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 appro...

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 border="0" cellpadding="0" cellspacing="0" style="width:728px;" width="728">
	<colgroup>
		<col />
		<col />
		<col />
		<col />
	</colgroup>
	<tbody>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">25% (w/v) Gluteraldehyde</td>
			<td style="width:99px;">Agar Scientific</td>
			<td style="width:196px;">AGR1010</td>
			<td style="width:187px;">aq. Solution, methanol free</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">8 wells Glass reaction slides</td>
			<td style="width:99px;">Marinfeld</td>
			<td style="width:196px;">MARI1216750</td>
			<td style="width:187px;">other brands may be used</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Acetic acid</td>
			<td style="width:99px;">Sigma-Aldrich</td>
			<td style="width:196px;">A6283</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Anti-Rat IgG (wole molecule)-FITC antibody produce in GOAT</td>
			<td style="width:99px;">Sigma-Aldrich</td>
			<td style="width:196px;">F6258</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:247px;">cover slips, 24 mm x 50 mm</td>
			<td style="width:99px;">Marinfeld</td>
			<td>MARI0100222</td>
			<td style="width:187px;">The cover slip should cover all the wells. Other brands may be used&#160;</td>
		</tr>
		<tr height="25">
			<td height="25" style="height:25px;width:247px;">ddH2O</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Ethanol absolute</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Fluorescent brightner 28</td>
			<td style="width:99px;">Sigma-Aldrich</td>
			<td style="width:196px;">F-6259</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Gelatin capsules</td>
			<td style="width:99px;">Agar scientific</td>
			<td>AGG29211</td>
			<td style="width:187px;">The capsule size sould feat the size of the sample.</td>
		</tr>
		<tr height="147">
			<td height="147" style="height:147px;width:247px;">Glass vials</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;">Any simple unexpensive glass vials that can be sealed, may be used. The vials may be clean with 96% etanol after use to remove LR-White residue and reused.&#160;</td>
		</tr>
		<tr height="294">
			<td height="294" style="height:294px;width:247px;">LR-white medium grade, embdeding resin</td>
			<td style="width:99px;">Agar Scientific</td>
			<td>AGR1281</td>
			<td style="width:187px;">LR-White comes in several forms the medium grade provides na adequate cutting suport for most tissues for harder tissues a harder grade of LR-white may be recomendable. If possible use a resin already mixed with the polimeration activator (benzoyl peroxide), if not please folow the instructions of the suplier to prepare the resin.</td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Non fat dry milk</td>
			<td style="width:99px;">Nestl&#233;</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;">any non fat dry milk is adequate</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Oven</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;">generic laboratory oven</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Petri dish, 10 cm x 10 cm square</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">PIPES</td>
			<td style="width:99px;">Sigma Aldrich</td>
			<td style="width:196px;">P1851</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="357">
			<td height="357" style="height:357px;width:247px;">Rat generated Monoclonal Anti-Body</td>
			<td style="width:99px;">Plant probes</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;">Several antibodies that recognize cell wall components are 
			available at both the Complex Carbohydrate research center 
			(CCRC, Georgia University USA) and Plant Probes (Paul 
			Knox Cell Wall Lab, at Leeds University UK). A short list of 
			some commonly used MABS and where they can be purchased 
			is presented in Supplemental Table 1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Razor blades</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;">regular razor blades</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">SDS</td>
			<td style="width:99px;">Sigma-Aldrich</td>
			<td style="width:196px;">L6026</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Toluidine Blue-O</td>
			<td style="width:99px;">Agar Scientific</td>
			<td>AGR1727</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">Tween 20</td>
			<td style="width:99px;">Sigma-Aldrich</td>
			<td style="width:196px;">P9416</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="42">
			<td height="42" style="height:42px;width:247px;">Ultramicrotome</td>
			<td style="width:99px;">Leica Microsystems</td>
			<td style="width:196px;">UC7</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="63">
			<td height="63" style="height:63px;width:247px;">upright epifluorescence microscope with UV and FITC fluorescence filters</td>
			<td style="width:99px;">Leica Mycrosistems</td>
			<td style="width:196px;">DMLb</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">vaccum chamber</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:247px;">vaccum pump</td>
			<td style="width:99px;">na</td>
			<td style="width:196px;">na</td>
			<td style="width:187px;"></td>
		</tr>
		<tr height="297">
			<td height="297" style="height:297px;width:247px;">Vectashield</td>
			<td style="width:99px;">vecta Labs</td>
			<td style="width:196px;">T-1000</td>
			<td style="width:187px;">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 198218) An antifade 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</st...

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

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

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the cephalochordate amphioxus, Branchiostoma lanceolatum. We use constructs for membrane and nuclear targeted mCherry and eGFP that have been modified to accommodate amphioxus codon usage and Kozak consensus sequences. We describe the type of injection needles to be used, the immobilization protocol for the unfertilized oocytes, and the overall injection set-up. This technique generates fluorescently labeled embryos, in which the dynamics of cell behaviors during early development can be analyzed using the latest in vivo imaging strategies. The development of a microinjection technique in this amphioxus species will allow live imaging analyses of cell behaviors in the embryo as well as gene-specific manipulations, including gene overexpression and knockdown. Altogether, this protocol will further consolidate the basal chordate amphioxus as an animal model for addressing questions related to the mechanisms of embryonic development and, more importantly, to their evolution.

Expression of Fluorescent Proteins in Branchiostoma lanceolatum by mRNA Injection into Unfertilized Oocytes

We report here the robust and efficient expression of fluorescent proteins after mRNA injection into unfertilized oocytes of Branchiostoma lanceolatum. The development of the microinjection technique in this basal chordate will pave the way for far-reaching technical innovations in this emerging model system, including in vivo imaging and gene-specific manipulations.

We report here a robust and efficient protocol for the expression of fluorescent proteins after mRNA injection into unfertilized oocytes of the ...

A Novel Mammary Fat Pad Transplantation Technique to Visualize the Vessel Generation of Vascular Endothelial Stem Cells

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper vessel development and homeostasis. However, studies on endothelial lineage hierarchy remain elusive due to the lack of tools to gain access as well as to directly evaluate their behavior in vivo. To address this shortcoming, a new tissue model to study angiogenesis using the mammary fat pad has been developed. The mammary gland develops mostly in the postnatal stages, including puberty and pregnancy, during which robust epithelium proliferation is accompanied by extensive vascular remodeling. Mammary fat pads provide space, matrix, and rich angiogenic stimuli from the growing mammary epithelium. Furthermore, mammary fat pads are located outside the peritoneal cavity, making them an easily accessible grafting site for assessing the angiogenic potential of exogenous cells. This work also describes an efficient tracing approach using fluorescent reporter mice to specifically label the targeted population of vascular endothelial stem cells (VESCs) in vivo. This lineage tracing method, coupled with subsequent tissue whole-mount microscopy, enable the direct visualization of targeted cells and their descendants, through which the proliferation capability can be quantified and the differentiation commitment can be fate-mapped. Using these methods, a population of bipotent protein C receptor (Procr) expressing VESCs has recently been identified in multiple vascular systems. Procr+ VESCs, giving rise to both new ECs and pericytes, actively contribute to angiogenesis during development, homeostasis, and injury repair. Overall, this manuscript describes a new mammary fat pad transplantation and in vivo lineage tracing techniques that can be used to evaluate the stem cell properties of VESCs.

This work demonstrates a novel approach to assess the proliferation, differentiation, and vessel-forming potential of vascular endothelial stem cells (VESCs) through mammary fat pad transplantation followed by whole-mount tissue preparation for microscopic observation. A lineage tracing strategy to investigate the behavior of VESCs in vivo is also presented.

Endothelial cells (ECs) are the fundamental building blocks of the vascular architecture and mediate vascular growth and remodeling to ensure proper ...

Live Imaging of Mouse Secondary Palate Fusion

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to cleft secondary palate, a common congenital anomaly in humans. Secondary palate fusion has been extensively studied leading to several proposed cellular mechanisms that may mediate this process. However, these studies have been mostly performed on fixed embryonic tissues at progressive timepoints during development or in fixed explant cultures analyzed at static timepoints. Static analysis is limited for the analysis of dynamic morphogenetic processes such a palate fusion and what types of dynamic cellular behaviors mediate palatal fusion is incompletely understood. Here we describe a protocol for live imaging of ex vivo secondary palate fusion in mouse embryos. To examine cellular behaviors of palate fusion, epithelial-specific Keratin14-cre was used to label palate epithelial cells in ROSA26-mTmGflox reporter embryos. To visualize filamentous actin, Lifeact-mRFPruby reporter mice were used. Live imaging of secondary palate fusion was performed by dissecting recently-adhered secondary palatal shelves of embryonic day (E) 14.5 stage embryos and culturing in agarose-containing media on a glass bottom dish to enable imaging with an inverted confocal microscope. Using this method, we have detected a variety of novel cellular behaviors during secondary palate fusion. An appreciation of how distinct cell behaviors are coordinated in space and time greatly contributes to our understanding of this dynamic morphogenetic process. This protocol can be applied to mutant mouse lines, or cultures treated with pharmacological inhibitors to further advance understanding of how secondary palate fusion is controlled.

Here, we present a protocol for live imaging of mouse secondary palate fusion using confocal microscopy. This protocol can be used in combination with a variety of fluorescent reporter mouse lines, and with pathway inhibitors for mechanistic insight. This protocol can be adapted for live imaging in other developmental systems.

The fusion of the secondary palatal shelves to form the intact secondary palate is a key process in mammalian development and its disruption can lead to ...

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the proteins involved in these pathways are evolutionarily conserved between mammals and other eukaryotes, such as the fruit fly Drosophila melanogaster, suggesting that similar organizing principles exist during the development of these organisms. Importantly, Drosophila has been used extensively to identify cellular and molecular mechanisms regulating processes that are required in mammals including neurogenesis, differentiation, axonal guidance, and synaptogenesis. Flies have also been used successfully to model a variety of human neurodevelopmental diseases. Here we describe a protocol for the step-by-step microdissection, fixation, and immunofluorescent localization of proteins within the adult Drosophila brain. This protocol focuses on two example neuronal populations, mushroom body neurons and retinal photoreceptors, and includes optional steps to trace individual mushroom body neurons using Mosaic Analysis with a Repressible Cell Marker (MARCM) technique. Example data from both wild-type and mutant brains are shown along with a brief description of a scoring criteria for axonal guidance defects. While this protocol highlights two well-established antibodies for investigating the morphology of mushroom body and photoreceptor neurons, other Drosophila brain regions and the localization of proteins within other brain regions can also be investigated using this protocol.

Dissection and Immunofluorescent Staining of Mushroom Body and Photoreceptor Neurons in Adult Drosophila melanogaster Brains

This protocol describes the dissection and immunostaining of adult Drosophila melanogaster brain tissues. Specifically, this protocol highlights the use of Drosophila mushroom body and photoreceptor neurons as example neuronal subsets that can be accurately used to uncover general principles underlying many aspects of neuronal development.

Nervous system development involves a sequential series of events that are coordinated by several signaling pathways and regulatory networks. Many of the ...

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological questions in cell and developmental biology. In addition, organoids together with recent technical advances in gene editing and viral gene delivery promises to advance medical research and development of new drugs for treatment of diseases. Organoids grown in vitro in basement matrix provide powerful model systems for studying the behavior and function of various proteins and are well suited for live-imaging of fluorescent-tagged proteins. However, establishing the expression and localization of the endogenous proteins in ex vivo tissue and in in vitro organoids is important to verify the behavior of the tagged proteins. To this end we have developed and modified tissue isolation, fixation, and immuno-labeling protocols for localization of microtubules, centrosomal, and associated proteins in ex vivo intestinal tissue and in in vitro intestinal organoids. The aim was for the fixative to preserve the 3D architecture of the organoids/tissue while also preserving antibody antigenicity and enabling good penetration and clearance of fixative and antibodies. Exposure to cold depolymerizes all but stable microtubules and this was a key factor when modifying the various protocols. We found that increasing the ethylenediaminetetraacetic acid (EDTA) concentration from 3 mM to 30 mM gave efficient detachment of villi and crypts in the small intestine while 3 mM EDTA was sufficient for colonic crypts. The developed formaldehyde/methanol fixation protocol gave very good structural preservation while also preserving antigenicity for effective labeling of microtubules, actin, and the end-binding (EB) proteins. It also worked for the centrosomal protein ninein although the methanol protocol worked more consistently. We further established that fixation and immuno-labeling of microtubules and associated proteins could be achieved with organoids isolated from or remaining within the basement matrix.

Immuno-fluorescent Labeling of Microtubules and Centrosomal Proteins in Ex Vivo Intestinal Tissue and 3D In Vitro Intestinal Organoids

We present protocols for isolation of intestinal 3D structures from in vivo tissue and in vitro basement matrix embedded organoids, and detail different fixation and staining protocols optimized for immuno-labeling of microtubule, centrosomal, and junctional proteins as well as cell markers including the stem cell protein Lgr5.

The advent of 3D in vitro organoids that mimic the in vivo tissue architecture and morphogenesis has greatly advanced the ability to study key biological ...

Cell Aggregation Assays to Evaluate the Binding of the Drosophila Notch with Trans-Ligands and its Inhibition by Cis-Ligands

Notch signaling is an evolutionarily conserved cell-cell communication system used broadly in animal development and adult maintenance. Interaction of the Notch receptor with ligands from neighboring cells induces activation of the signaling pathway (trans-activation), while interaction with ligands from the same cell inhibits signaling (cis-inhibition). Proper balance between trans-activation and cis-inhibition helps establish optimal levels of Notch signaling in some contexts during animal development. Because of the overlapping expression domains of Notch and its ligands in many cell types and the existence of feedback mechanisms, studying the effects of a given post-translational modification on trans- versus cis-interactions of Notch and its ligands in vivo is difficult. Here, we describe a protocol for using Drosophila S2 cells in cell-aggregation assays to assess the effects of knocking down a Notch pathway modifier on the binding of Notch to each ligand in trans and in cis. S2 cells stably or transiently transfected with a Notch-expressing vector are mixed with cells expressing each Notch ligand (S2-Delta or S2-Serrate). Trans-binding between the receptor and ligands results in the formation of heterotypic cell aggregates and is measured in terms of the number of aggregates per mL composed of &#62;6 cells. To examine the inhibitory effect of cis-ligands, S2 cells co-expressing Notch and each ligand are mixed with S2-Delta or S2-Serrate cells and the number of aggregates is quantified as described above. The relative decrease in the number of aggregates due to the presence of cis-ligands provides a measure of cis-ligand-mediated inhibition of trans-binding. These straightforward assays can provide semi-quantitative data on the effects of genetic or pharmacological manipulations on the binding of Notch to its ligands, and can help deciphering the molecular mechanisms underlying the in vivo effects of such manipulations on Notch signaling.

Cell Aggregation Assays to Evaluate the Binding of the Drosophila Notch with Trans-Ligands and its Inhibition by Cis-Ligands

Complexity of in vivo systems makes it difficult to distinguish between the activation and inhibition of Notch receptor by trans- and cis-ligands, respectively. Here, we present a protocol based on in vitro cell-aggregation assays for qualitative and semi-quantitative evaluation of the binding of Drosophila Notch to trans-ligands vs cis-ligands.

Notch signaling is an evolutionarily conserved cell-cell communication system used broadly in animal development and adult maintenance. Interaction of the ...

Tissue Preparation and Immunostaining of Mouse Craniofacial Tissues and Undecalcified Bone

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and simple protocol to detect protein expression during craniofacial morphogenesis/pathogenesis using mouse craniofacial tissues as examples. The protocol consists of preparation and cryosectioning of tissues, indirect immunofluorescence, image acquisition, and quantification. In addition, a method for preparation and cryosectioning of undecalcified hard tissues for immunostaining is described, using craniofacial tissues and long bones as examples. Those methods are key to determine the protein expression and morphological/anatomical changes in various tissues during craniofacial morphogenesis/pathogenesis. They are also applicable to other tissues with appropriate modifications. Knowledge of the histology and high quality of sections are critical to draw scientific conclusions from experimental outcomes. Potential limitations of this methodology include but are not limited to specificity of antibodies and difficulties of quantification, which are also discussed here.

Here, we present a detailed protocol to detect and quantify protein levels during craniofacial morphogenesis/pathogenesis by immunostaining using mouse craniofacial tissues as examples. In addition, we describe a method for preparation and cryosectioning of undecalcified hard tissues from young mice for immunostaining.

Tissue immunostaining provides highly specific and reliable detection of proteins of interest within a given tissue. Here we describe a complete and ...

A Multi-Omics Extraction Method for the In-Depth Analysis of Synchronized Cultures of the Green Alga Chlamydomonas reinhardtii

Microalgae have been the focus of research for their applications in the production of high value compounds, food and fuel. Moreover, they are valuable photosynthetic models facilitating the understanding of the basic cellular processes. System wide studies enable comprehensive and in-depth understanding of molecular functions of the organisms. However, multiple independent samples and protocols are required for proteomics, lipidomics and metabolomics studies introducing higher error and variability. A robust high throughput extraction method for the simultaneous extraction of chlorophyll, lipids, metabolites, proteins and starch from a single sample of the green alga Chlamydomonas reinhardtii is presented here. The illustrated experimental setup is for Chlamydomonas cultures synchronized using 12 h/12 h light/dark conditions. Samples were collected over a 24 h cell cycle to demonstrate that the metabolites, lipids and starch data obtained using various analytical platforms are well conformed. Furthermore, protein samples collected using the same extraction protocol were used to conduct detailed proteomics analysis to evaluate their quality and reproducibility. Based on the data, it can be inferred that the illustrated method provides a robust and reproducible approach to advance understanding of various biochemical pathways and their functions with greater confidence for both basic and applied research.

A Multi-Omics Extraction Method for the In-Depth Analysis of Synchronized Cultures of the Green Alga Chlamydomonas reinhardtii

System-wide analysis of multiple biomolecules is crucial to gain functional and mechanistic insights into biological processes. Hereby, an extensive protocol is described for high throughput extraction of lipids, metabolites, proteins and starch from a single sample harvested from synchronized Chlamydomonas culture.

Microalgae have been the focus of research for their applications in the production of high value compounds, food and fuel. Moreover, they are valuable ...

Use of Atomic Force Microscopy to Measure Mechanical Properties and Turgor Pressure of Plant Cells and Plant Tissues

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in indentation mode, we show how to measure an elastic modulus and use it to evaluate cell wall mechanical properties. In addition, we also explain how to evaluate turgor pressure. The main advantages of atomic force microscopy are that it is non-invasive, relatively rapid (5~20 min), and that virtually any type of living plant tissue that is superficially flat can be analyzed without the need for treatment. The resolution can be very good, depending on the tip size and on the number of measurements per unit area. One limitation of this method is that it only gives direct access to the superficial cell layer.

Here, we present atomic force microscopy (AFM), operated as a nano- and micro-indentation tool on cells and tissues. The instrument allows the simultaneous acquisition of 3D surface topography of the sample and its mechanical properties, including cell wall Young's modulus as well as turgor pressure.

We present here the use of atomic force microscopy to indent plant tissues and recover its mechanical properties. Using two different microscopes in ...

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in molecular and imaging technologies allowed us to gain insights into how biochemical signals influence morphogenetic events. However, it is increasingly evident that apart from biochemical signals, mechanical cues also impact several aspects of cell and tissue growth. The Arabidopsis shoot apical meristem (SAM) is a dome-shaped structure responsible for the generation of all aboveground organs. The organization of the cortical microtubule cytoskeleton that mediates apoplastic cellulose deposition in plant cells is spatially distinct. Visualization and quantitative assessment of patterns of cortical microtubules are necessary for understanding the biophysical nature of cells at the SAM, as cellulose is the stiffest component of the plant cell wall. The stereotypical form of cortical microtubule organization is also a consequence of tissue-wide physical forces existing at the SAM. Perturbation of these physical forces and subsequent monitoring of cortical microtubule organization allows for the identification of candidate proteins involved in mediating mechano-perception and transduction. Here we describe a protocol that helps investigate such processes.

Live Cell Imaging of Microtubule Cytoskeleton and Micromechanical Manipulation of the Arabidopsis Shoot Apical Meristem

Here we describe a protocol for live cell imaging of the cortical microtubule cytoskeleton at the shoot apical meristem and monitoring its response to changes in physical forces.

Understanding cell and tissue level regulation of growth and morphogenesis has been at the forefront of biological research for many decades. Advances in ...