them with a coverslip. Observe the sections under UV light.</li>
	</ol>
	</li>
</ol>
9. Toluidine Blue O Staining16,17
<ol>
	<li>Dissolve 0.02 g toluidine blue O in 100 ml distilled water to prepare a 0.02% solution. NOTE: The toluidine blue O solution can be stored for two weeks in a dark bottle at room temperature.</li>
	<li>Transfer stem sections to a 2.0 ml microcentrifuge tube. Add 1 ml of the 0.02% toluidine blue O solution to the tube. Gently pipette the 0.02% toluidine blue O solution up and down. Then incubate at room temperature for 5 min and repeat the pipetting once. NOTE: Be careful not to pipette the sections into the pipette tip as this may damage the sections. Let the microcentrifuge tube stand still until the sections settle down. This may be hard to see, as the solution is dark, so keep the tube containing the sections on test tube rack and allow them to settle for 2 min at room temperature.
	<ol>
		<li>Use a 1 ml pipette to draw out 700 &#181;l of 0.02% toluidine blue O solution. Add 700 &#181;l of distilled water to rinse out the toluidine blue O solution. Repeat 3-4x or until the wash solution is clear. Use a 1 ml pipette with the pipette tip cut in such a way that the sections can be pipetted out easily without damaging them.</li>
		<li>Gently pipette sections into the tip and onto a microscope slide and cover them with a coverslip. Observe the sections under bright-field lighting.</li>
	</ol>
	</li>
</ol>

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The authors have no financial conflicts to disclose.

A. thaliana stem sections are widely used to study the organization of the cells in the secondary cell wall and to qualitatively examine the differences between wild type and transgenic plants. The commonly used techniques for sectioning the specimens are direct hand cutting; or when specimens are embedded in agarose or fixative, sectioning can be done with a vibratome or microtome. In contrast to hand cutting, the last two allow reducing the risk of producing inaccurate differences between samples that would be...

The plant cell wall holds a plethora of information within its various components: lignin, cellulose, hemicelluloses (xylan, glucuronoxylan, xyloglucan, arabinoxylan, mixed linkage glucan, or glucomannan), and pectin. Histological techniques provide important visual cues in studying differences within the secondary cell walls at organizational and cellular levels. Various histological techniques were developed and can be found in the literature, but these techniques can be challenging and time-consuming for beginners because very detailed protocols with simple visual instructions are seldom if ever available. The goal of this study is to provide simple guidelines for histological staining techniques to obtain high-quality images.
Sectioning the stem tissue is the first step in visualization of cell walls and cell shapes. Although hand-cut sections are inexpensive and take less time to prepare, use of a vibratome offers consistency and yields high-quality images. Using a vibratome allows producing better data quality by generating even sections with the same thickness, which helps to produce sharp images and reduces significantly the risk of producing inaccurate differences between samples that would be simply caused by a bad sample preparation. Using resins to fix fresh specimens can be a challenge for beginners and may still be time-consuming even for experts when an analysis needs to be done quickly. In addition, it becomes impossible to measure any biological activity with the sample when it has been embedded in a resin. One simple technique that employs agarose and a homemade mold is helpful for embedding stem tissues, and it can also be utilized in other applications requiring dissection of soft tissues. Compared to embedding specimens in resin, this method has the advantage of keeping the tissues alive and reducing sample manipulation. Sectioning tissues via a vibratome is highly precise and generates homogenous sections, which, depending on the goal of the study, can then be used with several different staining techniques.
The simplest methods to visualize lignin and other aromatics employ ultraviolet (UV) light. Excitation of aromatic-based molecules by UV light is an old technique, but it is still one of the quickest approaches for visualization of lignin. However, UV visualization actually is not ideal for lignin detection because UV light will excite other aromatics. Lignin is made up primarily of three building blocks, the monolignols (hydroxycinnamyl alcohols: coniferyl alcohol,&#160;sinapyl alcohol,&#160;and p-coumaryl alcohol)1-3. Phloroglucinol stain can provide clues to the extent of cinnamaldehydes present in the xylem, fiber, and tracheal tissues4. Phloroglucinol is a good indicator of general cinnamaldehydes and can differentiate between cinnamaldehydes and other aromatics. The sinapyl alcohol monomers can be detected and differentiated by the use of M&#228;ule stain. Toluidine blue O is a polychromatic dye and therefore has the ability to stain different elements of the cell wall in different colors5,6. The primary use of toluidine blue O is to detect pectin and lignin5,6. The advantage of using toluidine blue O is that many elements of the cell wall can be visualized in a single step. Both calcofluor white and congo red are easy to work with and can be used to visualize cellulose. Calcofluor white stains cellulose, callose, and other non-substituted or weakly substituted &#946;-glucans6-9, whereas congo red stains directly to &#946;-(1&#8594;4)-glucans and particularly to cellulose10,11. The goal of this study is to provide simple guidelines for the use of the aforementioned staining techniques to obtain high-quality images from A. thaliana stems.

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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.

Stem Embedding and Sectioning: 
The use of the homemade plastic mold to embed the stems in 7% agarose proved to be fast and easy (Figure 1). The two parts (A and B; Figure 1) of the embedded vial system make it simple to easily release the stem embedded in agarose as the agarose does not stick to the vial parts that are inert, keeping the system clean. The vials can be reused multiple times for years. The convenience of storing the embedded stem also makes the next steps easier. In ...

Watch this Scientific Journal Video about Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements at JoVE.com

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.

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 ...

histochemical-staining-arabidopsis-thaliana-secondary-cell-wall

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40.3K Views.  Joint Bioenergy Institute. 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.

Video: Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Isolation of Protoplasts from Tissues of 14-day-old Seedlings of Arabidopsis thaliana

Protoplasts are plant cells that have had their cell walls enzymatically removed. Isolation of protoplasts from different plant tissues was first reported more than 40 years ago 1 and has since been adapted to study a variety of cellular processes, such as subcellular localization of proteins, isolation of intact organelles and targeted gene-inactivation by double stranded RNA interference (RNAi) 2-5. Most of the protoplast isolation protocols use leaf tissues of mature Arabidopsis (e.g. 35-day-old plants) 2-4. We modified existing protocols by employing 14-day-old Arabidopsis seedlings. In this procedure, one gram of 14-day-old seedlings yielded 5 106-107 protoplasts that remain intact at least 96 hours. The yield of protoplasts from seedlings is comparable with preparations from leaves of mature Arabidopsis, but instead of 35-36 days, isolation of protoplasts is completed in 15 days. This allows decreasing the time and growth chamber space that are required for isolating protoplasts when mature plants are used, and expedites the downstream studies that require intact protoplasts.

Isolation of Protoplasts from Tissues of 14-day-old Seedlings of Arabidopsis thaliana

This video shows a procedure for isolating intact protoplasts from tissues of 14-day-old seedlings of Arabidopsis. Given that the isolated protoplasts remain intact for at least 96h and are isolated from seedlings instead of one-month-old mature plants, this procedure expedites assays requiring intact protoplasts.

Protoplasts are plant cells that have had their cell walls enzymatically removed. Isolation of protoplasts  from different plant tissues was first ...

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called photoreceptors to sense and adapt to light. The red/far-red light-absorbing phytochrome photoreceptors have been studied extensively. Phytochromes exist as a family of proteins with distinct and overlapping functions in all higher plant systems in which they have been studied1. Phytochrome-mediated light responses, which range from seed germination through flowering and senescence, are often localized to specific plant tissues or organs2. Despite the discovery and elucidation of individual and redundant phytochrome functions through mutational analyses, conclusive reports on distinct sites of photoperception and the molecular mechanisms of localized pools of phytochromes that mediate spatial-specific phytochrome responses are limited. We designed experiments based on the hypotheses that specific sites of phytochrome photoperception regulate tissue- and organ-specific aspects of photomorphogenesis, and that localized phytochrome pools engage distinct subsets of downstream target genes in cell-to-cell signaling. We developed a biochemical approach to selectively reduce functional phytochromes in an organ- or tissue-specific manner within transgenic plants. Our studies are based on a bipartite enhancer-trap approach that results in transactivation of the expression of a gene under control of the Upstream Activation Sequence (UAS) element by the transcriptional activator GAL43. The biliverdin reductase (BVR) gene under the control of the UAS is silently maintained in the absence of GAL4 transactivation in the UAS-BVR parent4. Genetic crosses between a UAS-BVR transgenic line and a GAL4-GFP enhancer trap line result in specific expression of the BVR gene in cells marked by GFP expression4. BVR accumulation in Arabidopsis plants results in phytochrome chromophore deficiency in planta5-7. Thus, transgenic plants that we have produced exhibit GAL4-dependent activation of the BVR gene, resulting in the biochemical inactivation of phytochrome, as well as GAL4-dependent GFP expression. Photobiological and molecular genetic analyses of BVR transgenic lines are yielding insight into tissue- and organ-specific phytochrome-mediated responses that have been associated with corresponding sites of photoperception4, 7, 8. Fluorescence Activated Cell Sorting (FACS) of GFP-positive, enhancer-trap-induced BVR-expressing plant protoplasts coupled with cell-type-specific gene expression profiling through microarray analysis is being used to identify putative downstream target genes involved in mediating spatial-specific phytochrome responses. This research is expanding our understanding of sites of light perception, the mechanisms through which various tissues or organs cooperate in light-regulated plant growth and development, and advancing the molecular dissection of complex phytochrome-mediated cell-to-cell signaling cascades.

Investigating Tissue- and Organ-specific Phytochrome Responses using FACS-assisted Cell-type Specific Expression Profiling in Arabidopsis thaliana

The molecular basis of spatial-specific phytochrome responses is being investigated using transgenic plants that exhibit tissue- and organ-specific phytochrome deficiencies. The isolation of specific cells exhibiting induced phytochrome chromophore depletion by Fluorescence-Activated Cell Sorting followed by microarray analyses is being utilized to identify genes involved in spatial-specific phytochrome responses.

Light mediates an array of developmental and adaptive processes throughout the life cycle of a plant. Plants utilize light-absorbing molecules called ...

Floral-dip Transformation of Arabidopsis thaliana to Examine pTSO2::&beta;-glucuronidase Reporter Gene Expression

The ability to introduce foreign genes into an organism is the foundation for modern biology and biotechnology. In the model flowering plant Arabidopsis thaliana, the floral-dip transformation method1-2 has replaced all previous methods because of its simplicity, efficiency, and low cost. Specifically, shoots of young flowering Arabidopsis plants are dipped in a solution of Agrobacterium tumefaciens carrying specific plasmid constructs. After dipping, the plants are returned to normal growth and yield seeds, a small percentage of which are transformed with the foreign gene and can be selected for on medium containing antibiotics. This floral-dip method significantly facilitated Arabidopsis research and contributed greatly to our understanding of plant gene function. In this study, we use the floral-dip method to transform a reporter gene, &beta;-glucuronidase (GUS), under the control of TSO2 promoter. TSO2, coding for the Ribonucleotide Reductase (RNR) small subunit3, is a cell cycle regulated gene essential for dNDP biosynthesis in the S-phase of the cell cycle. Examination of GUS expression in transgenic Arabidopsis seedlings shows that TSO2 is expressed in actively dividing tissues. The reported experimental method and materials can be easily adapted not only for research but also for education at high school and college levels.

Floral-dip Transformation of Arabidopsis thaliana to Examine pTSO2::&#946;-glucuronidase Reporter Gene Expression

This article illustrates the floral-dip method of Agrobacterium tumefaciens -mediated transformation of Arabidopsis thaliana. By introducing a cell-cycle regulated promoter-reporter, pTSO2::&beta;-glucuronidase (GUS), into Arabidopsis, we illustrates how one detects GUS reporter expression in transgenic seedlings.

The ability to introduce foreign genes into an organism is the foundation for modern biology and biotechnology. In the model flowering plant Arabidopsis ...

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography (TLC) Coupled with Gas-Liquid Chromatography (GLC)

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as phosphatidylcholine or phosphatidylethanolamine, form bilayer membranes which act as both boundaries and interfaces for chemical exchange between cells and their surroundings. Unlike animals, plant cells have a special organelle for photosynthesis, the chloroplast. The intricate membrane system of the chloroplast contains unique glycerolipids, namely glycolipids lacking phosphorus: monogalactosyldiacylglycerol (MGDG), digalactosyldiacylglycerol (DGDG), sulfoquinovosyldiacylglycerol (SQDG)4. The roles of these lipids are beyond simply structural. These glycolipids and other glycerolipids were found in the crystal structures of photosystem I and II indicating the involvement of glycerolipids in photosynthesis8,11. During phosphate starvation, DGDG is transferred to extraplastidic membranes to compensate the loss of phospholipids9,12.
Much of our knowledge of the biosynthesis and function of these lipids has been derived from a combination of genetic and biochemical studies with Arabidopsis thaliana14. During these studies, a simple procedure for the analysis of polar lipids has been essential for the screening and analysis of lipid mutants and will be outlined in detail. A leaf lipid extract is first separated by thin layer chromatography (TLC) and glycerolipids are stained reversibly with iodine vapor. The individual lipids are scraped from the TLC plate and converted to fatty acyl methylesters (FAMEs), which are analyzed by gas-liquid chromatography coupled with flame ionization detection (FID-GLC) (Figure 1). This method has been proven to be a reliable tool for mutant screening. For example, the tgd1,2,3,4 endoplasmic reticulum-to-plastid lipid trafficking mutants were discovered based on the accumulation of an abnormal galactoglycerolipid: trigalactosyldiacylglycerol (TGDG) and a decrease in the relative amount of 18:3 (carbons : double bonds) fatty acyl groups in membrane lipids 3,13,18,20. This method is also applicable for determining enzymatic activities of proteins using lipids as substrate6.

Arabidopsis thaliana Polar Glycerolipid Profiling by Thin Layer Chromatography TLC Coupled with Gas-Liquid Chromatography GLC

Composition of polar lipid extracts and the fatty acid composition of individual glycerolipids are determined in a simple and robust lipid profiling experiment. For this purpose, glycerolipids are isolated by thin layer chromatography and subjected to transmethylation of their acyl groups. Fatty acyl methylesters are quantified by gas-liquid chromatography.

Biological membranes separate cells from the environment. From a single cell to multicellular plants and animals, glycerolipids, such as ...

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the Arabidopsis leaf is a complex organ made up of many different cell types and several structures. At the same time, the growing leaf contains cells at different stages of development, with the cells furthest from the petiole being the first to stop expanding and undergo senescence1. Different cells within the leaf are therefore dividing, elongating or differentiating; active, stressed or dead; and/or responding to stimuli in sub-sets of their cellular type at any one time. This makes genomic study of the leaf challenging: for example when analyzing expression data from whole leaves, signals from genetic networks operating in distinct cellular response zones or cell types will be confounded, resulting in an inaccurate profile being generated.
To address this, several methods have been described which enable studies of cell specific gene expression. These include laser-capture microdissection (LCM)2 or GFP expressing plants used for protoplast generation and subsequent fluorescence activated cell sorting (FACS)3,4, the recently described INTACT system for nuclear precipitation5 and immunoprecipitation of polysomes6.
FACS has been successfully used for a number of studies, including showing that the cell identity and distance from the root tip had a significant effect on the expression profiles of a large number of genes3,7. FACS of GFP lines have also been used to demonstrate cell-specific transcriptional regulation during root nitrogen responses and lateral root development8, salt stress9 auxin distribution in the root10 and to create a gene expression map of the Arabidopsis shoot apical meristem11. Although FACS has previously been used to sort Arabidopsis leaf derived protoplasts based on autofluorescence12,13, so far the use of FACS on Arabidopsis lines expressing GFP in the leaves has been very limited4. In the following protocol we describe a method for obtaining Arabidopsis leaf protoplasts that are compatible with FACS while minimizing the impact of the protoplast generation regime. We demonstrate the method using the KC464 Arabidopsis line, which express GFP in the adaxial epidermis14, the KC274 line, which express GFP in the vascular tissue14 and the TP382 Arabidopsis line, which express a double GFP construct linked to a nuclear localization signal in the guard cells (data not shown; Figure 2). We are currently using this method to study both cell-type specific expression during development and stress, as well as heterogeneous cell populations at various stages of senescence.

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

A method for producing Arabidopsis leaf protoplasts that are compatible with fluorescence activated cell sorting (FACS), allowing for studies of specific cell populations. This method is compatible with any Arabidopsis line that expresses GFP in a subset of cells.

After initiation of the leaf primordium, biomass accumulation is controlled mainly by cell proliferation and expansion in the leaves1. However, the ...

Glycan Profiling of Plant Cell Wall Polymers using Microarrays

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the raw materials for human societies (e.g. wood, paper, textile and biofuel industries)1,2. However, understanding the biosynthesis and function of these components remains challenging.
Cell wall glycans are chemically and conformationally diverse due to the complexity of their building blocks, the glycosyl residues. These form linkages at multiple positions and differ in ring structure, isomeric or anomeric configuration, and in addition, are substituted with an array of non-sugar residues. Glycan composition varies in different cell and/or tissue types or even sub-domains of a single cell wall3. Furthermore, their composition is also modified during development1, or in response to environmental cues4.
In excess of 2,000 genes have Plant cell walls are complex matrixes of heterogeneous glycans been predicted to be involved in cell wall glycan biosynthesis and modification in Arabidopsis5. However, relatively few of the biosynthetic genes have been functionally characterized 4,5. Reverse genetics approaches are difficult because the genes are often differentially expressed, often at low levels, between cell types6. Also, mutant studies are often hindered by gene redundancy or compensatory mechanisms to ensure appropriate cell wall function is maintained7. Thus novel approaches are needed to rapidly characterise the diverse range of glycan structures and to facilitate functional genomics approaches to understanding cell wall biosynthesis and modification.
Monoclonal antibodies (mAbs)8,9 have emerged as an important tool for determining glycan structure and distribution in plants. These recognise distinct epitopes present within major classes of plant cell wall glycans, including pectins, xyloglucans, xylans, mannans, glucans and arabinogalactans. Recently their use has been extended to large-scale screening experiments to determine the relative abundance of glycans in a broad range of plant and tissue types simultaneously9,10,11.
Here we present a microarray-based glycan screening method called Comprehensive Microarray Polymer Profiling (CoMPP) (Figures 1 &amp; 2)10,11 that enables multiple samples (100 sec) to be screened using a miniaturised microarray platform with reduced reagent and sample volumes. The spot signals on the microarray can be formally quantified to give semi-quantitative data about glycan epitope occurrence. This approach is well suited to tracking glycan changes in complex biological systems12 and providing a global overview of cell wall composition particularly when prior knowledge of this is unavailable.

A technique called Comprehensive Microarray Polymer Profiling (CoMPP) for the characterisation of plant cell wall glycans is described. This method combines the specificity of monoclonal antibodies directed to defined glycan-epitopes with a miniature microarray analytical platform allowing screening of glycan occurrence in a broad range of biological contexts.

Plant cell walls are complex matrixes of heterogeneous glycans which play an important role in the physiology and development of plants and provide the ...

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a consequence, the isolation of plant embryos at early stages (1 cell to globular stage) is technically challenging due to their relative inaccessibility. Efficient manual dissection at early stages is strongly impaired by the small size of young Arabidopsis seeds and the adhesiveness of the embryo to the surrounding tissues. Here, we describe a method that allows the efficient isolation of young Arabidopsis embryos, yielding up to 40 embryos in 1 hr to 4 hr, depending on the downstream application. Embryos are released into isolation buffer by slightly crushing 250-750 seeds with a plastic pestle in an Eppendorf tube. A glass microcapillary attached to either a standard laboratory pipette (via a rubber tube) or a hydraulically controlled microinjector is used to collect embryos from droplets placed on a multi-well slide on an inverted light microscope. The technical skills required are simple and easily transferable, and the basic setup does not require costly equipment. Collected embryos are suitable for a variety of downstream applications such as RT-PCR, RNA sequencing, DNA methylation analyses, fluorescence in situ hybridization (FISH), immunostaining, and reporter gene assays.

Efficient and Rapid Isolation of Early-stage Embryos from Arabidopsis thaliana Seeds

We report an efficient and simple method to isolate embryos at early stages of development from Arabidopsis thaliana seeds. Up to 40 embryos can be isolated in 1 hr to 4 hr, depending on the downstream application. The procedure is suitable for transcriptome, DNA methylation, reporter gene expression, immunostaining and fluorescence in situ hybridization analyses.

In flowering plants, the embryo develops within a nourishing tissue - the endosperm - surrounded by the maternal seed integuments (or seed coat). As a ...

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling processes. Extracting sterol-enriched membrane microdomains from plant cells for proteomic analysis is a difficult task mainly due to multiple preparation steps and sources for contaminations from other cellular compartments. The plasma membrane constitutes only about 5-20% of all the membranes in a plant cell, and therefore isolation of highly purified plasma membrane fraction is challenging. A frequently used method involves aqueous two-phase partitioning in polyethylene glycol and dextran, which yields plasma membrane vesicles with a purity of 95% 1. Sterol-rich membrane microdomains within the plasma membrane are insoluble upon treatment with cold nonionic detergents at alkaline pH. This detergent-resistant membrane fraction can be separated from the bulk plasma membrane by ultracentrifugation in a sucrose gradient 2. Subsequently, proteins can be extracted from the low density band of the sucrose gradient by methanol/chloroform precipitation. Extracted protein will then be trypsin digested, desalted and finally analyzed by LC-MS/MS. Our extraction protocol for sterol-rich microdomains is optimized for the preparation of clean detergent-resistant membrane fractions from Arabidopsis thaliana cell cultures.
We use full metabolic labeling of Arabidopsis thaliana suspension cell cultures with K15NO3 as the only nitrogen source for quantitative comparative proteomic studies following biological treatment of interest 3. By mixing equal ratios of labeled and unlabeled cell cultures for joint protein extraction the influence of preparation steps on final quantitative result is kept at a minimum. Also loss of material during extraction will affect both control and treatment samples in the same way, and therefore the ratio of light and heave peptide will remain constant. In the proposed method either labeled or unlabeled cell culture undergoes a biological treatment, while the other serves as control 4.

Metabolic Labeling and Membrane Fractionation for Comparative Proteomic Analysis of Arabidopsis thaliana Suspension Cell Cultures

Here we describe a robust method for the fractionation of plant plasma membranes into detergent resistant and detergent soluble membranes based on a mixture of unlabeled and in vivo fully 15N labeled Arabidopsis thaliana cell cultures. The procedure is applied for comparative proteomic studies to understand signaling processes.

Plasma membrane microdomains are features based on the physical properties of the lipid and sterol environment and have particular roles in signaling ...

A Seed Coat Bedding Assay to Genetically Explore In Vitro How the Endosperm Controls Seed Germination in Arabidopsis thaliana

The Arabidopsis endosperm consists of a single cell layer surrounding the mature embryo and playing an essential role to prevent the germination of dormant seeds or that of nondormant seeds irradiated by a far red (FR) light pulse. In order to further gain insight into the molecular genetic mechanisms underlying the germination repressive activity exerted by the endosperm, a &#34;seed coat bedding&#34; assay (SCBA) was devised. The SCBA is a dissection procedure physically separating seed coats and embryos from seeds, which allows monitoring the growth of embryos on an underlying layer of seed coats. Remarkably, the SCBA reconstitutes the germination repressive activities of the seed coat in the context of seed dormancy and FR-dependent control of seed germination. Since the SCBA allows the combinatorial use of dormant, nondormant and genetically modified seed coat and embryonic materials, the genetic pathways controlling germination and specifically operating in the endosperm and embryo can be dissected. Here we detail the procedure to assemble a SCBA.

A Seed Coat Bedding Assay to Genetically Explore In Vitro How the Endosperm Controls Seed Germination in Arabidopsis thaliana

We present the procedure to assemble a seed coat bedding assay (SCBA) from Arabidopsis thaliana seeds. The SCBA was shown to be a powerful tool to explore genetically and in vitro how the endosperm controls seed germination in dormant seeds and in response to light cues. The SCBA is in principle applicable under any situation where the endosperm is suspected to influence embryonic growth.

The Arabidopsis endosperm consists of a single cell layer surrounding the mature embryo and playing an essential role to prevent the germination of ...

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.

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.

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 ...