The authors gratefully thank Maria Herrero and Eliseo Rivas for their helpful discussion and advice. This work was supported by the Ministerio de Econom&#237;a y Competitividad &#8212; European Regional Development Fund, European Union [grant number BES- 2010-037992 to E. F.]; the Instituto Nacional de Investigaci&#243;n y Tecnolog&#237;a Agraria y Alimentaria [grant numbers RFP2015-00015-00, RTA2014-00085-00, RTA2017-00003-00]; and the Gobierno de Arag&#243;n &#8212; European Social Fund, European Union [Grupo Consolidado A12-17R].

1. Dormancy Determination and Plant Material Collection
<ol>
	<li>Sample the flower buds in the field. Dormancy studies are long-term experiments and require adult trees big enough to collect buds and shoots all winter without compromising the trees&#8217; development during the next spring. Special orchard management could be required depending on the training system; thus, pruning may be less severe than for fruit production purposes.
	<ol>
		<li>Each week, from the start of autumn until the onset of bud break, collect and weigh 10 flower buds.</li>
		<li>Fix the flower buds by placing them into a 10-mL glass tube with a cap and soak the samples in a fixative solution of ethanol/acetic acid (3:1) for at least 24 h at 4 &#176;C. Then, discard the fixative and add 75% ethanol, generously ensuring that it covers the samples. The samples can be conserved in this solution at 4 &#176;C until use. 
		Note: Vacuum infiltration can be used to remove the air bubbles inside the bud and prevent them from floating. This facilitates the penetration of the fixative into the tissues but could damage the structure of the tissues. Try to avoid it if not necessary.</li>
	</ol>
	</li>
	<li>Sample the shoots in the field.
	<ol>
		<li>Each week, from the start of autumn until the onset of bud break, take three shoots of 15 - 30 cm in length and 5 mm in diameter, containing at least 10 flower buds each. Place them on water-soaked florist foam in a growth chamber at 22 &#177; 1 &#176;C with a 12 h light photoperiod.</li>
		<li>After 10 days in the growth chamber, pick and weigh 10 flower buds from the shoots.</li>
	</ol>
	</li>
	<li>Evaluate the bud growth and determine the dormancy status. The sampling period has to be adjusted to the conditions of the location. In the orchard conditions (Zaragoza, Spain, 41&#176;44&#8217;30&#8221; N, 0&#176;47&#8217;00&#8221; W, and 220 m above sea level), the sampling of the shoots was carried out from November 30 until the end of February or the beginning of March.
	<ol>
		<li>Weekly evaluate the response of the flower buds to the suitable growth conditions of the chamber, from the beginning of autumn until budburst in the spring, by comparing the weight of the 10 buds picked from the field.</li>
		<li>If there are no differences or those differences are less than 30%, consider that the buds have not fulfilled their chilling requirements and are endodormant22. If the differences are more than 30%, consider that the buds have fulfilled their chilling requirements and are ecodormant22.</li>
	</ol>
	</li>
</ol>
2. Plant Material Preparation for Starch Quantification
<ol>
	<li>Select about six fixed buds from each sampling date (see step 1.1). Depending on cultivars, each sweet cherry flower bud contains up to five flower primordia.
	<ol>
		<li>Remove the external bud scales and place the bud on a watchmaker&#8217;s glass with 75% ethanol to prevent it from drying out.</li>
		<li>Dissect the bud and extract at least one flower primordium per bud with the help of precision forceps and an ophthalmologic scalpel under a stereoscopic microscope. The flower primordium can be conserved in a 10-mL glass tube with 75% ethanol at 4 &#176;C, or proceed immediately to the next step.</li>
	</ol>
	</li>
	<li>Dehydrate the samples in a tertiary butyl alcohol series.
	<ol>
		<li>Replace the 75% ethanol solution with 10 mL of 75% tertiary butyl alcohol (TBA) to generously cover the samples and incubate them for 1.5 h. A Pasteur pipette can be useful to discard the solution.</li>
		<li>Repeat step 2.2.1, adding TBA with rising concentrations (85%, 95%, and 100%, v/v; and 3x with pure TBA) in a drying stove at 30 &#176;C with air extraction, since pure TBA crystalizes at room temperature (&#60; 20 &#176;C) and is very volatile and toxic. If TBA crystalizes with the sample, it would damage the tissues.</li>
	</ol>
	</li>
	<li>Embed the samples in paraffin18.
	<ol>
		<li>Melt the paraffin by introducing the paraffin pearls in the drying stove at 60 &#176;C with air extraction the previous day. The paraffin pearls can be also melted on a hotplate, but ensure that the liquid paraffin is at 60 &#176;C and not at a higher temperature, to avoid tissue damage.</li>
		<li>Replace the TBA with a mix of TBA and paraffin oil (1:1) and incubate for 24 h inside a drying stove at 60 &#176;C. Then, replace the mix of TBA and paraffin oil with pure melt paraffin and incubate for at least 6 h inside a drying stove at 60 &#176;C. Repeat it 2x and incubate the last change for at least 4 - 6 d.</li>
		<li>Place each sample on a small metal base mold over a heat surface, embedded in paraffin wax, and pace the embedding cassette. Place it over a cold surface and remove the block once the wax is solidified.</li>
	</ol>
	</li>
	<li>Section and rehydrate the preparations.
	<ol>
		<li>Prepare Haupt&#8217;s adhesive: dissolve 1 g of plain Knox gelatin in 100 mL of distilled water at 30 &#176;C; then, add 2 g of phenol loose crystals (C6H5OH) and 15 mL of glycerol. Spread the Haupt&#8217;s adhesive over a glass slide with a brush and add a drop of a 1% formaldehyde solution.</li>
		<li>Section each paraffin block at 10 &#181;m in a rotatory microtome and place the sections onto the glass slide covered with Haupt&#8217;s adhesive.</li>
		<li>Place the glass slides with the sections over a heat surface at 35 - 40 &#176;C until dry; then, move the glass slides to a slide rack.</li>
		<li>Prepare the dewax and rehydration solutions. Place 200 mL of Histoclear II in three glass-staining dishes, another one with Histoclear II:ethanol (1:1, v/v), an ethanol series (100%, 70%, 40%, v/v), and a final distilled-water wash.</li>
		<li>Dewax the sections with three washes, each 5 min, in Histoclear II and in Histoclear II:ethanol (1:1, v/v). Place the slide rack inside the glass-staining dishes, ensuring that the solution completely covers the slides, and, then, move the slide rack from one solution to the next.</li>
		<li>Rehydrate the sections by the following series of 2-min washes: an ethanol series (100%, 70%, 40%, v/v) and a final distilled-water wash. Finally, dry the glass slides at room temperature.</li>
	</ol>
	</li>
	<li>Stain the sections18.
	<ol>
		<li>Prepare I2KI stain: dissolve 2 g of potassium iodide (KI) and 0.2 g of iodine (I2) in 100 mL of distilled water. Apply a drop of fresh I2KI over the section(s) for 5 min and, then, discard the excess of stain by absorbing it with a blotting paper. Quickly proceed to the next step.</li>
		<li>Apply a small drop of a synthetic mounting media, place a small cover glass on top, and press hard. Once the mounting media dries, observe under a bright-field microscope for a preliminary evaluation of the ovary starch. 
		Note: This step is not mandatory, although a better contrast between the starch and the background is obtained. If the section has to be reused with other stains after starch quantification, place the cover glass over the drop of stain and discard the excess by absorbing it with a blotting paper (see 2.5.1). Then, after starch quantification, wash out the I2KI stain with distilled water and place the preparations over a heat surface at 35 - 40 &#176;C until dry.</li>
	</ol>
	</li>
</ol>
3. Quantification of the Starch Content
<ol>
	<li>Calibrate the optical conditions. 
	Note: The detection levels used by the image analyzer to detect the stained starch are directly dependent on light conditions and magnification of the microscope; thus, fix these conditions for all the preparations evaluated. Adapt the adjustment proposed here to the available microscope and light conditions.
	<ol>
		<li>Adjust the aperture diaphragm at the 20X magnification and the brightness control or light intensity.</li>
		<li>Make sure that there are no filters on the filter holder, and select a bright-field condition in the microscope. Select a suitable magnification (e.g., 40X for sweet cherry ovary primordium).</li>
	</ol>
	</li>
	<li>Control the image acquisition conditions. Adjust the camera settings with a stained preparation without tissue via Image | Acquisition | Pre-view.
	<ol>
		<li>Fix the brightness at 50%, the gain at 1.0x, and the histogram indicators the gamma value at 1.00 and the contrast at 0 - 100, lining up with the limits of the brightness distribution histogram.</li>
		<li>Activate the overexposure/underexposure function and adjust the exposure time at the limit of overexposure.</li>
		<li>Apply the white balance function to the complete image to display all neutral-colored components of the image without any color tone and the shading correction to the complete image to create a corrected and homogeneous image.</li>
	</ol>
	</li>
	<li>Calibrate the Image Analysis System to obtain the control values of the grey level (0, black; 255, white) of different optical densities (OD) that are obtained by the transmittance (T) values.
	<ol>
		<li>Acquire an image of a stained preparation without tissue, considered control white, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 0 | Measure | Calibrate | OK. This corresponds with a 100% transmittance; thus, an optical density of 0, according to OD = 2 - log T.</li>
		<li>Acquire an image of the same preparation with a 4N filter, which reduces the amount of light 4x, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 0.6 | Measure | Calibrate | OK. This corresponds with a 25% transmittance; thus, an optical density of 0.6, according to OD = 2 - log T.</li>
		<li>Acquire an image of the same preparation without light, considered black, and measure the grey level of the black and white image via Measure | Grey measure | Calibrate grey | Reference value = 1 | Measure | Calibrate | OK. This corresponds with a 0% transmittance; thus, an optical density of 1, according to OD = 2 - log T.</li>
	</ol>
	</li>
	<li>Detect starch.
	<ol>
		<li>Acquire a color image of the field to measure in TIFF format with a resolution of at least 300 dots per inch (dpi).</li>
		<li>Create a binary image corresponding to the area stained. Set the three color thresholds (values between 0 - 255 for each) until the binary image exactly reflects the stained starch granules observed, via Image | Detect | Select the thresholds of red, blue and green | OK. Make repeatedly visual comparisons in various preparations and tissues to tune up the final detection levels. Store and use these levels for all preparations.</li>
		<li>Quantify the starch. Convert the original color image into a black and white image with the image analysis system. Use the binary image as a superimposed mask on the black and white image via Image | Binary Edit. Measure the sum of the optical density of every pixel under the mask via Measure | Grey level | OK and consider this value as the starch content in the measured field.</li>
		<li>Repeat steps 3.4.1 - 3.4.3 in four places of the ovary primordia to obtain a representative value of the starch content in the ovary of flower primordia.</li>
		<li>Repeat steps 3.4.1 - 3.4.3 and step 3.4.5 in different flowers of each collecting date.</li>
	</ol>
	</li>
</ol>

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The authors have nothing to disclose.

Dormancy in woody perennials presents clear implications in fruit production and forestry in a changing climate, although the biological process behind dormancy remains unclear. Dormancy studies can be approached from different points of view, but the research looking for a biological marker for winter dormancy has intensified over the last years. However, most attempts to find an unequivocal indicator showing when a bud has broken dormancy have been unsuccessful3. The methodology described herein...

Temperate woody perennials adapt to the seasons by modulating their growth and development. While they develop during spring and summer, they stop growing during the autumn to go dormant in winter1. Although dormancy allows them to survive at low winter temperatures, chilling is a prerequisite for a proper budburst in spring2. The important implications of dormancy in temperate fruit production and forestry have led to diverse efforts to determine and predict the dormancy period3. In fruit tree species, empirical experiments transferring shoots to forcing conditions and statistical predictions based on data of flowering are current approaches to determine the date of the breaking of dormancy, which allows researchers to estimate the chilling requirements for each cultivar. However, how to determine the dormancy status based on biological processes remains unclear3. 
Flowering in temperate fruit trees, such as sweet cherry (Prunus avium L.), occurs once a year and lasts about a fortnight. However, flowers start to differentiate and develop about 10 months earlier, during the previous summer4. Flower primordia stop growing during the autumn to remain dormant inside the buds during winter. In this period, each cultivar needs to accumulate a particular chilling requirement for proper flowering4. Despite the lack of phenological changes in the buds during winter, flower primordia are physiologically active during dormancy, and the accumulation of chilling temperatures has been recently associated with the dynamic of starch accumulation or decrease within the cells of the ovary primordium, offering a new approach for dormancy determination5. However, the small size and the location of the ovary primordium require a special methodology.
Starch is the major storage carbohydrate in woody plant species6. Thus, changes in starch have been related to the physiological activity of the flower tissues, which need carbohydrates to support their development7,8. Different key events during the reproductive process are also related to variations in starch content in different floral structures, such as anther meiosis9, the growth of the pollen tubes through the style or ovule fertilization10. Histochemical techniques allow the detection of starch in each particular tissue of the flower primordia during dormancy. However, the difficulty remains in quantifying that starch to allow following its pattern of accumulation/decrease over time or comparing the starch content among tissues, cultivars, or years. This is due to the little amount of tissue available for the analytical techniques11. As an alternative, image analysis linked to microscopy12 allows the quantification of the starch in very small samples of plant tissue13. 
Approaches combining microscopy and image analysis have been used to quantify the content of different components in plant tissues, such as callose14, microtubes15, or starch16, by measuring the size of the area dyed by specific stains. For starch, it can be easily detected using the potassium iodide-iodine (I2KI) reaction17. This method is highly specific; I2KI intercalates within the laminar structure of starch grains and forms a dark blue or reddish-brown color, depending on the amylose content of the starch18. Sections stained with I2KI stain show adequate contrast between starch and the background tissue, allowing an unequivocal starch detection and the subsequent quantification by the image analysis system19. Although this dye is not stoichiometric, the accumulation of iodine is proportional to the length of the starch molecule, which can highly vary17. Thus, the size of the stained area expressed as the number of pixels may not reflect accurately the content of starch, since high differences in starch content could be found between fields with stained areas of similar size. As an alternative, the starch content may be evaluated by measuring the optical density of the stained granules on black and white images obtained from the microscope, as it has been reported in different tissues in apricot8,13,19, avocado10,20, and olive21.
Here, we describe a methodology that combines the experimental determination of dormancy status with the quantification of starch content in the ovary primordium tissue from autumn to spring in sweet cherry, offering a new tool for the understanding and prediction of dormancy based on the study of the biological mechanisms linked with dormancy.

<table><tbody><tr><td>Precision scale</td><td>Sartorius</td><td>CP225D</td><td></td></tr><tr><td>Stereoscopic microscope</td><td>Leica Microsystems</td><td>MZ-16</td><td></td></tr><tr><td>Drying-stove</td><td>Memmert</td><td>U15</td><td></td></tr><tr><td>Paraffin Embedding station</td><td>Leica Microsystems</td><td>EG1140H</td><td></td></tr><tr><td>Rotatory microtome</td><td>Reichert-Jung</td><td>1130/Biocut</td><td></td></tr><tr><td>Microtome blade</td><td>Feather</td><td>S35</td><td>Stainless steel</td></tr><tr><td>Bright field microscope</td><td>Leica Microsystems</td><td>DM2500</td><td></td></tr><tr><td>Digital Camera</td><td>Leica Microsystems</td><td>DC-300</td><td></td></tr><tr><td>Image Analysis System</td><td>Leica Microsystems</td><td>Quantiment Q550</td><td></td></tr></tbody></table>

combining histochemical staining and image analysis to quantify starch in the ovary primordia of sweet cherry during winter dormancy

Changes in starch in small structures are associated with key events during several plant developmental processes, including the reproductive phase from pollination to fertilization and the onset of fruiting. However, variations in starch during flower differentiation are not completely known, mainly due to the difficulty of quantifying the starch content in the particularly small structures of the flower primordia. Here, we describe a method for the quantification of starch in the ovary primordia of sweet cherry (Prunus avium L.) by using an image analysis system attached to the microscope, which allows relating the changes in starch content with the different phases of dormancy from autumn to spring. For this purpose, the dormancy status of flower buds is determined by evaluating the bud growth of shoots transferred to controlled conditions at different moments in winter time. For the quantification of starch in the ovary primordia, flower buds are sequentially collected, fixed, embedded in paraffin wax, sectioned, and stained with I2Kl (potassium iodide-iodine). Preparations are observed under the microscope and analyzed by an image analyzer that clearly distinguishes starch from the background. Starch content values are obtained by measuring the optical density of the image that corresponds to the stained starch, considering the sum of the optical density of each pixel as an estimation of the starch content of the frame studied.

Dormancy studies require the determination of the moment when the chilling requirements are fulfilled. Despite the lack of phenological changes during winter under field conditions (Figure 1A), cherry trees do not recover the capacity of growth in suitable conditions until they pass a certain period under low temperatures. The regular transference of shoots to a controlled conditions chamber (Figure 1B) during winter time allowed...

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Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

We present a methodology to quantify the starch content in the ovary primordia in sweet cherry (Prunus avium L.) during winter dormancy by using an image analysis system combined with histochemical techniques.

Changes in starch in small structures are associated with key events during several plant developmental processes, including the reproductive phase from ...

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Research

JoVE Journal

Developmental Biology

6.2K Views.  Centro de Investigación y Tecnología Agroalimentaria de Aragón. Deze methode kan helpen bij het beantwoorden van belangrijke vragen over rust in houtachtige plant, zoals waarom de bloemknoppen moeten verkouden accumuleren in de winter om goed te bloeien. Het belangrijkste voordeel van deze techniek is het mogelijk maakt de kwantificering van het zetmeel in zeer kleine monsters van bloem primordium. Deze aanpak is zeer nuttig om de fysiologische activiteit van het bloem primordium tijdens de rust te detecteren en kan ook worden toegepast op andere boomsoor...

Video: Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

Identification of Critical Conditions for Immunostaining in the Pea Aphid Embryos: Increasing Tissue Permeability and Decreasing Background Staining

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental studies. Like other aphids, A. pisum propagate rapidly via parthenogenetic viviparous reproduction, where the embryos develop within egg chambers in an assembly-line fashion in the ovariole. Previously we have established a robust platform of whole-mount in situ hybridization allowing detection of mRNA expression in the aphid embryos. For analyzing the expression of protein, though, established protocols for immunostaining the ovarioles of asexual viviparous aphids did not produce satisfactory results. Here we report conditions optimized for increasing tissue permeability and decreasing background staining, both of which were problems when applying established approaches. Optimizations include: (1) incubation of proteinase K (1 &#181;g/ml, 10 min), which was found essential for antibody penetration in mid- and late-stage aphid embryos; (2) replacement of normal goat serum/bovine serum albumin with a blocking reagent supplied by a Digoxigenin (DIG)-based buffer set and (3) application of methanol rather hydrogen peroxide (H2O2) for bleaching endogenous peroxidase; which significantly reduced the background staining in the aphid tissues. These critical conditions optimized for immunostaining will allow effective detection of gene products in the embryos of A. pisum and other aphids.

A protocol of whole-mount immunostaining on aphid embryos is presented, in which critical conditions for decreasing background staining and increasing tissue permeability are addressed. These conditions are specifically developed for effective detection of protein expression in the embryonic tissues of aphids.

The pea aphid Acyrthosiphon pisum, with a sequenced genome and abundant phenotypic plasticity, has become an emerging model for genomic and developmental ...

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

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.

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

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.

Due to its formidable tools for molecular genetic studies, Arabidopsis thaliana is one of the most prominent model species in plant biology and, ...

Whole Mount Immunofluorescence and Follicle Quantification of Cultured Mouse Ovaries

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ovarian fitness is often assessed by visualizing and quantifying follicles and oocytes. Because the ovary is an opaque three-dimensional tissue, traditional approaches require laboriously slicing the tissue into numerous serial sections in order to visualize cells throughout the entire organ. Furthermore, because quantification by this method typically entails scoring only a subset of the sections separated by the approximate diameter of an oocyte, it is prone to inaccuracy. Here, a protocol is described that instead utilizes whole organ tissue clearing and immunofluorescence staining of mouse ovaries to visualize follicles and oocytes. Compared to more traditional approaches, this protocol is advantageous for visualizing cells within the ovary for numerous reasons: 1) the ovary remains intact throughout sample preparation and processing; 2) small ovaries, which are difficult to section, can be examined with ease; 3) cellular quantification is more readily and accurately achieved; and 4) the whole organ imaged.

Here, we present a protocol to quantify follicles in cultured ovaries of young mice without serial sectioning. Using whole organ immunofluorescence and tissue clearing, physical sectioning is replaced with optical sectioning. This method of sample preparation and visualization maintains organ integrity and facilitates automated quantification of specific cells.

Research in the field of mammalian reproductive biology often involves evaluating the overall health of ovaries and testes. Specifically, in females, ...

Whole Ovary Immunofluorescence, Clearing, and Multiphoton Microscopy for Quantitative 3D Analysis of the Developing Ovarian Reserve in Mouse

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells entering meiotic prophase I are eliminated during Fetal Oocyte Attrition (FOA) and the first week of postnatal life. Three major mechanisms regulate the number of oocytes that survive during development and establish the ovarian reserve in females entering puberty. In the first wave of oocyte loss, 30-50% of the oocytes are eliminated during early FOA, a phenomenon that is attributed to high Long interspersed nuclear element-1 (LINE-1) expression. The second wave of oocyte loss is the elimination of oocytes with meiotic defects by a meiotic quality checkpoint. The third wave of oocyte loss occurs perinatally during primordial follicle formation when some oocytes fail to form follicles. It remains unclear what regulates each of these three waves of oocyte loss and how they shape the ovarian reserve in either mice or humans.
Immunofluorescence and 3D visualization have opened a new avenue to image and analyze oocyte development in the context of the whole ovary rather than in less informative 2D sections. This article provides a comprehensive protocol for whole ovary immunostaining and optical clearing, yielding preparations for imaging using multiphoton microscopy and 3D modeling using commercially available software. It shows how this method can be used to show the dynamics of oocyte attrition during ovarian development in C57BL/6J mice and quantify oocyte loss during the three waves of oocyte elimination. This protocol can be applied to prenatal and early postnatal ovaries for oocyte visualization and quantification, as well as other quantitative approaches. Importantly, the protocol was strategically developed to accommodate high-throughput, reliable, and repeatable processing that can meet the needs in toxicology, clinical diagnostics, and genomic assays of ovarian function.

Here, we present an optimized protocol for imaging entire ovaries for quantitative and qualitative analyses using whole-mount immunostaining, multiphoton microscopy, and 3D visualization and analysis. This protocol accommodates high-throughput, reliable, and repeatable processing that is applicable for toxicology, clinical diagnostics, and genomic assays of ovarian function.

Female fertility and reproductive lifespan depend on the quality and quantity of the ovarian oocyte reserve. An estimated 80% of female germ cells ...

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

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

Histochemical Staining of Arabidopsis thaliana Secondary Cell Wall Elements

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

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

Biology

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the adult plant. The female SMC (megaspore mother cell, MMC) differentiates in the ovule primordium and undergoes meiosis. The selected haploid megaspore then undergoes mitosis to form the multicellular female gametophyte, which will give rise to the gametes, the egg cell and central cell, together with accessory cells. The limited accessibility of the MMC, meiocyte and female gametophyte inside the ovule is technically challenging for cytological and cytogenetic analyses at single cell level. Particularly, direct or indirect immunodetection of cellular or nuclear epitopes is impaired by poor penetration of the reagents inside the plant cell and single-cell imaging is demised by the lack of optical clarity in whole-mount tissues.
Thus, we developed an efficient method to analyze the nuclear organization and chromatin modification at high resolution of single cell in whole-mount embedded Arabidopsis ovules. It is based on dissection and embedding of fixed ovules in a thin layer of acrylamide gel on a microscopic slide. The embedded ovules are subjected to chemical and enzymatic treatments aiming at improving tissue clarity and permeability to the immunostaining reagents. Those treatments preserve cellular and chromatin organization, DNA and protein epitopes. The samples can be used for different downstream cytological analyses, including chromatin immunostaining, fluorescence in situ hybridization (FISH), and DNA staining for heterochromatin analysis. Confocal laser scanning microscopy (CLSM) imaging, with high resolution, followed by 3D reconstruction allows for quantitative measurements at single-cell resolution.

An Efficient Method for Quantitative, Single-cell Analysis of Chromatin Modification and Nuclear Architecture in Whole-mount Ovules in Arabidopsis

We provide here an efficient and reliable protocol for immunostaining, Fluorescence in&#160;situ Hybridization, DNA staining followed by quantitative, high-resolution imaging in whole-mount Arabidopsis thaliana ovules. This method was successfully used to analyze chromatin modifications and nuclear architecture.

In flowering plants, the somatic-to-reproductive cell fate transition is marked by the specification of spore mother cells (SMCs) in floral organs of the ...

A Simple Dry Sectioning Method for Obtaining Whole-Seed-Sized Resin Section and Its Applications

The morphology, size and quantity of cells, starch granules&#160;and protein bodies in seed determine the weight and quality of seed. They are significantly different among different regions of seed. In order to view the morphologies of cells, starch granules and protein bodies clearly, and quantitatively analyze their morphology parameters accurately, the whole-seed-sized section is needed. Though the whole-seed-sized paraffin section can investigate the accumulation of storage materials in seeds, it is very difficult to quantitatively analyze the morphology parameters of cells and storage materials due to the low resolution of the thick section. The thin resin section has high resolution, but the routine resin sectioning method is not suitable to prepare the whole-seed-sized section of mature seeds with a large volume and high starch content. In this study, we present a simple dry sectioning method for preparing the whole-seed-sized resin section. The technique can prepare the cross and longitudinal whole-seed-sized sections of developing, mature, germinated, and cooked seeds embedded in LR White resin, even for large seeds with high starch content. The whole-seed-sized section can be stained with fluorescent brightener 28, iodine, and Coomassie brilliant blue R250 to specifically exhibit the morphology of cells, starch granules, and protein bodies clearly, respectively. The image obtained can also be analyzed quantitatively to show the morphology parameters of cells, starch granules, and protein bodies in different regions of seed.

This technique allows for the fast and simple preparation of whole-seed-sized resin section for the observation and analysis of cells, starch granules, and protein bodies in different regions of the seed.

The morphology, size and quantity of cells, starch granules&#160;and protein bodies in seed determine the weight and quality of seed. They are ...

Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp

Cytosolic Ca2+ plays a key role in plant development. Calcium imaging is the most versatile method to detect dynamic changes in Ca2+ in the cytoplasm. In this study, we obtained viable protoplasts of pulp cells by enzymatic hydrolysis. Isolated protoplasts were incubated with the small-molecule fluorescent reagent (Fluo-4/AM) for 30 min at 37 &#176;C. The fluorescent probes successfully stained cytosolic Ca2+ but did not accumulate in vacuoles. La3+, a Ca2+ channel blocker, decreased cytoplasmic fluorescence intensity. These results suggest that Fluo-4/AM can be used to detect changes in cytosolic Ca2+ in the fruit flesh. In summary, we present a method to effectively isolate protoplasts from flesh cells of the fruit and detect Ca2+ by loading a small-molecule calcium fluorescent reagent in the cytoplasm of pulp cells.

Staining the Cytoplasmic Ca2+ with Fluo-4/AM in Apple Pulp

Isolated protoplasts of apple pulp cells were loaded with a calcium fluorescent reagent to detect cytoplasmic Ca2+ concentration.

Cytosolic Ca2+ plays a key role in plant development. Calcium imaging is the most versatile method to detect dynamic changes in Ca2+ in the cytoplasm. In ...

Microscopy Techniques for Interpreting Fungal Colonization in Mycoheterotrophic Plants Tissues and Symbiotic Germination of Seeds

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching mycoheterotrophic plants (i.e., plants that obtain carbon from fungi), remarkable aspects of their structural adaptations, the patterns of tissue colonization by fungi, and the morphoanatomy of subterranean organs can enlighten their developmental strategies and their relationships with hyphae, the source of nutrients. Another important role of symbiotic fungi is related to the germination of orchid seeds; all Orchidaceae species are mycoheterotrophic during germination and seedling stage (initial mycoheterotrophy), even the ones that photosynthesize in adult stages. Due to the lack of nutritional reserves in orchid seeds, fungal symbionts are essential to provide substrates and enable germination. Analyzing germination stages by structural perspectives can also answer important questions regarding the fungi interaction with the seeds. Different imaging techniques can be applied to unveil fungi endophytes in plant tissues, as are proposed in this article. Freehand and thin sections of plant organs can be stained and then observed using light microscopy. A fluorochrome conjugated to wheat germ agglutinin can be applied to the fungi and co-incubated with Calcofluor White to highlight plant cell walls in confocal microscopy. In addition, the methodologies of scanning and transmission electron microscopy are detailed for mycoheterotrophic orchids, and the possibilities of applying such protocols in related plants is explored. Symbiotic germination of orchid seeds (i.e., in the presence of mycorrhizal fungi) is described in the protocol in detail, along with possibilities of preparing the structures obtained from different stages of germination for analyses with light, confocal, and electron microscopy.

This protocol aims to provide detailed procedures for collecting, fixing, and maintaining mycoheterotrophic plant samples, applying different microscopy techniques such as scanning and transmission electron microscopy, light, confocal, and fluorescence microscopy to study fungal colonization in plants tissues and seeds germinated with mycorrhizal fungi.

Structural botany is an indispensable perspective to fully understand the ecology, physiology, development, and evolution of plants. When researching ...

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls requires cellular reprogramming and changes in the metabolism of the infected plant. This is necessary to establish a pathogen-oriented physiological sink toward which the host nutrients are redirected. For a complete understanding of this particular plant-pathogen interaction and the mechanisms by which host growth and development are subverted and repatterned, it is essential to track and observe the internal changes accompanying gall formation with cellular resolution. Methods combining fluorescent stains and fluorescent proteins are often employed to study anatomical and physiological responses in plants. Unfortunately, the large size of galls and their low transparency act as major hurdles in performing whole-mount observations under the microscope. Moreover, low transparency limits the employment of fluorescence microscopy to study clubroot disease progression and gall formation. This article presents an optimized method for fixing and clearing galls to facilitate epifluorescence and confocal microscopy for inspecting P. brassicae-infected galls. A tissue-clearing protocol for rapid optical clearing was used followed by vibratome sectioning to detect anatomical changes and localize gene expression with promoter fusions and reporter lines tagged with fluorescent proteins. This method will prove useful for studying cellular and physiological responses in other pathogen-triggered structures in plants, such as nematode-induced syncytia and root knots, as well as leaf galls and deformations caused by insects.

Combining Clearing and Fluorescence Microscopy for Visualising Changes in Gene Expression and Physiological Responses to Plasmodiophora brassicae

The present protocol describes an optimized method for the histological observation of galls induced by Plasmodiophora brassicae. Vibratome sections of hypocotyls are cleared before fluorescence imaging to study the involvement of transcription factors and phytohormones during disease progression. This protocol overcomes resin embedding limitations, enabling in planta visualization of fluorescent proteins.

Infection of Brassica crops by the soilborne protist Plasmodiophora brassicae leads to gall formation on the underground organs. The formation of galls ...

Combining Histochemical Staining and Image Analysis to Quantify Starch in the Ovary Primordia of Sweet Cherry during Winter Dormancy

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