Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

The overall goal of this procedure is to show histochemical staining of Arabidopsis ANA secondary cell wall elements. This is accomplished by first embedding the plant stem into aros. Then the aros embedded stem is de casted and transferred to specimen discs, which are then sectioned into STEM cross-sections using a vibrato.

Next, the sections are subjected to various histochemical staining procedures. Ultimately, microscopy is used to show the differential changes in the elements of the secondary cell wall obtained by the histochemical staining. This method can help to answer key questions in cell wall, such as changes of macular tissues in plant stems.

Generally, individuals new to this method will struggle because simple yet critical steps mentioned in the protocols here have not been thoroughly documented or presented in a simplistic manner before. To begin, prepare a homemade mold for embedding the stems using plastic vials. First, use a razor blade to cut off the conical bottom of a two milliliter screw cap micro centrifuge tube.

This component is referred to as 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 half a centimeter of the bottom of a 0.6 milliliter micro centrifuge tube. This component will be referred to as Part B.Add the cutoff half centimeter portion of the 0.6 milliliter micro centrifuge tube into the precut two milliliter micro centrifuge tube.

Seal the two parts using paraform. Using a razor blade, cut a stem section from the area of interest, melt a 7%aros solution in the microwave at low intensity. Once the melted aros cools to about 50 degrees Celsius, slowly pipette five milliliters of the agros into the pre-made plastic mold to fill the vial.

After the aros has cooled for 30 to 60 seconds and is a semi-solid, place the stem into the aeros filled vial. Then place the perforated cap in such a way that the tip of the stem is held by the cap. Leave the vial at room temperature or at four degrees Celsius for 10 to 30 minutes until it solidifies.

Remove the mold by gently sliding it out of the tube. Open the para film and push the bottom of part B of the mole gently with the thumb to release the solidified aros from part A onto a glass microscope. Slide cut.

The aros embedded stem into three approximately equal pieces of 1.2 centimeters in length. Cut out the part of the stem that was not in the aros and discarded. Make sure the specimen disc is clean and completely dry.

Use soft paper wipes to remove any moisture from the aros block containing the specimens. Place a small drop of tissue adhesive on the specimen disc. Quickly place the aros block so that the specimens are either perpendicular to the plate for transversal cross-sections or in parallel with the plate.

For longitudinal cross-sections, allow the adhesive to fix the sample to the specimen disc at room temperature or at four degrees Celsius for 10 to 30 minutes. Fix the specimen disc in place on the buffer tray or trough the block of aros 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.

Cut a razor blade in half and then trim the ends with sturdy scissors so that the blade stays completely flat. Attach one half of the precut razor blade onto the knife holder. Set the angle on the knife holder to 84 degrees the speed to 0.90 millimeters per second and the frequency to 50 hertz.

Cut the sections to 100 micron thickness using continuous mode. Collect the sections using a disposable plastic pipette during sectioning in the buffer tray. Transfer a few sections to the two milliliter micro centrifuge tubes for fluorol staining.

Transfer the stem sections to a micro centrifuge tube. Add one milliliter of the fluorol solution to the tube and cap it immediately. Because the hydrochloric acid in the fluoro gluc anol stain is highly corrosive, gently move the tube to assure that all the sections are stained.

Gently pipee the sections into a cut pipette tip and onto a microscope.Slide. Cover the sections with a cover slip. Observe the sections under brightfield lighting.

The solution dries up in five to 10 minutes causing a deterioration of the specimens. Therefore, the imaging has to be completed within that period of time. For mal staining, transfer the stem sections to a micro centrifuge tube.

Add one milliliter of the 0.5%potassium permanganate solution to the tube containing the sections. Pipette the 0.5%potassium permanganate solution up and down gently, preferably without disturbing the sections. After a two minute incubation and repeat pipetting, let the micro centrifuge tube stand until all the sections settle down.

Using a one milliliter pipette, draw out 700 microliters of 0.5%Potassium permanganate solution, add 700 microliters of distilled water to rinse out the potassium permanganate. Repeat three to four times or until the water solution stays clear. After discarding the water quickly add one milliliter of 3%hydrochloric acid until the deep brown color is discharged from the sections.

This may happen within three to five minutes or may require two washes of five minutes. Each pipette out all of the 3%hydrochloric acid solution and immediately add one milliliter of concentrated ammonium hydroxide solution. Before imaging under brightfield lighting for Congo red staining, transfer the stem sections to a micro centrifuge tube and add one milliliter of the 0.5%Congo red solution.

Gently pipette the 0.5%Congo red solution up and down without disturbing the sections. Before incubating and washing the sections as described in the text protocol, gently pipette the dissects into a cut pipette tip and onto a microscope slide. Then cover them with a cover slip.

Observe the samples on the microscope. Slide under blue light excitation using a 560 nanometer emission filter for calcior white staining. Transfer the stem sections to a micro centrifuge tube and add one milliliter of the 0.02%calcior white solution.

Gently pipette the solution up and down. Proceed to incubate and wash the sections as detailed in the text protocol before observing the sections under UV light. For toluidine blue o staining transfer stem sections to a micro centrifuge tube.

Then add one milliliter of the 0.02%toluidine blue O solution to the tube and gently pipette the solution up and down. Then incubate and wash the sections as described in the text protocol before observing them under brightfield lighting. See the text protocol for details on microscope and camera settings for imaging as well as for the ultraviolet and brightfield imaging procedure.

The aromatic compounds, including the lignin in the cells, can be visualized by their autofluorescence under UV light. Xylem and interf fibers showed autofluorescence under UV illumination, but not in pith and cortex or epidermis because lignin and aromatic polymer is deposited during secondary cell while biosynthesis fluorol stain reacts with aldehyde end groups of lignin to give a pink or fuchsia color as observed under UV illumination. A fuchsia coloration is observed in xylem and interf fibers, but is absent in pith and cortex or epidermis.

Usually, the intensity of the color correlates with the level of lignification in a qualitative manner. The MALS stain is specific in detecting the ingal lignin units in xylem and interf fibers. Red coloration indicates the presence of ingal lignin units in the lignin elements.

However, a brighter red coloration is observed in the fibers when compared to the xylem tissues, which suggests that the fibers contain a higher level of ingal lignin, whereas the xylem is more enriched in gua lignin units. Stem sections stained with calcior white were observed under UV light and revealed that the epidermis cortex and pith were stained because all of these tissues contained cellulose as a major polysaccharide polymer in their cell walls. Congo red stained sections were observed under blue light excitation and also stained the epidermis cortex and pith.

In contrast to calcior white Congo red stained polysaccharides better in the xylem and interf fibers and seemed less affected by the presence of lignin odine. Blue O is classified as a polychromatic dye because it results in a multicolored specimen. Odine blue O is a onic dye that binds to negatively charged groups and generates different colors.

When the dye binds with different onic groups in the cell, taine blue O of stem cross sections revealed that xylem and inter fascicular fibers are lignified since they show a greenish blue or blue coloration, which is in agreement with the UV and fluoro glucan. All staining observations in contrast, the pith and cortex and epidermis tissues show greenish blue or blue coloration because although they are not lignified, they contain some pectin polymers in their cell walls. After watching this video, you should have a good understanding of how to section plant stems and to differentially stain the various tissues in the plant cell wall with different histochemical stains.