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

Summary

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.

Abstract

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 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.

Introduction

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 stages^2,3,4. 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–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.

Protocol

1. Sample Preparation: fixation, dehydration, and LR-White embedding

1. 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).
   1. 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 °C and 8 h darkness at 18 °C. Select the plant tissues to be analyzed, and trim samples to be no more than 16 mm² in size.
   2. 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.
   3. After gathering all the samples, refresh the fixative.
   4. 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).
   5. Keep under a vacuum of no more than -80 kPa for 2 h at room temperature.
   6. Slowly release the vacuum, seal the glass vial and place overnight at 4 °C.
   7. 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.
   8. 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.
2. 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.
   1. 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 °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.
   2. Refresh the LR-white resin and incubate for an additional 12 h at 4 °C.
   3. 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.
   4. Apply one drop of fresh LR-white resin to the bottom of each capsule.
   5. 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).
   6. Polymerize the resin for 24 to 48 h at 58 °C, or until fully hardened.
   7. Store samples at room temperature.
      NOTE: Post polymerization LR-white resin is inert.

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’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.

1. Slide washing
   1. 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)).
   2. Maintain a mild agitation for 20 min.
   3. Transfer the staining racks to a ddH₂O bath with mild agitation for 10 min. Repeat 4 times.
   4. Carefully drain the racks before dipping briefly in 100% ethanol and let the slides dry in a dust-free environment.
   5. Store until use.
2. Poly-L-lysine coating (optional)
   NOTE: Small sections usually stick very well to clean glass slides. This step is only recommendable for larger sections (>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.
   1. 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.
   2. Let the slides dry overnight at 40 °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.

3. Sample trimming and sectioning

1. Sample trimming
   1. 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.
   2. Trim the pyramidal structure by removing fine slices of the excess resin perpendicularly to the pyramid major axis.
   3. 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).
2. 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.
   1. With an ultra-microtome, make sections with a thickness between 200 and 700 nm (Figure 1G).
   2. Check the section area by transferring some sections to a ddH₂O drop on a glass slide. Place the slide on a 50 °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.
   3. 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.
   4. Place each slide in a closed clean 10 cm square petri dish and let it dry at 50 °C.
   5. Store slides in a clean archive box until use.

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.

1. 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.
2. Pipette a 50 μL 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.
3. Prepare the primary antibody solutions (see Supplemental Table 1), 1:5 (v/v) antibody in blocking solution; make an estimate of about 40 μL per 8 mm well.
   NOTE: The concentration of the antibody must be adjusted according to the manufacture’s protocol.
4. Perform a final wash with ddH₂O for 5 min, never let the wells dry completely.
5. Pipette the primary antibody solution to the reaction wells. Pipette blocking solution to the control wells.
6. Close the incubation chamber and let stand for 2 h at room temperature followed by overnight at 4 °C. Prepare the secondary antibody solution, 1% in blocking solution (v/v) about 40 μL per well. Keep it covered with tin foil.
7. 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.
8. Pipette the secondary antibody solution to all wells. From now on, protect the slides from light.
9. Incubate for 3–4 h in the dark at room temperature.
10. Wash all wells twice for 10 min with PBS followed by another wash of 10 min with ddH₂O.
11. 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).
12. 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.
13. For a better visualization of the results overlap both images with ImageJ or similar.

Results

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

Discussion

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...

Materials

Name	Company	Catalog Number	Comments
25% (w/v) Gluteraldehyde	Agar Scientific	AGR1010	aq. Solution, methanol free
8 wells Glass reaction slides	Marinfeld	MARI1216750	other brands may be used
Acetic acid	Sigma-Aldrich	A6283
Anti-Rat IgG (wole molecule)-FITC antibody produce in GOAT	Sigma-Aldrich	F6258
cover slips, 24 mm x 50 mm	Marinfeld	MARI0100222	The cover slip should cover all the wells. Other brands may be used
ddH2O	na	na
Ethanol absolute	na	na
Fluorescent brightner 28	Sigma-Aldrich	F-6259
Gelatin capsules	Agar scientific	AGG29211	The capsule size sould feat the size of the sample.
Glass vials	na	na	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.
LR-white medium grade, embdeding resin	Agar Scientific	AGR1281	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.
Non fat dry milk	Nestlé	na	any non fat dry milk is adequate
Oven	na	na	generic laboratory oven
Petri dish, 10 cm x 10 cm square	na	na
PIPES	Sigma Aldrich	P1851
Rat generated Monoclonal Anti-Body	Plant probes	na	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
Razor blades	na	na	regular razor blades
SDS	Sigma-Aldrich	L6026
Toluidine Blue-O	Agar Scientific	AGR1727
Tween 20	Sigma-Aldrich	P9416
Ultramicrotome	Leica Microsystems	UC7
upright epifluorescence microscope with UV and FITC fluorescence filters	Leica Mycrosistems	DMLb
vaccum chamber	na	na
vaccum pump	na	na
Vectashield	vecta Labs	T-1000	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.)