Whole-mount Clearing and Staining of Arabidopsis Flower Organs and Siliques

Summary

In this protocol, we describe techniques for the proper dissection of Arabidopsis flowers and siliques, some basic clearing techniques, and selected staining procedures for whole-mount observations of reproductive structures.

Abstract

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, especially, in plant reproductive biology. However, plant morphological, anatomical, and ultrastructural analyses traditionally involve time-consuming embedding and sectioning procedures for bright field, scanning, and electron microscopy. Recent progress in confocal fluorescence microscopy, state-of-the-art 3-D computer-aided microscopic analyses, and the continuous refinement of molecular techniques to be used on minimally processed whole-mount specimens, has led to an increased demand for developing efficient and minimal sample processing techniques. In this protocol, we describe techniques for properly dissecting Arabidopsis flowers and siliques, basic clearing techniques, and some staining procedures for whole-mount observations of reproductive structures.

Introduction

Flowers are among the most important defining organs of angiosperms. Flowering plants appeared some 90–130 million years ago¹, and diversified so fast that their rapid appearance was described as an "abominable mystery" by Charles Darwin². The interests of plant researchers in flower development are diverse. Some research has focused on understanding the evolutionary origin of the flower as a whole, or the evolution of specific anatomical, structural, and functional properties of flowers^3,4,5,6. The high variation in floral form and structure, as well as the modes of sexual and asexual reproduction relying on them, make the flower a highly complex structure. This has led to diverse efforts to characterize the anatomy and structural features of floral organs, using light and electron microscopical techniques that could be combined with genetic and molecular investigations⁷. Furthermore, as the source of fruits and seeds, flowers are of paramount importance for human and animal nutrition. Therefore, the characterization of flower and fruit development has many implications for applied research, including food security for an ever-increasing human population and ecological conservation strategies under a changing environment^8,9,10.

Flower development in Arabidopsis starts with flower induction and the transformation of the vegetative meristem to an inflorescence (group of flowers) meristem. Flower primordia are initiated laterally on the flank of the inflorescence meristem¹¹. The floral organ primordia form progressively in concentric whorls from the outside to the center of the flower, and eventually develop into sepals, petals, stamens, and carpels⁷. These floral organs fulfill distinct nutritive, protective, and functional (e.g., pollinator attraction) roles in different plant species, with the sexual organs sustaining the development of male and female gametophytes, respectively^12,13. The gametophytes, in turn, each differentiate a pair of male (sperm) and female gametes (egg and central cell), which unite upon double fertilization to form the next generation, the zygote, and the primary endosperm, a terminal tissue supporting the development of the embryo^14,15. Fruit and seed development support the growth, maturation, and preservation of the embryo and, eventually, its dispersal. Extensive research has been performed to characterize flower and embryo development in diverse plant species, especially in the model species Arabidopsis^7,16,17.

Early microscopic analyses of flower development were based on time-consuming sample processing and observation techniques, such as paraffin or resin embedding and sectioning, combined with light or electron microscopy. These traditional microscopic techniques were often used in combination with molecular genetic investigations, such as microscopical analyses of mutants, the localization of RNA by in situ hybridization, or the immuno-detection of proteins. Recent progress in wide-field and confocal fluorescence microscopy, in state-of-the-art 3-D computer-aided image analyses, and the continuous refinement of molecular methods that can be used on minimally processed whole-mount specimens, has led to a need for efficient and minimal sample processing techniques that are preferentially amenable to quantitative analyses. In recent years, significant progress has been made in developing clearing techniques on whole-mount animal specimen. They render the sample transparent either by using aqueous urea- or sugar-based reagents (e.g., SCALE, SeeDB, CUBIC)^18,19,20, or by selectively removing lipids (using the detergent SDS) after embedding samples in stable hydrogels; the removal of lipids can be achieved either by passive diffusion (e.g., modified CLARITY protocol²¹, PACT-PARS-RIMS²²) or actively by electrophoresis (original CLARITY protocol²³ and ACT-PRESTO²⁴). Encouraged by this fast progress, some derived techniques are also emerging for use in plants.

In this methods paper focused on the model Arabidopsis, we describe the procedure for the proper dissection of flower buds, flowers, and young siliques, and the clearing of whole-mount samples for diverse staining and observation procedures using classical or a recent SDS-based clearing method. Examples for starch, callose, and chromatin staining are given. Although these procedures may need further improvements and adaptations when used with other species, we hope they will set the stage for further research on these simple but critical methods that are the starting point of many research projects.

Protocol

1. Flower and Silique Fixation

1. Harvest flowers and siliques from plants synchronized at the opening of the first flower.
   NOTE: Under the experimental conditions used here, plants start flowering approximately 21 days after transplantation from Murashige and Skoog (MS) plates to soil. Seeds are stratified for 3–4 days at 4 °C and germinated/grown on MS plates at 22 °C/16 h light and 18 °C/8 h dark for 8 to 10 days, before transplanting the seedlings on nutrient-rich soil in pots kept under the same conditions (Figure 1). The number of replicates depends on the specific research objective, but a minimum of 5 inflorescences (from 5 individual plants) for each treatment is recommended. If flowers are the experimental unit, a minimum of 10 flowers per replicate is recommended.
2. Cut inflorescences or flowers (Figure 1E-H) using small scissors (e.g., nail scissors) and place them immediately in a microcentrifuge tube (for single inflorescence/flower fixation) or a conical tube (for collective fixations) containing freshly made Carnoy's, methanol/acetic acid, or FPA50 fixatives (depending on the application, see below) on ice.
   NOTE: Samples should be completely submerged in the fixative.
3. Leave the tissue within the fixative for 4 h to overnight at 4 °C.
   NOTE: Vacuum infiltration can be used for speeding up the penetration of the fixative into the tissue, but this may be detrimental to preserving the structure of the tissue. The authors do not implement vacuum infiltration.
4. Remove the fixative, add enough 70% ethanol to cover the samples, and return them to 4 °C for at least 24 h; samples can stay indefinitely in this solution. After removing ethanol, proceed immediately to the next step; either dissection, or SDS clearing.

2. Dissection under the Stereomicroscope

1. Put freshly made 70% ethanol in a watch maker's glass placed on a small Petri dish for support.
2. Place the inflorescence/flower/silique on this freshly made fixative and dissect under the stereomicroscope using forceps and a syringe with a needle.
   1. Use a watch maker's glass or similar device that allows the samples to be totally covered with ethanol (recommended) for the dissection of inflorescences, and for tearing apart floral organs of mature flowers (sepals, petals, and stamen can be dissected at this stage) to avoid the risk of samples drying out.
3. Dissect siliques and small flower buds on a slide as described below.
   1. Put the watch maker's glass with pre-processed samples aside, and use a normal slide for the final dissection.
   2. Move the sample of interest to the slide and add 10 µL of fresh 70% ethanol. Work swiftly on the final dissection and add 10 µL of ethanol, if necessary, to keep the sample moist without excessive liquid.
   3. For releasing pollen grains from stamens, see step 5.1. For dissecting carpels and ovules, follow the procedure described in Figure 2. This procedure applies to all stages of development of the carpel, including the stage of development of green siliques.
      NOTE: Do not add ethanol if there is sufficient ethanol carry-over from moving the samples; dissecting very small samples is much easier in a minimal amount of liquid. Always keep only the organs of interest (sepals, petals, stamens, carpels, ovules, seeds, or pollen grains) on the slide. This procedure will ensure a uniform thickness between the slide and coverslip, corresponding to the thickness of the organ under examination, resulting in a higher homogeneity, and thus efficiency for microscopic examinations (higher number of target specimens in the same focal plane).
4. Use a small piece of blotting paper to absorb and remove as much ethanol as possible. Quickly proceed to the next step (section 3, 4, or 6) before the sample dries out.

3. Chloral Hydrate-Based Clearing and Combined Clearing-staining

NOTE: Best results for chloral hydrate-based clearing are obtained with FPA50 fixed material.

1. Place 20 µL of clearing solution (chloral hydrate/glycerol, modified Hoyer's medium, Herr's 4½, or Herr's IKI-4½ solutions) on the specimen-bearing slide. Using a pair of syringes with hypodermal needles, position the specimens as needed by reducing the space between them, and flipping those where the organ of interest faces downward. Remove any remaining air bubble using the needle.
2. Lower a coverslip sideways and gently place it, pressing very gently, and wait until the clearing solution fills the space in between. Add minimal clearing solution, if needed, to fill the entire space under the coverslip.
3. Place the slide on a slide holder and leave it under the fume hood. Proceed to microscopic observations after at least 4 h and during the following 4–5 days.

4. Combined Alexander Staining and Herr's 4½ Clearing of Anthers

NOTE: The original Alexander protocol is based on releasing pollen grains on the slide before staining. An efficient and more informative, modified Alexander staining and clearing procedure is the staining of mature pollen grains within mature but non-dehiscent anthers.

1. Choose freshly harvested flower buds that are about to open with non-dehiscent anthers.
   NOTE: Do not take younger flower buds because Alexander staining is intended for mature pollen grains and does not efficiently stain immature pollen grains. A minimum of 5 flowers per treatment harvested from different plants and inflorescences is recommended.
2. Prepare a 96-well plate with enough Alexander's solution to submerge a flower bud. Expose the anthers by removing the sepals and petals and immerse them in Alexander's solution. Keep the samples under the fume hood for 1–3 h.
3. Replace Alexander's solution with Herr's 4½ solution and place the multi-well plate back under the fume hood for an overnight incubation.
4. Using forceps, gently move the cleared flower buds onto a new slide. Since some carry-over of Alexander's solution may occur, use a blotting paper to remove as much clearing solution as possible.
5. Dissect the stamens and remove all other tissues. Follow the steps in section 3 using Herr's 4½ solution.

5. Removing the Exine from Pollen Grains

NOTE: We recommend using Carnoy's fixative for DAPI staining.

1. Follow the fixation and dissection procedures described in sections 1 and 2. By now, one should have one to many stamens on the slide within a minimum amount of 70% ethanol.
2. Use a blotting paper to remove excess ethanol, and add 5–10 µL of DAPI solution. Using a couple of syringes with needles, release the pollen grains within that small amount of solution (e.g., use one syringe to immobilize the stamen and the other to release the pollen grains). Add another 5 µL of DAPI if the solution dries out.
3. Removing all debris of the stamen is a critical step, such that only pollen grains are left between the slide and the coverslip.
4. Place a coverslip on the specimen-bearing slide by lowering it sideways and add the minimal amount of DAPI solution required to fill the space. Place an index finger on the coverslip, and without squashing too much make a quick, firm, and short sliding movement.
   NOTE: To gauge the necessary force and amplitude of the sliding movement, this step should be practiced several times, checking the results each time under the microscope.
5. Add DAPI solution if needed and place the slide in a humid box in the dark at 4 °C for 15 min to 1 h. Proceed to observe samples under wide-field fluorescence or confocal microscopy within 24 h. Try to combine this procedure with SDS clearing to check whether further improvements can be obtained.

6. Sodium Dodecyl Sulfate (SDS) Clearing

NOTE: Depending on the floral organ to be analyzed, and on the researcher's skills to dissect very soft and small specimens, the SDS treatment can be carried out either before (for experienced researchers) or after the dissection step (for less experienced researchers) on methanol-acetic acid fixed material.

1. Use a watch maker's glass, a concave slide, or a microcentrifuge tube to incubate dissected or non-dissected samples in 1% SDS and 0.2 N NaOH solution overnight at room temperature.
2. Handle with care because the sample is very soft and has a transparent appearance. Rinse several times with water.
   NOTE: Remnants of the SDS solution might lead to precipitation when adding the staining solution, e.g., aniline blue.
3. For non-dissected samples, follow the dissection procedure described in section 2, using water instead of 70% ethanol. After dissecting the desired tissue, remove any remnants and debris, and any excess water with a blotting paper, then add 20–40 µL of staining solution, and proceed with step 6.5.
4. For dissected samples, carefully move the sample from the washing solution and place it onto a clean slide, and add 20–40 µL of staining solution.
5. Gently place a coverslip on the slide by lowering it sideways. Take care not to squash the preparation. If the tissue is too soft, place any thin separator in between the slide and the coverslip at its corners (e.g., tape or a broken coverslip), leaving a chamber-like space between slide and coverslip, such that the coverslip doesn't crush the specimen.
6. Place all the slides within a humid box, cover it with aluminum foil, and place it at 4 °C for at least 1 h. Proceed to observe the specimen using widefield fluorescence or confocal microscopy within 24 h.

Results

Arabidopsis belongs to the Brassicacea family, bearing inflorescences with bisexual flowers arranged in a corymb (Figure 1). Each flower has four sepals, four petals, six stamens (four long and two short), and a syncarpous gynoecium consisting of two congenitally fused carpels (Figure 1F-H) arranged in four concentric whorls^25,26. Arabidopsis...

Discussion

The existence of many flower buds within a single inflorescence of Arabidopsis, spanning all flower developmental stages, offers a unique opportunity for studies aimed at characterizing an effect of a treatment or a developmental feature simultaneously across the different stages of flower development. A good reference point between different individual plants is the opening of the first flower of the main inflorescence. Plants are treated in such a way that flowering is synchronized as much as possible (e.g...

Materials

Name	Company	Catalog Number	Comments
Reagents and Materials
Ethanol	Scharlau	ET00102500
Acetic Acid	Applichem	A3686,2500	100% Molecular biology grade
Glacial Acetic Acid	Sigma-Aldrich	320099	Molecular Biology Grade
Methanol	Scharlau	ME03062500
Formaldehyde Solution	Sigma-Aldrich	F1635
Propionic acid	Sigma-Aldrich	81910-250 ml
Chloral hydrate	Sigma-Aldrich	15307
Glycerol	Roth	3783.1
Gum arabic	Fluka	51198
Lactic acid	Fluka	69773
Phenol	Sigma-Aldrich	77607-250ML	We used liquid phenol (use the density to find the required volume for your solution)
Clove oil	Sigma-Aldrich	C8392-100ML
Xylene	Roth	4436.1
Iodine	Fluka	57665
Potassium iodide	Merck	5043
Malachite Green	Fluka	63160
Fuchsin acid	Fluka	84600
Orange G	Sigma	7252
Sodium Dodecyl Sulfate	Sigma-Aldrich	L3771	Molecular Biology Grade
Sodium hydroxide	Sigma-Aldrich	71690
Sodium di-Hydrogen Phosphate	Applichem	A1047,1000
Sodium phosphate dibasic	Sigma-Aldrich	S9763-1KG
Potassium phosphate	Sigma-Aldrich	04347
EDTA	Applichem	A2937,1000
Calcofluor	Sigma	F6259	Fluorescent brightener 28
Auramine	Chroma	10120
DAPI	Sigma	D9542	toxic
Triton-X-100	Sigma	T8787
Aniline blue	Merck	1275
MS medium	Carolina	19-57030
Nutrient-rich substrate	Einheitserde	ED73
Watch maker's glass			No specific brand
15 ml falcon centrifuge tubes	VWR	62406-200
Dumond Forceps	Actimed	0208-5SPSF-PS
Forceps	DUMONT BIOLOGY	0108-5
Syringe	BD	BD Plastipak 300013	1 ml
Preparation needle	BD	BD Microlance 304000
Microscope slides	Thermo Scientific	10143562CE	cut edges
Coverslips	Thermo Scientific	DV40008
Humid box			A plastic box with damp paper towel and slide supports inside
Name	Company	Catalog Number	Comments
Solutions
Fixatives
Carnoy's (Farmer's) fixative			Absolute ethanol : glacial acetic acid, 3:1 (ml:ml)
Methanol/acetic acid fixative			50 % (v/v) methanol, 10 % (v/v) glacial acetic acid in deionized water
FPA50 fixative			Formalin, propionic acid, 50% ethanol; 5:5:90 (ml:ml:ml)
Clearing solutions
Chloral hydrate/glycerol			Chloral hydrate : glycerol : water, 8:1:2 (g:ml:ml). Can be used for all flower developmental stages and for silique development with DIC microscopy. The best fixative is the formaline based FPA50
Modified Hoyer			Gum arabic 7.5 g, chloral hydrate 100 g, glycerol 5 ml , water 30 ml. Can be used for all flower developmental stages and for silique development with DIC microscopy. The best fixative is the formaline based FPA50
Herr's 4½ clearing fluid			Lactic acid, chloral hydrate, phenol crystals, clove oil, xylene; 2:2:2:2:1, by weight. Can be used for all flower developmental stages (especially for stamen development) and for silique development with DIC microscopy. The best fixative is the formaline based FPA50
SDS/NaOH solution			Mix-dilute the the SDS and the NaOH stock solution to 1% SDS / 0.2 N NaOH (10x dilution). For all stages of flower and silique developmental stages. The best fixative is the methano/acetic acid fixative; the other two fixatives can also be used. Can be combined with calcofluor, auramine, DAPI, and aniline blue staining solution.
SDS stock solution			10 % (w/v) sodium dodecyl sulphate. Dissolve 10 g sodium dodecyl sulphate in 80 ml deionized water and make up to 100 ml with deionized water.
NaOH stock solution			2 N NaOH solution: dissolve 4 g of NaOH in 40 ml of deionized water and make up to 100 ml with deionized water
Combined clearing and staining solutions
Herr's IKI-4½			To a standard 4½ (9 g in total) add: 100 mg iodine, 500 mg potassium iodide. This clearing solution can be used for all flower developmental stages and for silique development, either for increasing contrast or for characterizing starch dynamics. Use FPA50 for structural analysis and Carnoy's fixative for quantitative starch analysis.
Alexander staining			Ethanol 95% 10ml, malachite green (1% in 95% EtOH) 1 ml, fuchsin acid (1% in ddH2O) 5ml, orange G (1% in ddH2O) 0.5ml, phenol 5g, chloral hydrate 5g, glacial acetic acid 2ml, glycerol 25ml . This clearing/staining alone or in combination with Herr's 4½ solution can be used to evaluate pollen abortion in flowers with mature and tricellular pollen grains. It's used on freshly harved non-fixed material.
Staining solutions
Calcofluor solution			Calcofluor 0.007% in water (g:ml). Originally used as an optical brightner. Can be used for staining cellulose, carboxylated polysaccharides and callose in cell walls. Frequently used to stain the intine of the pollen grain. All three fixatives can be used with this solution.
Auramine solution			Auramine 0.01% in water (g:ml). This lipophilic fluorscent dye can be used for staining cuticles, cutin, and exine among others. All three fixatives can be used with this solution
Calcofluor-Auramine mixture			Auramine solution : Calcofluor solution, 3:1. Can be used for a combined staining by both solutions. Other proportions can be assayed maintaining a smaller proportion of calcofluor with respect to auramine.
DAPI solution			DAPI 0.4 ug/ml, 0.1 M sodium phosphate buffer (pH 7), 0.1% Triton-X-100, 1 mM EDTA. This solution can be used for staining chromosome spreads during male and female meiosis, and cell nuclei of any tissue. Frequently used for studying pollen grain development. Carnoy's and methanol/ acetic acid are the best fixatives for this solution. Formaldehyde-based fixatives such as FPA50 may interfere with the staining. Excitation in the UV and maximum emission around 461 nm.
Sodium phosphate buffer (0.1 M)			Proton receptor: 0.2 M Na2HPO4, proton donor: 0.2 M NaH2PO4, ratio proton donor / proton receptor: 1.364 ( for a pH 7)
Aniline blue solution			0.1% (w/v) aniline blue, 108 mM K3PO4 (pH 11), 2% glycerol. This solution can be used for staining callose and cellulose of many stages of development (e.g callose deposition in male and female terads, callose plugs in pollen tubes). Excitation in the UV and maximum emission around 455. It can also be excited at 514 nm with emission in the red for cell content staining.