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 ...

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40.0K 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 ...

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::&amp;#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 ...

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 ...

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 ...

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light conditions, which are preferably variable in quantity and quality so that studies under different setups can be conducted. These requirements are met by climatic chambers featuring light emitting diodes (LED) lights, which can &#8211; in contrast to fluorescent lights &#8211; be set to different wavelengths. LEDs are energy conserving and emit virtually no heat even at light intensities, which often constitutes a problem with other light sources. The presented protocol provides a step-by-step guidance of how to program a climatic chamber equipped with variable LED lights as well as describing several approaches for in depth analysis of growth phenotypes. Depending on the experimental set-up various characteristics of the growing plants can be observed and analyzed. Here we describe how to determine fresh weight, leaf area, photosynthetic activity, and stomatal density. We demonstrate that in order to obtain reliable data and draw valid conclusions it is mandatory to use a sufficient number of individuals for statistical evaluation. Taking too few plants for this kind of analysis results in high statistical errors and consequently in less clear interpretations of the data.

Analysis of Arabidopsis thaliana Growth Behavior in Different Light Qualities

Here, we present a protocol for studying plant growth behavior and especially phenotypes in a reproducible manner. We show how to provide variable and at the same time stable light conditions. Proper analyses depend on sufficient sample numbers and valid statistical evaluations.

Plant biologists often need to observe the growth behavior of their chosen species. To this end, the plants need constant environmental and stable light ...

Spatiotemporal Investigation of ERL1 Dynamics via Confocal Imaging

Image-Based Methods to Study Membrane Trafficking Events in Stomatal Lineage Cells

In eukaryotic cells, membrane components, including proteins and lipids, are spatiotemporally transported to their destination within the endomembrane system. This includes the secretory transport of newly synthesized proteins to the cell surface or the outside of the cell, the endocytic transport of extracellular cargoes or plasma membrane components into the cell, and the recycling or shuttling transport of cargoes between the subcellular organelles, etc. Membrane trafficking events are crucial to the development, growth, and environmental adaptation of all eukaryotic cells and, thus, are under stringent regulation. Cell-surface receptor kinases, which perceive ligand signals from the extracellular space, undergo both secretory and endocytic transport. Commonly used approaches to study the membrane trafficking events using a plasma membrane-localized leucine-rich-repeat receptor kinase, ERL1, are described here. The approaches include plant material preparation, pharmacological treatment, and confocal imaging setup. To monitor the spatiotemporal regulation of ERL1, this study describes the co-localization analysis between ERL1 and a multi-vesicular body marker protein, RFP-Ara7, the time series analysis of these two proteins, and the z-stack analysis of ERL1-YFP treated with the membrane trafficking inhibitors brefeldin A and wortmannin.

Author Spotlight: Image-Based Methods to Study Membrane Trafficking Events in Stomatal Lineage Cells  

Several commonly used methods are introduced here to study the membrane trafficking events of a plasma membrane receptor kinase. This manuscript describes detailed protocols including the plant material preparation, pharmacological treatment, and confocal imaging setup.

In eukaryotic cells, membrane components, including proteins and lipids, are spatiotemporally transported to their destination within the endomembrane ...

An Improved Method to Measure Pollen Hydration Profiles in &lt;em&gt;Arabidopsis thaliana&lt;/em&gt;

A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is established between the interacting partners. Studies across a range of species have revealed that a series of molecular checkpoints regulate the pollen-stigma interaction to ensure that only compatible, generally intraspecific pollen is successful in effecting fertilization. In species that possess a 'dry stigma', such as the model plant Arabidopsis thaliana, the first post-pollination, prezygotic compatibility checkpoint is the establishment of pollen hydration. 
This phase of pollination is tightly regulated, whereby signals from the pollen grain elicit the release of water from the stigma, thus permitting pollen hydration. The ability to accurately measure and track pollen hydration over time is key to the design of experiments directed at understanding the regulation of this critical step in reproduction. Published protocols frequently utilize flowers that have been excised from the parent plant, maintained on liquid or solid media, and bulk pollinated. 
This paper describes a noninvasive, in vivo pollination bioassay that permits minute-by-minute hydration tracking of individual A. thaliana pollen grains at high resolution. The assay is highly reproducible, able to detect very subtle variations of pollen hydration profiles, and thus is suitable for the analysis of mutants that affect pathways regulating pollination. Although the protocol is lengthier than those described for bulk pollinations, the precision and reproducibility it provides, along with its in vivo nature, make it ideal for the detailed dissection of pollination phenotypes.

Author Spotlight: A High-Resolution, Single-Grain, In Vivo Pollen Hydration Bioassay for Arabidopsis thaliana  

An improved method to measure pollen hydration profiles in Arabidopsis thaliana is described here. The new method offers higher resolution, is noninvasive, and is highly reproducible. The protocol represents a new tool for a finer dissection of the processes that regulate the early stages of pollination.

Sexual reproduction in flowering plants requires initial interaction between the pollen grain and the stigmatic surface, where a molecular dialog is ...

Pollen Hydration Assay in Arabidopsis thaliana: Raw Data Acquisition and its Measurements

Pollen Hydration Assay in Arabidopsis thaliana: Raw Data Acquisition and its Measurements  

Begin the assay by laying the labeled A.thaliana pollen recipient plant on its side, positioning the emasculated flower on the stage of an inverted microscope to image the stigma. To fix the position of the emasculated recipient flower, immobilize the stem using strips of masking tape. From the pollen donor plant, remove a healthy and freshly-opened flower and place the pollen donor plant under a dissecting microscope.Gather pollen grains using fine-tipped forceps. Remove excess pollen grains by lightly touching the forceps against the donor petals until the pollen grain's monolayer is formed. Return to the pollen recipient plant and use a low power objective lens to focus the inverted microscope on the stigma.Holding the forceps along the opening between the arms of the forceps, carefully approach and unpollinated stigmatic papilla cell. Continue approaching the unpollinated stigmatic papilla cell until the suitably positioned pollen grain on the forceps makes light contact with its surface. Once pollen attachment is confirmed, slowly withdraw the forceps.Orient the pollen grain with its equatorial axis visible and in sharp focus, and immediately switch to a higher power objective lens, like 20x. Capture the first pollen grain image as T-0, and continue capturing images at one-minute intervals for 10 minutes. Adjust the focus to accommodate small movements in the pollen grain or stigma.Record the room's ambient temperature and relative humidity every two minutes, allowing future comparisons between experimental replicates. Save the images in a lossless format, such as ND2, before repeating the pollination and imaging steps for additional pollen grains to acquire data for control and experimental pollinations. Begin the data measurements for pollen hydration assay using image analysis software.Use similar parameters of digital zoom and the approach defining the pollen boundary for all measurements in the datasets. Record the semiminor axis values for all time points in the dataset. Once the measurements are complete for a dataset, export the raw semiminor axis values of each individual time point to a spreadsheet.Present the data in columns per image stack and calculate the percentage semiminor axis change. Repeat the steps and obtain data from at least 15 hydrated pollen grains for each plant line. A few pollen grains may fail to hydrate or may hydrate significantly slower than expected.They're known as dud grains. Exclude these from the dataset. Calculate the mean values for each time point per plant using the specialized software package GraphPad Prism.Analyze the hydration data from wild-type and mutant lines at each time point using unpaired T-test and one-way ANOVA to compare the means of the specific time point of interest. To compare the means across all the time points, perform multiple T-tests between wild-type and mutant lines across multiple time points. The pollen hydration time series data for wild-type plants collected on different days showed that the minimum and maximum values for the means between replicates for all time points were within 3%This representative data for wild-type pollinations demonstrated a high degree of consistency for relatively low sample numbers and across different days.

﻿Preparation of Dual-Colored F1 Transgenic Arabidopsis thaliana

Preparation of Dual-Colored F1 Transgenic Arabidopsis thaliana  

To begin, grow homozygous transgenic Arabidopsis thaliana plants bearing ERL1-YFP and a red fluorescent protein, or RFP-labeled marker protein RFP ARA7 side-by-side in a growth room until flowering. On the young inflorescence of plant A, remove all the older flowers and calyx, except for one recently opened flower. Carefully remove the younger flowers and floral merostems.In the recently opened flower, remove the sepals, petals, and stamen using a pair of sharp tweezers, leaving only the pistil on the inflorescence. From plant B, take mature stamen from an opened flower and deposit the pollen grains into the stigma of the dissected pistil of plant A.Label this manually crossed flower, indicating its parents and the date of the genetic cross. When the calyx turns yellow or brown, harvest the F1 seeds to check for a successful F1 plant having yellow fluorescent protein, or YFP, and RFP signals.