Name: fluorescent immunolocalization of arabinogalactan proteins and pectins in the cell wall of plant tissues
Uploaded: 2021-02-27T13:00:00.000+00:00
Duration: 10 min 14 s
Description: Watch this Scientific Journal Video about Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues at JoVE.com

The authors received support from the EU project 690946 &#8216;SexSeed&#8217; (Sexual Plant Reproduction &#8211; Seed Formation) funded by H2020-MSCA-RISE-2015 and SeedWheels FCT PTDC/BIA-FBT/27839/2017. AMP received a grant from the European Union&#8217;s MSCA-IF-2016 project (no. 753328). MC received a grant from FCT PhD grant SFRH/BD/111781/2015.

1. Sample Preparation: fixation, dehydration, and LR-White embedding
<ol>
	<li>Fixation and dehydration 
	NOTE: The fixation process is critical to preserve the sample; by crosslinking molecules the cellular metabolism is stopped, ensuring cellular integrity and preventing molecular diffusion. Fixative agents and the concentration used must be adjusted to that purpose, leaving the antigens sufficiently exposed to interact with the antibodies. The following protocol combines the mild fixative capability of paraformaldehyde, with the stronger effect of glutaraldehyde. Their proportions were optimized for AGPs and cell wall components, but it is suitable for other proteins and cell structures. The subsequent dehydration process will prepare the samples for LR-White embedding. Plant tissues from several plant species were used in this experiment. Quercus suber samples were collected from trees grown in the field. Trithuria submersa samples were kindly shared with us by Paula Rudall (Kew Gardens, London, UK).
	<ol>
		<li>Sow all Arabidopsis plants directly on soil and grow in an indoor growth facility with 60% relative humidity and a day/night cycle of 16 h light at 21 &#176;C and 8 h darkness at 18 &#176;C. Select the plant tissues to be analyzed, and trim samples to be no more than 16 mm2 in size.</li>
		<li>Immediately transfer the sample to a glass vial previously filled with enough cold fixative solution (2% formaldehyde (w/v), 2.5% glutaraldehyde (w/v), 25 mM PIPES pH 7 and 0.001% Tween-20 (v/v)) to completely submerge the samples (Figure 1A). 
		CAUTION: Formaldehyde and glutaraldehyde are both fixative agents that can be harmful by inhalation and contact. Perform the fixation step on a fume hood and wear adequate protective clothing and nitrile gloves. All solutions from this step onwards, including alcohol series until 70%, should be stored for later decontamination by specialized staff.</li>
		<li>After gathering all the samples, refresh the fixative.</li>
		<li>Transfer the vials to a vacuum chamber, and slowly apply vacuum. Upon reaching -60 kPa, the floating material will start to sink to the bottom of the vial (Figure 1B).</li>
		<li>Keep under a vacuum of no more than -80 kPa for 2 h at room temperature.</li>
		<li>Slowly release the vacuum, seal the glass vial and place overnight at 4 &#176;C.</li>
		<li>Discard any samples that did not sink during the overnight fixation. Wash the remaining samples with 25 mM PBS pH 7 for 10 min, followed by a 20 min wash in 25 mM PIPES buffer pH 7.2.</li>
		<li>Dehydrate the samples in an ethanol series (25%, 35%, 50%, 70%, 80%, 90%, and 3x 100% ethanol) for 20 min each. Immediately transfer the dehydrated samples to labeled glass vials for embedding (Figure 1C). 
		NOTE: The dehydration process can be paused at the 70% ethanol step.</li>
	</ol>
	</li>
	<li>LR-White resin embedding 
	NOTE: LR-white is a non-hydrophobic acrylic resin with low viscosity, making it ideal for penetrating tissues with many thick cell walls layers. LR-White also comes in several hardness grades compatible for cutting most plant samples. Please do make sure that the used resin is already supplied with the proper catalyst mixed in, or follow the supplier instructions to prepare the resin. The following embedding process is slow but the results justify greatly the means.
	<ol>
		<li>Perfuse the samples by incubating with the LR-White resin in a series of crescent concentration of resin (1:3, 2:3, 1:1, 3:2, 3:1, 1:0) in ethanol, incubating for 24 h at 4 &#176;C in each step. 
		CAUTION: LR-White resin is a low toxicity acrylic resin; however, it may be an irritant to the skin by contact and inhalation. Working in a well-ventilated area or under a fume hood with appropriate protective wear and nitrile gloves is highly recommended.</li>
		<li>Refresh the LR-white resin and incubate for an additional 12 h at 4 &#176;C.</li>
		<li>Prepare appropriate size embedding gelatin capsules (size 1 (0.5 mL) for samples up to 3 mm, 2 x 0.37 mL for samples up to 5 mm or 3 x 0.3 mL for samples up to 8 mm, and paper tags (Figure 1D). 
		NOTE: Select the capsule size to be slightly larger than the sample, so that the specimen can be completely enclosed in the resin. Also, remember to label the tags with a pencil, because pen or printed inks will contaminate the resin, ruining the sample.</li>
		<li>Apply one drop of fresh LR-white resin to the bottom of each capsule.</li>
		<li>Place a sample in each gelatin capsule and fill to maximum capacity with fresh resin. Place the capsule cap and press gently to form a hermetic seal (Figure 1E).</li>
		<li>Polymerize the resin for 24 to 48 h at 58 &#176;C, or until fully hardened.</li>
		<li>Store samples at room temperature. 
		NOTE: Post polymerization LR-white resin is inert.</li>
	</ol>
	</li>
</ol>
2. Slide preparation
NOTE: Glass slides must be clean, free of any dust, grease or any other contaminants. Even new slides must be cleaned as some suppliers use oils and detergents to prevent the slides from sticking together. Any grease or detergent will interfere with the section adhesion to the slide, even if treated with poly-L-lysine. Lint and dust will affect the specimen&#8217;s observations and very possibly ruin the experiment. Teflon coated slides with reaction wells are perfect for this task. They are affordable, reusable and drastically reduce the amount of antibody solution needed. With proper cleaning, excellent quality fluorescent immunolocalization can be performed at a very affordable cost.
<ol>
	<li>Slide washing
	<ol>
		<li>Place the slides in a staining rack and cover with cleaning solution (0.1% SDS (w/v), 1% acetic acid (v/v), 10% ethanol (v/v)).</li>
		<li>Maintain a mild agitation for 20 min.</li>
		<li>Transfer the staining racks to a ddH2O bath with mild agitation for 10 min. Repeat 4 times.</li>
		<li>Carefully drain the racks before dipping briefly in 100% ethanol and let the slides dry in a dust-free environment.</li>
		<li>Store until use.</li>
	</ol>
	</li>
	<li>Poly-L-lysine coating (optional) 
	NOTE: Small sections usually stick very well to clean glass slides. This step is only recommendable for larger sections (&#62;2 mm2). Larger sections tend to fold and wrinkle, and are not advisable. Nevertheless, if needed use reaction slides with larger holes. Clean them as above and proceed as follows.
	<ol>
		<li>Place clean slides in a square Petri dish. Pipette 0.001% poly-L-lysine solution (w/v) to cover the holes of the slides, without overflowing.</li>
		<li>Let the slides dry overnight at 40 &#176;C in closed Petri dishes. 
		NOTE: The coated slides are ready for immediate use and can be stored in a dust free environment at room temperature, for several months.</li>
	</ol>
	</li>
</ol>
3. Sample trimming and sectioning
<ol>
	<li>Sample trimming
	<ol>
		<li>With a sharp razor blade, trim the LR-White blocks under a stereomicroscope to form a pyramid shaped structure where the apex is perpendicular to the area of interest of the sample (Supplemental Figure 1A). 
		NOTE: The ultra-microtome specimen holder is an excellent tool to secure the block. If the device does not have a universal specimen holder, use the one most fit for round blocks.</li>
		<li>Trim the pyramidal structure by removing fine slices of the excess resin perpendicularly to the pyramid major axis.</li>
		<li>Proceed until the sample is reached forming the cutting surface. Within the cutting surface, the target sample should ideally be enclosed in a trapezoid shape. 
		NOTE: The resin block can be further trimmed at a slit angle to reduce the section surface area with a sharp blade (Figure 1F).</li>
	</ol>
	</li>
	<li>Semi-thin sectioning 
	NOTE: A microtome with glass knives will be used. The ability of the antibody to bind to the epitope will condition the immunolabeling reaction success. The hydrophilic nature of the LR-White resin will allow for good contact of the antibodies with the sample sections. Thinner sections will present fainter Calcofluor coloration that will be used to help locate the sections and assist with the imaging process. The same holds true for other stains that may later be applied. Also excessive thickness will affect image acquisition quality. A good compromise for section thickness can be found between 200 and 700 nm, depending on the tissue characteristics. Mount the blocks tightly on the specimen holder of the ultra-microtome.
	<ol>
		<li>With an ultra-microtome, make sections with a thickness between 200 and 700 nm (Figure 1G).</li>
		<li>Check the section area by transferring some sections to a ddH2O drop on a glass slide. Place the slide on a 50 &#176;C hot plate until water evaporates. Stain placing a drop of 1% Toluidine Blue O (w/v) in 1% boric acid solution (w/v) over the sections for 30 s. Rinse and observe under the microscope.</li>
		<li>Upon finding the desired structure, transfer one or two sections to each drop of ddH2O previously placed on each well of a clean reaction slide.</li>
		<li>Place each slide in a closed clean 10 cm square petri dish and let it dry at 50 &#176;C.</li>
		<li>Store slides in a clean archive box until use.</li>
	</ol>
	</li>
</ol>
4. Immunolocalization
NOTE: The fluorescent immunolocalization procedure relies on sequential use of two antibodies. The primary antibody is raised against a specific target antigen. The secondary antibody is raised specifically against the primary antibody and for fluorescent techniques is conjugated to a fluorophore (FITC in this specific protocol). The primary antibody will be used to detect the target antigen in the sample, and the secondary antibody will be used to mark the location where the primary antibody connected to the sample after washing off the excess of primary antibody. Controls are an important part of this assay and must always be performed to insure the accuracy of the observations. One well on the slide should be reserved for use as a negative control, where the primary antibody treatment will be skipped, and therefore no signal should be observed at the end of the experiment. A positive control must be included in the experiment by treating one well with an antibody which labelling is already known and certain. The positive control is used to confirm the secondary antibody labelling effectiveness and reaction conditions while the negative control tests the secondary antibody specificity.
<ol>
	<li>Prepare an incubation chamber by placing some damped paper towels at the bottom of a pipette tips box and wrapping it with tin foil (Supplemental Figure 1B-1E). Place the slides in the incubation chamber.</li>
	<li>Pipette a 50 &#956;L drop per 8 mm well of blocking solution (5% nonfat dry milk (w/v) in 1 M PBS) and incubate for 10 min. Remove the blocking solution and wash all wells twice with PBS for 10 min.</li>
	<li>Prepare the primary antibody solutions (see Supplemental Table 1), 1:5 (v/v) antibody in blocking solution; make an estimate of about 40 &#956;L per 8 mm well. 
	NOTE: The concentration of the antibody must be adjusted according to the manufacture&#8217;s protocol.</li>
	<li>Perform a final wash with ddH2O for 5 min, never let the wells dry completely.</li>
	<li>Pipette the primary antibody solution to the reaction wells. Pipette blocking solution to the control wells.</li>
	<li>Close the incubation chamber and let stand for 2 h at room temperature followed by overnight at 4 &#176;C. Prepare the secondary antibody solution, 1% in blocking solution (v/v) about 40 &#956;L per well. Keep it covered with tin foil.</li>
	<li>Wash all wells twice with PBS for 10 min, followed by 10 additional minutes with ddH2O. Make sure that no trace of the blocking solution or deposits is visible on the wells.</li>
	<li>Pipette the secondary antibody solution to all wells. From now on, protect the slides from light.</li>
	<li>Incubate for 3&#8211;4 h in the dark at room temperature.</li>
	<li>Wash all wells twice for 10 min with PBS followed by another wash of 10 min with ddH2O.</li>
	<li>Apply a drop of calcofluor (1:10,000 (w/v) fluorescent brighter 28 in PBS) to each well. Without washing, apply a drop of mounting medium (see Table of Materials) to each well and place a coverslip (Figure 1H).</li>
	<li>Observe with a fluorescence microscope, equipped with 10x/0.45, 20x/0.75, 40x/0.95 and 100x/1.40 lens, use UV (for calcofluor stain) and FITC filters, to detect cell wall and immunolocalization respectively (Figure 1I). Use the following wavelengths: excitation/emission (nm) 358/461 for UV and 485/530 for FITC.</li>
	<li>For a better visualization of the results overlap both images with ImageJ or similar.</li>
</ol>

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The authors declare no conflicts of interest.

The fluorescent immunolocalization method in plants here described, while seamlessly straightforward, relies on the success of several small steps. The first of which is sample preparation and fixation. During this first step, a mixture of formaldehyde and glutaraldehyde is used to crosslink the majority of the cell components. The formaldehyde in the solution provides a mild and reversible fixation while the glutaraldehyde provides a strong more permanent linkage; the balance between the two fixatives provides the appropriate amount of crosslinking, allowing exposure of the epitopes to react with the primary antibody10. For the fixative solution to work properly, the sample must be small. Large structures more than 7 mm in diameter are very difficult to fix and should be clipped to ensure fixative penetration.
After wounding or stress, some plants secrete large amounts of tannins forming dark precipitates that may react with the formaldehyde interfering with the fixative process. Keeping the samples on ice and replacing the fixative solution whenever it becomes cloudy helps to reduce this effect. Air can also interfere with the fixative process and later on in the embedding and hardening of the resin. The proposed vacuum treatment should remove most of the air. For particularly airy tissues, the vacuum step may be extended further than 2 h, until no more air comes out of the samples and they sink to the bottom. Any sample found floating after the overnight fixative step should be discarded. Resin embedding provides support for cutting the preserved sample, and its hardness should be approximate to the embedded tissues10. LR-White comes in three hardness grades: hard for woody heavily sclerified tissues, medium grade for the vast majority of tissues from leaves to pollen grains, and soft grade for more delicate tissues. For a perfect embedding, the resin must completely penetrate the sample; this is better obtained with a slow and gradual infiltration. With prior testing, shorter incubation periods may be used. The gelatin capsules are both small and hermetically sealable, which makes them perfect for LR-White resin curing. For a size 2 (0.37 mL), curing should be complete after 24 h at 58 &#176;C. For other capsule sizes, curing time should be adjusted. The LR-White cured resin should go from almost colorless to a light golden/amber color and feel hard to the nails. Sample trimming and the use of the ultra-microtome requires practice but will produce clear figures. The thickness and size of the section is important for the imaging and immunolocalization assay. The section thickness should be kept around 500 nm. Sections that are too thin will result in poor staining and section thicknesses over 700 nm will interfere with the image acquisition and resolution. The hydrophilic nature of LR-White resin allows for direct use of the sections in staining and immunolocalization without removing the resin.
The blocking solution (5% nonfat dry milk in 1 M PBS) reduces unspecific binding of antibodies. Nonfat dry milk has been a reliable alternative to BSA or other more traditional blocking agents, and is significantly less expensive. The blocking solution must be filtered through a paper filter prior to use, to avoid precipitates that form hard to wash away aggregates, retain antibodies, and compromise the experiment. The proposed antibody ratios and incubation times have been optimized and successfully applied to diverse species in several studies9,11,12,13,14,15,16.
Evaporation and exposition to light are two issues that must be avoided during the immunolocalization procedure. A humid and dark incubation chamber solves both of these problems. A tutorial on how to make a dark chamber can be found in Supplemental Figure 1B-1E. This immunolocalization protocol calls for the use of a primary antibody directed to the target epitope with no label of its own, and a secondary antibody conjugated to a fluorochrome (FITC) that is raised against the primary antibody IgG. This method offers several advantages over the use of a single antibody detection system due to an increased stringency of the detection, as the secondary antibody has no affinity to the target sample species. This system offers increased signal as the primary antibody molecules may be targeted by several molecules of the secondary antibody, each carrying a fluorochrome17.
The decay of fluorochromes by intense light, or photo bleaching, is an important issue in this technique. Especially when exploring for faint localized signals, the signal may become irreversibly lost before imaging. Mounting medium greatly increases the stability and lifespan of the fluorochromes18; however, it is highly advisable to test the mounting media, as some may react with the stain and/or the fluorochrome, forming precipitates and blurring the image. The configuration of the optical system used for observing and registering the immunolocalization is one of the most important aspects for the success of this experiment. Due to the sections&#8217; reduced thickness, the benefits of using the confocal microscope are limited. The most successful setup requires a conventional upright epifluorescence microscope equipped with a fluorescent light source, LED or mercury light bulb19, with adequate fluorescence grade objectives. Also good quality light filters set for excitation/emission (nm) 358/461 for UV and 485/530 for FITC are required. For the fluorescence image acquisition, refrigerated monochromatic digital cameras are recommended due to their high speed and sensitivity, but sacrifice true color information20. Polychromatic cameras provide the ability of easily sort out signal from background fluorescence but are slower and far less sensitive than their monochromatic counterparts.
The method is limited by the availability of specific antibodies. Despite the availability of a vast and expanding collection of antibodies aimed at plants epitopes, understandably not all plant compounds are yet covered. Also lipids tend to be extracted by the described embedding method, and is therefore not recommended for tissues with a high oil content. Cryostat section may be an alternative, despite sacrificing image resolution. The temperature of LR-White resin polymerization may in some cases alter more sensitive epitopes. If the target epitope is temperature sensitive, switch LR-White for LR-Gold resin. Polymerizing at -25 &#176;C under white light, LR-Gold offers an excellent preservation of the thermosensitive epitopes, but will require the acquisition of a specialized polymerization apparatus and is slightly more toxic then LR-White.
This method has been used across a broad range of species and tissues. It allows fast access to reliable information on specific epitope distribution. Offering supporting data for evolutionary models and identifying markers and specific adaptations in cells and tissues while providing visually enticing results. The use of antibodies conjugated with fluorochromes over peroxidase or alkaline phosphatase conjugation alternatives10, is less time consuming, significantly less prone to overreaction artefacts and provides a clear subcellular resolution hard to match with any other technique. The use of antibodies specific to cell wall related polymers allows an insight into the arrangement of compounds into very restrict domains. Such resolution would be challenging to obtain from traditional biochemical tools. The ever-expanding set of primary antibodies available offers constant new discovery opportunities into the plant cell inner workings.

Plant cell walls are complex structures composed of polysaccharides and glycoproteins. Cell walls are extremely dynamic structures whose architecture, organization and composition vary according to cell type, localization, developmental stage, external and internal stimuli1. Arabinogalactan proteins (AGPs) and pectins are important components of the plant cell wall. AGPs are highly glycosylated proteins and pectins are homogalacturonan polysaccharides whose composition, amount and structure vary greatly during different plant developmental stages2,3,4. AGPs and pectin studies have revealed their involvement in several plant processes such as programmed cell death, response to abiotic stresses, sexual plant reproduction, among many others5. Most of these studies started with information obtained from immunolocalization studies.
Given its complexity, the study of cell walls requires many different tools. Detection of glycan epitopes using monoclonal antibodies (mAbs) is a valuable approach to resolve polysaccharide and glycoprotein distribution along this structure. There is a large collection of mAbs available to detect glycan epitopes and the specificity of each mAb is continuously being improved as well6. The technique here described is applicable to all plant species, and is a perfect tool to guide future research directions that might involve more expensive and complex techniques.
In this technique, specific antibodies are chemically conjugated to fluorescent dyes such as FITC (fluorescein isothiocyanate), TRITC (tetramethylrhodamine-5-(and 6)-isothiocyanate) or several Alexa Fluor dyes. Immunofluorescence offers several advantages, allowing a clear and quick subcellular localization of glycans that can be directly observed under a fluorescence microscope. It is highly specific and sensitive, since the preparation of the sample can effectively protect the natural structure of the antigen, even if present in lower amounts. It allows the detection of multiple antigens in the same sample and most important, offers high quality and visually beautiful results. Despite the great power offered by fluorescence immunolocalization studies, they are often regarded as difficult to perform and implement most probably due to the lack of detailed protocols allowing the visualization of the different steps of the procedure. Here, we provide some simple guidelines on how to perform this technique and how to obtain high quality images.
For the protocol presented here, samples must first be fixed and embedded using the most appropriate fixative. Although considered as a time consuming and relatively tedious technique, proper fixation and embedding of the plant sample is the key to ensure a successful immunolocalization assay. For this purpose, the most usual is chemical fixation using crosslinking fixatives, like aldehydes. Cross-linking fixatives establish chemical bonds between molecules of the tissue, stabilizing and hardening the sample. Formaldehyde and glutaraldehyde are cross-linking fixatives, and sometimes a mix of both fixatives is used7. Formaldehyde offers great structural preservation of tissues and for extended periods of time, producing small tissue retractions and being compatible with immunostaining. Glutaraldehyde is a stronger and stable fixative usually used in combination with formaldehyde. The use of glutaraldehyde has some disadvantages that must be taken into account as it introduces some free aldehyde groups into the fixed tissue, which may generate some unspecific labeling. Also the crosslinking between proteins and other molecules occasionally may render some target epitopes inaccessible for the antibodies. To avoid this, the quantity and duration of the fixation must be carefully defined.
After fixation, samples are embedded in the proper resin to harden before obtaining the sections. London Resin (LR-White) acrylic resin is the resin of choice for immunolocalization studies. Unlike other resins, LR-White is hydrophilic, allowing the antibodies to reach their antigens, with no need of any treatment to facilitate it. LR-White has also the advantage of offering low auto-fluorescence, allowing a reduction in background noise during immunofluorescence imaging.
There are many staining techniques available to detect different components of the cell wall, such as Alcian blue staining, toluidine blue staining or Periodic acid&#8211;Schiff (PAS) staining. None of these offers the power of immunolocalization analyses8. This approach gives greater specificity in the detection of glycans, offering vaster information regarding cell wall composition and structure.

<table><tbody><tr><td>25% (w/v) Gluteraldehyde</td><td>Agar Scientific</td><td>AGR1010</td><td>aq. Solution, methanol free</td></tr><tr><td>8 wells Glass reaction slides</td><td>Marinfeld</td><td>MARI1216750</td><td>other brands may be used</td></tr><tr><td>Acetic acid</td><td>Sigma-Aldrich</td><td>A6283</td><td></td></tr><tr><td>Anti-Rat IgG (wole molecule)-FITC antibody produced in GOAT</td><td>Sigma-Aldrich</td><td>F6258</td><td></td></tr><tr><td>cover slips, 24 mm x 50 mm</td><td>Marinfeld</td><td>MARI0100222</td><td>The cover slip should cover all the wells. Other brands may be used&amp;nbsp;</td></tr><tr><td>ddH&lt;sub&gt;2&lt;/sub&gt;O</td><td>na</td><td>na</td><td></td></tr><tr><td>Absolute ethanol</td><td>na</td><td>na</td><td></td></tr><tr><td>Fluorescent brightner 28</td><td>Sigma-Aldrich</td><td>F-6259</td><td></td></tr><tr><td>Gelatin capsules</td><td>Agar scientific</td><td>AGG29211</td><td>The capsule size should fit the size of the sample.</td></tr><tr><td>Glass vials</td><td>na</td><td>na</td><td>Any simple unexpensive glass vials that can be sealed, may be used. The vials may be cleaned with 96% ethanol after use to remove LR-White residue and reused.&amp;nbsp;</td></tr><tr><td>LR-white medium grade, embdeding resin</td><td>Agar Scientific</td><td>AGR1281</td><td>LR-White comes in several forms; the medium grade provides an adequate cutting support for most tissues, while for harder tissues a harder grade of LR-white may be recommendable. If possible use a resin already mixed with the polimeration activator (benzoyl peroxide), if not please follow the instructions of the supplier to prepare the resin.</td></tr><tr><td>Non fat dry milk</td><td>Nestl&amp;eacute;</td><td>na</td><td>any non fat dry milk is adequate</td></tr><tr><td>Oven</td><td>na</td><td>na</td><td>generic laboratory oven</td></tr><tr><td>Petri dish, 10 cm x 10 cm square</td><td>na</td><td>na</td><td></td></tr><tr><td>PIPES</td><td>Sigma Aldrich</td><td>P1851</td><td></td></tr><tr><td>Rat generated Monoclonal Anti-Body</td><td>Plant probes</td><td>na</td><td>Several antibodies that recognize cell wall components are&lt;br/&gt; available at both the Complex Carbohydrate research center&lt;br/&gt; (CCRC, Georgia University USA) and Plant Probes (Paul&lt;br/&gt; Knox Cell Wall Lab, at Leeds University UK). A short list of&lt;br/&gt; some commonly used MABS and where they can be purchased&lt;br/&gt; is presented in Supplemental Table 1</td></tr><tr><td>Razor blades</td><td>na</td><td>na</td><td>regular razor blades</td></tr><tr><td>SDS</td><td>Sigma-Aldrich</td><td>L6026</td><td></td></tr><tr><td>Toluidine Blue-O</td><td>Agar Scientific</td><td>AGR1727</td><td></td></tr><tr><td>Tween 20</td><td>Sigma-Aldrich</td><td>P9416</td><td></td></tr><tr><td>Ultramicrotome</td><td>Leica Microsystems</td><td>UC7</td><td></td></tr><tr><td>Upright epifluorescence microscope with UV and FITC fluorescence filters</td><td>Leica Mycrosistems</td><td>DMLb</td><td></td></tr><tr><td>Vacuum chamber</td><td>na</td><td>na</td><td></td></tr><tr><td>Vacuum pump</td><td>na</td><td>na</td><td></td></tr><tr><td>Vectashield</td><td>vecta Labs</td><td>T-1000</td><td>Other anti-fade may be used. Please do check for compatibility with FITC and the Fluorescente brightner 28. (Note: for a non-commercial alternative, (see Jonhson et al 1982&lt;sup&gt;18&lt;/sup&gt;) An anti-fade medium can be made by mixing 25 mg/mL of 1,4-Diazobicyclo-(2,2,2)octane (DABCO) in 9:1 (v/v) glycerol to 1xPBS. Adjust pH to 8.6 with diluted HCl.)</td></tr></tbody></table>

fluorescent immunolocalization of arabinogalactan proteins and pectins in the cell wall of plant tissues

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls are composed of cellulose and non-cellulosic polysaccharides together with proteins, phenolic compounds and water. 90% of the cell wall is composed of&#160;polysaccharides (e.g., pectins) and arabinogalactan proteins (AGPs). The fluorescent immunolocalization of specific glycan epitopes in plant histological sections remains a key tool to uncover remodeling of wall polysaccharide networks, structure and components.
Here, we report an optimized fluorescent immunolocalization procedure to detect glycan epitopes from AGPs and pectins in plant tissues. Paraformaldehyde/glutaraldehyde fixation was used along with LR-White embedding of the plant samples, allowing for a better preservation of the tissue structure and composition. Thin sections of the embedded samples obtained with an ultra-microtome were used for immunolocalization with specific antibodies. This technique offers great resolution, high specificity, and the chance to detect multiple glycan epitopes in the same sample. This technique allows subcellular localization of glycans and detects their level of accumulation in the cell wall. It also permits the determination of spatio-temporal patterns of AGP and pectin distribution during developmental processes. The use of this tool may ultimately guide research directions and link glycans to specific functions in plants. Furthermore, the information obtained can complement biochemical and gene expression studies.

In a successful experiment, the secondary antibody will specifically pinpoint the location of the specific epitope in bright green, in a consistent manner, allowing for the characterization of the cell wall composition at a certain development stage of the cell, tissue or organ. For example the LM6 antibody has an high affinity for 1,5-arabinan, a compound with type-I rhamnogalacturonan that can be found abundantly labelling the cell wall of the developing Quercus suber anther (Figure 2A), thus allowing to conclude that this type of pectin is abundant and part of the primary cell wall composition. JIM5 has affinity for homogalacturonans scarcely esterified that are typically found at the root tip of Quercus suber embryo, specifying mechanical properties of the organ (Figure 2B). Xylogalacturonan are a type of pectin rich in xylose associated with cell wall loosening, they are found in degenerating cells. The antibody LM8 specifically recognizes xylogalacturonans, in maturing organs it may be used to detect degenerating cells or tissues, like the endosperm cells during the final stages of the Quercus suber acorn maturation (Figure 2C).
JIM13 has affinity for AGPs found on structures related with reproduction, cell lines related to microgametogenesis in Arabidopsis thaliana (Figure 2D). JIM8 also recognizes epitopes of AGPs present in cells and organs related to reproduction like the stigmatic papillae and micropyle of the Basal Angiosperm Trithuria submersa (Figure 2E-2F). Both antibodies have been suggested to be molecular markers for the gametophytic cell lines in plants9.
Common mistakes to this protocol are normally easy to detect and identify. When the washes are skipped or the reaction wells are let to dry, the secondary antibody will usually appear as an unspecific smear covering indiscriminately over cells, tissues, resin and slide (Figure 3A). Aggregates of green fluorochrome will form if the unbounded primary antibody has not been properly washed away (Figure 3B). Another common cause for failure of this technique is related with the folding and/or detachment of the sections (Figure 3C), rendering the experiment useless. This problem is usually related with either poor adhesion of the sections to the slides, probably due to the use of unclean slides, and/or aggressive washing.
The sample preparation is also a critical step in this procedure. Fortunately, the most common fixative and embedding issues are easy to spot (Figure 3D). If all goes well the resin block will be free of cracks and the sample will be clearly visible with a pale yellow to light brown color (Figure 3D-1). Samples inefficiently embedded will show powdery white spots or areas (Figure 3D-2). Keeping the sample size under 8 mm is important to guarantee the penetration of the fixative solution. When samples are poorly fixed, they will appear dark brown almost black (Figure 3D-3). Also the temperature of polymerization is important for both the preservation of the epitopes and proper hardening of the resin. An excessive temperature can cause the resin to crack making the sectioning of the sample impossible (Figure 3D-4).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/61034/61034fig1.jpg" /> 
Figure 1. Overview of the complete protocol. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61034/61034fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/61034/61034fig2.jpg" /> 
Figure 2: Typical results of AGPs and Pectin immunolocalization in plant samples.&#160;(A) LM6 labeling of arabinan moiety of pectins in a Quercus suber anther in meiosis I. (B) Specific labeling of low methyl-esterified pectins in a Quercus suber embryo root tip by JIM5. (C) Xylogalacturonan labeled by LM8 in the receding endosperm of a Quercus suber maturing acorn. (D) JIM13 labeling of AGPs in the tapetum and tetrads of an Arabidopsis thaliana anther. (E) AGPs labeled by JIM8 in the stigmatic papillae of Trithuria submersa.&#160;(F) JIM8 labelling AGPs at the micropyle (white arrow) of a Trithuria submersa ovule. Meiotic microspore (Mc), Tapetum (Tp), Root (R), Root tip (Rt), Testa (Ts), Endosperm (Ed), Embryo (Em), Epidermis (Ep), Stigmatic papillae (SP), Ovule (OV). Scale bars: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61034/61034fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/61034/61034fig3.jpg" /> 
Figure 3: Common mistakes and problems during the protocol.&#160;(A) Failure of wash steps are identified by the presence of unspecific smear of secondary antibody (+) over sample resin. (B) The poor specificity or wash failure of the primary antibody results in the formation of FITC aggregates (+) not bonded to a specific location. (C) Folds and section detachment are often the result of aggressive wash and unclean slides. (D) Examples of sample fixation and embedding; (1) perfect sample size and embedding, (2) embedding failure, (3) sample fixation failure, (4) resin hardening failure. FITC labeled antibody (green) and Calcofluor-white stain (blue). Scale bars: 100 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61034/61034fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure 1. (A) Schematic representation of the block trimming sequence for preparing the sample (Yellow block) for sectioning with the ultramicrotome. Firstly, the gelatin capsule is removed (1). Then they are first trimmed at a 40&#176; to 45&#176; angle to the sides of the capsule tangentially to the sample (2), a second cut is made at 90&#176; of the first (3) followed by a third (4) and fourth following the same rule (5). From this first phase of trimming results a pyramidal shaped structure which summit is located above the sample. Finally, with a sharp blade the summit is shaved off perpendicularly to the resin block major axis to reach the embedded sample. A square or trapezoid surface should be obtained at the end of the procedure (6). (B) Required supplies to make an incubation chamber; tin foil (1), double sided duct tape (2), paper towels (3) and a pipette tip box. (C) First step, the tin foil is fixed to the box with the double-sided duct tape. (D) Second step, the pipette tips holder is removed to place paper towels at the bottom of the box. (E) Third step, the paper towels are damped with water and the pipette tips holder is placed back to act has a tray for the slides. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61034/61034supfig1large.jpg" target="_blank">Please click here to download this file.</a>
Supplemental Table 1: List of useful monoclonal antibodies. The above table represents an example of available antibodies, with information about their targets and where they can be purchased. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/61034/Supplemental_Table_1.xlsx" target="_blank">Please click here to download this file.</a>

Watch this Scientific Journal Video about Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues at JoVE.com

Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

This protocol describes in detail how the plant material for immunolocalization of Arabinogalactan proteins and pectins is fixed, embedded in a hydrophilic acrylic resin, sectioned and mounted on glass slides. We show cell wall related epitopes will be detected with specific antibodies.

Plant development involves constant adjustments of the cell wall composition and structure in response to both internal and external stimuli. Cell walls ...

fluorescent-immunolocalization-arabinogalactan-proteins-pectins-cell

Nanyang Technological University

Research

JoVE Journal

Developmental Biology

6.1K Views.  Faculdade de Ciências da Universidade do Porto. This protocol describes in detail how the plant material for immunolocalization of Arabinogalactan proteins and pectins is fixed, embedded in a hydrophilic acrylic resin, sectioned and mounted on glass slides. We show cell wall related epitopes will be detected with specific antibodies.

Video: Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

Fluorescence Activated Cell Sorting of Plant Protoplasts

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental stimuli in any multicellular organism. In situ hybridization and reporter gene visualization can to a limited extent be used to this end but for high resolution quantitative RT-PCR or high-throughput transcriptome-wide analysis the isolation of RNA from particular cell types is requisite. Cellular dissociation of tissue expressing a fluorescent protein marker in a specific cell type and subsequent Fluorescence Activated Cell Sorting (FACS) makes it possible to collect sufficient amounts of material for RNA extraction, cDNA synthesis/amplification and microarray analysis.
An extensive set of cell type-specific fluorescent reporter lines is available to the plant research community. In this case, two marker lines of the Arabidopsis thaliana root are used: PSCR::GFP (endodermis and quiescent center) and PWOX5::GFP (quiescent center). Large numbers (thousands) of seedlings are grown hydroponically or on agar plates and harvested to obtain enough root material for further analysis. Cellular dissociation of plant material is achieved by enzymatic digestion of the cell wall. This procedure makes use of high osmolarity-induced plasmolysis and commercially available cellulases, pectinases and hemicellulases to release protoplasts into solution.
FACS of GFP-positive cells makes use of the visualization of the green versus the red emission spectra of protoplasts excited by a 488 nm laser. GFP-positive protoplasts can be distinguished by their increased ratio of green to red emission. Protoplasts are typically sorted directly into RNA extraction buffer and stored for further processing at a later time.
This technique is revealed to be straightforward and practicable. Furthermore, it is shown that it can be used without difficulty to isolate sufficient numbers of cells for transcriptome analysis, even for very scarce cell types (e.g. quiescent center cells). Lastly, a growth setup for Arabidopsis seedlings is demonstrated that enables uncomplicated treatment of the plants prior to cell sorting (e.g. for the cell type-specific analysis of biotic or abiotic stress responses). Potential supplementary uses for FACS of plant protoplasts are discussed.

A method for isolating specific cell types from plant material is demonstrated. This technique employs transgenic marker lines expressing fluorescent proteins in particular cell types, cellular dissociation and Fluorescence Activated Cell Sorting. Additionally, a growth setup is established here that facilitates treatment of Arabidopsis thaliana seedlings prior to cell sorting.

High-resolution, cell type-specific analysis of gene expression greatly enhances understanding of developmental regulation and responses to environmental ...

Biology

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

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

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

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

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

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

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

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

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

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

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation (BiFC) System

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein (YFP) into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest. These constructs can then be co-transformed into Nicotiana benthamiana via Agrobacterium mediated transformation, allowing the transit expression of fusion proteins. The reconstitution of YFP signal only occurs when the inquest proteins interact 1-7. To test and validate the protein-protein interactions, BiFC can be used together with yeast two hybrid (Y2H) assay. This may detect indirect interactions which can be overlooked in the Y2H. Gateway technology is a universal platform that enables researchers to shuttle the gene of interest (GOI) into as many expression and functional analysis systems as possible8,9. Both the orientation and reading frame can be maintained without using restriction enzymes or ligation to make expression-ready clones. As a result, one can eliminate all the re-sequencing steps to ensure consistent results throughout the experiments. We have created a series of Gateway compatible BiFC and Y2H vectors which provide researchers with easy-to-use tools to perform both BiFC and Y2H assays10. Here, we demonstrate the ease of using our BiFC system to test protein-protein interactions in N. benthamiana plants.

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation BiFC System

We have developed a technique to test protein-protein interactions in plant. A yellow fluorescent protein (YFP) is split into two non-overlapping fragments. Each fragment is cloned in-frame to a gene of interest via Gateway system, enabling expression of fusion proteins. Reconstitution of YFP signal only occurs when the inquest proteins interact.

We have developed a BiFC technique to test the interaction between two proteins in vivo. This is accomplished by splitting a yellow fluorescent protein ...

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 ...

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

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

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

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

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

Application of Membrane and Cell Wall Selective Fluorescent Dyes for Live-Cell Imaging of Filamentous Fungi

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two decades ago and since then continues to contribute greatly to our understanding of the filamentous fungal lifestyle. This paper provides a practical guide for the utilization of the two membrane dyes FM 1-43 and FM 4-64 and the four cell wall stains Calcofluor White M2R, Solophenyl Flavine 7GFE 500, Pontamine Fast Scarlet 48 and Congo Red. The focus is on their low-dose application to ascertain artefact-free staining, their co-imaging properties, and their quantitative evaluation. The presented methods are applicable to all filamentous fungal samples that can be prepared in the described ways. The fundamental staining approaches can serve as starting points for adaptations to species that might require different cultivation conditions. First, biophysical and biochemical properties are reviewed as their understanding is essential for using these dyes as truly vital fluorescent stains. Secondly, step-by-step protocols are presented that detail the preparation of various fungal sample types for fluorescent live-cell imaging. Finally, example experiments illustrate different approaches to: (1) identify defects in the spatio-temporal organization of endocytosis in genetic mutants, (2) comparatively characterize shared and distinct co-localization of GFP-labeled target proteins in the endocytic pathway, (3) identify morphogenetic cell wall defects in a genetic mutant, and (4) monitor cell wall biogenesis in real time.

Vital fluorescent dyes are essential tools for live-cell imaging analyses in modern fungal cell biology. This paper details the application of established and lesser-known fluorescent dyes for tracking plasma membrane dynamics, endo-/exocytosis and cell wall morphogenesis in filamentous fungi.

The application of membrane and cell wall selective fluorescent dyes for live-cell imaging analyses of organelle dynamics in fungal cells started two ...

Double-Staining Method to Detect Pectin in Plant-Fungus Interaction

Plant cells use different structural mechanisms, either constitutive or inducible, to defend themselves from fungal infection. Encapsulation is an efficient inducible mechanism to isolate the fungal haustoria from the plant cell protoplast. Conversely, pectin, one of the polymeric components of the cell wall, is a target of several pectolytic enzymes in necrotrophic interactions. Here, a protocol to detect pectin and fungal hyphae through optical microscopy is presented. The pectin-rich encapsulation in the cells of coffee leaves infected by the rust fungus Hemileia vastatrix and the mesophyll cell wall modification induced by Cercospora coffeicola are investigated. Lesioned leaf samples were fixed with the Karnovsky solution, dehydrated, and embedded in glycol methacrylate for 2-4 days. All steps were followed by vacuum-pumping to remove air in the intercellular spaces and improve the embedding process. The embedded blocks were sectioned into 5-7 &#181;m thick sections, which were deposited on a glass slide covered with water and subsequently heated at 40 &#176;C for 30 min. Next, the slides were double-stained with 5% cotton blue in lactophenol to detect the fungus and 0.05% ruthenium red in water to detect pectin (acidic groups of polyuronic acids of pectin). Fungal haustoria of Hemileia vastatrix were found to be encapsulated by pectin. In coffee cercosporiosis, mesophyll cells exhibited dissolution of cell walls, and intercellular hyphae and conidiophores were observed. The method presented here is effective to detect a pectin-associated response in the plant-fungi interaction.

This protocol describes a microscopical method to detect pectin in coffee-fungus interaction.

Plant cells use different structural mechanisms, either constitutive or inducible, to defend themselves from fungal infection. Encapsulation is an ...

Experimental Screening Protocols, Immunocytochemistry and Microscopy-based Imaging Techniques for Penium margaritaceum

The cell wall is the first component of signal reception/transduction for a plant cell during development and when responding to environmental abiotic and biotic stressors. The cell constantly monitors the integrity of its cell wall and modulates it in response to stress. Elucidating the specific structural and biochemical modulations occurring in the cell wall is a difficult task especially when employing multicellular plants and their organs/tissues. This is due to limits as to what can be resolved in an individual cell that is part of a complex multicellular network. The unicellular streptophyte alga, Penium margaritaceum, has recently been used in investigations of pectin dynamics, cell wall-based phenotypic plasticity and multiple aspects of algal cell biology. Its simple phenotype, distinct cell wall that has many components notably similar to land plant cell walls, and ease in immunocytochemical and experimental studies make it a powerful model organism in plant cell wall biology. The goal of this study is to provide the basic techniques for culturing, experimental manipulation and screening of applied stressors. Screening protocols for immunocytochemistry, confocal laser scanning microscopy imaging and scanning electron microscopy imaging&#160;of cell wall structure. Likewise, many of the described techniques may be modified for a wide array of other cell and molecular studies.

This report describes the fundamental methods used to culture and experimentally manipulate the unicellular streptophyte alga, Penium margaritaceum. It also provides fundamental protocols of microscopy-based imaging, including live cell labeling with monoclonal antibodies and other fluorescent probes and scanning electron microscopy.

The cell wall is the first component of signal reception/transduction for a plant cell during development and when responding to environmental abiotic and ...

Calcofluor White Staining: A Simple Method for UV-Induced Fluorescence Visualization of Cellulose Deposits in Cell Walls of Dicot Plant Stem Sections

Source: Pradhan M. et al. <a href="https://www-jove-com.remotexs.ntu.edu.sg/v/51381">Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements.</a> J. Vis. Exp. (2014)
In this video, we describe a staining procedure to visualize cellulose deposition within cell walls of Arabidopsis thaliana, a dicot plant, using calcofluor white, which specifically binds to &#946;-linked glucans, exhibiting fluorescence under UV light.

Source: Pradhan M. et al. Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements. J. Vis. Exp. (2014)
In this video, we describe a ...

Fluorescent Immunolocalization of Arabinogalactan Proteins and Pectins in the Cell Wall of Plant Tissues

Summary

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Chapters in this video

Fluorescence Activated Cell Sorting of Plant Protoplasts

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

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

Detection of Protein Interactions in Plant using a Gateway Compatible Bimolecular Fluorescence Complementation (BiFC) System

Cell Specific Analysis of Arabidopsis Leaves Using Fluorescence Activated Cell Sorting

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

Application of Membrane and Cell Wall Selective Fluorescent Dyes for Live-Cell Imaging of Filamentous Fungi

Double-Staining Method to Detect Pectin in Plant-Fungus Interaction

Experimental Screening Protocols, Immunocytochemistry and Microscopy-based Imaging Techniques for Penium margaritaceum

Calcofluor White Staining: A Simple Method for UV-Induced Fluorescence Visualization of Cellulose Deposits in Cell Walls of Dicot Plant Stem Sections