The authors acknowledge the European Union H2020 Program (project GAIN4CROPS, GA no. 862087). The Centre of Excellence AgroCropFuture Agroecology and new crops in future climates is financed by the Ministry of Education and Research, Estonia. We wish to thank Professor Evelin Loit-Harro for providing seeds of T. aestivum and Z. mays, Paula Palmet and Vaiko Vainola for their assistance in preparing leaf cross-sections, and Jo&#227;o Paulo de Silva Souza for assistance with analysis. All images were obtained from the microscopy unit of the Estonian University of Life Sciences under various projects.

1. Plant growth conditions
<ol>
	<li>Select wheat (Triticum aestivum L. Var. Winter Wheat, &#34;Fredis&#34;) and maize (Zea mays L. Var. Saccharata, &#34;Golden Bantam&#34;) as representative C3 and C4 plants, respectively. Grow both plant species in an environment-controlled growth chamber from seeds.</li>
	<li>Maintain relative humidity at 55% with the air temperature at 25 &#176;C/18 &#176;C (day/night) with a 12 h light period. Maintain photosynthetic photon flux density (Q) at plant surface at 1200 &#181;mol/m2/s to represent their natural growing conditions. Grow all plants at an ambient CO2 concentration of 400 &#181;mol/mol.</li>
	<li>Water plants every 2nd day with distilled water and fertilize with Hoagland&#39;s solution 10 days after seedling emergence. Grow the plants at the given conditions for 60 days and use fully expanded leaves for microscopic analyses.</li>
</ol>
2. Preparation and viewing of free-hand sections
<ol>
	<li>Select 3 mature non-senescent leaves from each plant and keep them in distilled water. Using a new single-sided razor blade, make one leveling cut to form a right angle with the leaf surface to ensure the sections are transverse cross-sections made perpendicular to the leaf surface.</li>
	<li>Place the leaf specimen on a glass plate on a piece of dental wax to stabilize the sample and avoid slipping. Cut by moving the blade straight down on the leaf specimen to cut cleanly through cells. Keep the cross-sections in distilled water.</li>
	<li>Mount the samples on a glass slide by placing them on a water droplet and covering them with a glass coverslip for microscopical viewing and imaging. Avoid trapping air bubbles.</li>
	<li>View the slide under a compound light microscope at 10x and 20x magnification. Save images as per the software guide, along with the relevant scale.</li>
</ol>
3. Preparation of semi-thin sections
<ol>
	<li>Cut sections of approximately 3 mm x 5 mm in size from intercostal areas alongside the midrib from leaves from both Z. mays and T. aestivum on a glass plate as in step 2.2. Ensure that the length of the cut section runs along the grain of the leaf.</li>
	<li>Place the plant material in a syringe with 1 mL of fixation buffer (4% glutaric aldehyde in 0.1 M phosphate buffer, pH = 6.9, Table 1).</li>
	<li>Hold the syringe vertically, remove the air, and, holding a finger on the tip, pump the syringe to create a vacuum. Repeat until the samples lie at the bottom of the syringe, indicating successful infiltration.</li>
	<li>Transfer the contents of the syringe to a labeled glass vial and store at 4 &#176;C for 48 h before the next step (Table 2).</li>
	<li>For each vial, gently remove the fixative with a pipette and add the phosphate buffer until the sample is covered. Leave for 30 min. Repeat this step 3x.</li>
	<li>Remove the phosphate buffer with a pipette and add the osmium tetroxide (CAUTION) until the sample is covered. Samples and other organic matter will turn black. Use double gloves. Leave for 1 h.</li>
	<li>Remove the previous solution with a pipette and cover the samples with distilled water. Leave for 30 min Repeat this step 3x and use double gloves.</li>
	<li>Remove the previous solution with a pipette and cover the samples with the 50/50 distilled water-ethanol solution. Leave for 30 min. Repeat this step with distilled water-ethanol solutions in the following series: 30/70, 20/80, 10/90, and 2x at 100 % ethanol. Samples in any of these solutions can be left overnight at 4 &#176;C.</li>
	<li>Remove the previous solution with a pipette and cover the samples with 1/3 resin-ethanol solution. Put the samples on a mixer plate and leave for 2 h. Repeat this step with resin-ethanol solutions in the following series: 1/2, 1/1, 2/1, 3/1, and 2x at 100% resin.&#160;Samples in any of these solutions can be left overnight at 4 &#176;C.</li>
	<li>Cover the cavities of a flat embedding mold (16 cavities, cavity dimensions: 14 L x 4.8 W x 3 D (mm)) with 100% resin and place the samples on one end of the cavity. Prepare pencil-written paper labels for each sample and place them on the other end of the cavity.</li>
	<li>Completely fill all the cavities with 100% resin and straighten the samples and labels if they moved. Cover the mold with embedding film and polymerize in an oven as per acrylic resin product guidelines.</li>
	<li>Set the polymerized block with the sample in the specimen holder of the ultramicrotome. Trim the block using a rough glass knife until the tissue becomes visible and excess resin is eliminated.</li>
	<li>Cut semi-thin sections transversely (1 &#181;m) using a glass or diamond knife. Collect the sections using a metal inoculation loop from the water surface and place them on a glass slide. Dry the slide on a hot plate (60 &#176;C) to fix the sections onto the glass.</li>
	<li>Stain the fixed sections with toluidine blue (1% aqueous solution) for 5 s before rinsing with distilled water and drying again on the hot plate. 
	NOTE: Ensure the chemical fixation process is performed under a fume hood. Glass knives should be replaced periodically to ensure sharp cuts that allow for sections without tears or distortions.</li>
</ol>
4. Imaging of samples
<ol>
	<li>Imaging of fresh leaf sections
	<ol>
		<li>Set up the compound light microscope with the accompanying software as per the manual.</li>
		<li>Place the glass slide on the stage. Adjust the eyepiece and view the section at 10x to obtain an overview, then at 20x and 40x objectives.</li>
		<li>At each objective, navigate the Live channel, focus on the area of interest, and locate the bundle sheath cells around the vein.</li>
		<li>Once focused, click the Freeze button and add the scale by pressing the Scale Bar button in the Tools tab. Save the image in .TIF format for image analysis.</li>
	</ol>
	</li>
	<li>Imaging of resin-infiltrated sections
	<ol>
		<li>Set up the automated light microscope in connection with the accompanying software on the computer.</li>
		<li>Load the transparent channel and place the slide on the microscope stage. Focus on the section using a 40x objective magnification and adjust the brightness to ensure all cell structures are visible.</li>
		<li>Set the location area of the section using the navigator function to ensure the entire section is imaged and click the Stitch button.</li>
		<li>Click Done and then Start to allow the microscope to image the entire section. Open the image analysis software and open the tiles taken by the microscope.</li>
		<li>Using the Align feature, ensure that the tiles do not overlap. After generating the image, adjust the positioning, brightness, and rotation using the Adjust tab.</li>
		<li>Print the scale on the image before saving in .TIF format.</li>
	</ol>
	</li>
	<li>Anatomical measurements using ImageJ software
	<ol>
		<li>Open ImageJ software and load the image by dragging it into the window.</li>
		<li>Using the Line tool, calibrate the scale as follows: Draw a line over the scale bar, choose Analyze &#62; Set Scale, change the known distance to the length of the scale bar, and change the unit of length to the correct unit.</li>
		<li>Measure the prospective trait by selecting the Line Tool, drawing a line across the area of interest, and pressing the M key.</li>
		<li>For area measurements, select the Polygon Selection Tool and draw around the tissue of interest. Conclude by pressing the M key. The measurements show up as a pop-up window, which can be copied to a spreadsheet for further data analysis.</li>
	</ol>
	</li>
</ol>

Microscopical Analysis for Enhancing Photosynthetic Efficiency in C4 Crops

The authors declare no conflicts of interest.

In this article, we discuss both the quantitative and qualitative methods of measuring leaf anatomy and ways in which they can be optimized. Furthermore, the methodology is applied to representative crop species so as to determine which anatomical traits are most useful in distinguishing between C3 and C4 cross-sections. Understanding these traits is essential as hybrid species, termed C2 photosynthesis, is becoming a more promising avenue of research. As of now, only one crop species, <e...

Photosynthesis is a fundamental process where light energy is converted into chemical energy, serving as the cornerstone of terrestrial trophic networks. The majority of plants follow the C3 pathway of photosynthesis, where the primary photosynthetic product is the three-carbon compound glycerate 3-phosphate. C3 photosynthesis evolved over 2 billion years ago in the atmosphere abundant in CO2 and low in O21. The key photosynthetic enzyme ribulose 1,5 bisphosphate carboxylase/oxygenase (Rubisco), which evolved under these conditions, is suboptimal for current low CO2 high O2 conditions as it competitively reacts with O2, initiating photorespiration2. Photorespiration is a wasteful pathway that consumes energy instead of producing it and releasing CO2 as a byproduct. Consequently, it is crucial to maintain high CO2 concentration around Rubisco in chloroplasts to prevent oxygenation3,4. Due to the inability of C3 plants to concentrate CO2, there is a significant CO2 drawdown from ambient air to chloroplasts, curbing photosynthesis and affecting plant growth and biomass production2,5,6.
In C3 plants, photosynthesis is limited by the entrance of CO2 through stomata, its diffusion through the mesophyll, and the biochemical activity of photosynthetic enzymes. The entry of CO2 is first limited by stomatal conductance, which is controlled by environmental conditions such as air temperature and humidity. Then CO2 diffuses through the leaf&#39;s internal gaseous and liquid phase to Rubisco7. In C3 plants, all stages of photosynthesis occur in the chloroplasts of mesophyll cells, and plants need to maintain a constant influx of CO2 from the atmosphere into chloroplasts. The dependence of CO2 availability in chloroplasts on stomatal openness, mesophyll architecture, and individual cell and chloroplast characteristics leaves plants susceptible to environmental constraints that eventually affect photosynthesis, like low water availability and high temperatures7,8,9,10, particularly highlighting their vulnerability to climate change conditions11.
Given the challenges posed by the inefficiencies of the C3 pathway, as well as limitations in maintaining optimal CO2 levels and susceptibility to environmental factors, in certain plants, another pathway, the C4 photosynthesis pathway, has evolved. Characteristically, C4 plants have two spatially separated biochemical pathways; the initial CO2 fixation occurs in mesophyll cells by phosphoenolpyruvate carboxylase, which has a higher affinity for CO2 than Rubisco and also lacks oxygenation activity. The formed C4 product is further transported to bundle sheath cells, where it is decarboxylated, and CO2 is again released and fixed by Rubisco (C3 photosynthesis)12,13,14. The greater affinity of PEP carboxylase to CO2 and thick cell walls of bundle sheath cells allows CO2 concentration in bundle sheath cells, and thus, C4 plants minimize the photorespiration by spatially segregating CO2 fixation and the Calvin cycle. The adoption of the C4 pathway showcases nature&#39;s adaptive response to environmental constraints, offering insights into potential strategies for improving crop productivity and resilience in changing climate conditions15.
The specialized anatomy of the leaf structure in C4 plants is characterized by veins surrounded by enlarged vascular bundle sheath cells containing chloroplasts and with a radial arrangement of mesophyll cells in an outer ring patterning around bundle sheath cells. The mesophyll cells are in close proximity to the bundle sheath cells, which enables a rapid and continuous transport of metabolites between the two cell types. This cell&#39;s arrangement is typical of C4 plants and is referred to as Kranz anatomy16. In C3 species, mesophyll cell specialization and disposition can vary, but bundle sheath cells are distinctly smaller and have a few chloroplasts or no chloroplasts at all. Specific Kranz anatomy allows concentrating CO2 in chloroplasts in bundle sheath cells where the C3-carboxylating enzyme Rubisco is located, effectively hindering photorespiration4, 17, 18. Despite its seemingly complex arrangement, these changes have occurred independently multiple times in the evolution of angiosperms, indicating that it is a relatively feasible evolutionary pathway19,20,21, and various taxa have been shown to be at an intermediate stage between C3 and C4 carbon metabolism, referred to as C3-C4 or C2, having abilities to concentrate and re-assimilate CO222,23,24,25.
Many C4 plants are crops with high economic importance, and genetically engineering C3 crops, like rice, to improve their climate resilience and secure the yield has been a topic of interest in the last decades26,27. However, the engineering efforts call for a detailed understanding of C4 specialized anatomy and how it controls photosynthesis2,28.
Establishing the C4 Kranz anatomy is a prerequisite to achieving the ambitious goal of engineering C4 photosynthesis into C3 crops25. However, the current understanding of the regulation of Kranz anatomy and methods for quickly screening its key anatomical traits is limited, making it difficult to identify hybrid species. Previous studies have shown that key traits regulating photosynthetic efficiency in C3 and C4 plants include the interveinal distance, the diameter of the bundle sheath complex, and the size of bundle sheath cells14,29. These traits can be easily screened using free-hand sections and quantitatively analyzed using semi-thin sections. Here, we describe the method of assessing the traits that allow for C3 and C4 anatomical differentiation diagnostics through free-hand cross and light microscopy, namely bundle sheath area, interveinal distance, and vein frequency.

<table><tbody><tr><td>Disodium hydrogen phosphate dihydrate (Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;2H&lt;sub&gt;2&lt;/sub&gt;O) pure</td><td>PENTA, CZ</td><td>10028-24-7</td><td></td></tr><tr><td>Embedding Film, 7.8 mil Thick, 8 x 12.5, (203 x 318mm)</td><td>ACLAR, US</td><td>10501-10</td><td></td></tr><tr><td>Ethanol, abs. 100% a.r.</td><td>Chem-Lab NV, BE</td><td>CL00.0505.1000</td><td>Danger: Highly inflammable liquid and vapour.</td></tr><tr><td>EVOS Invitrogen FL Auto 2 Imaging System</td><td>Thermo Fisher Scientific, US</td><td></td><td></td></tr><tr><td>Flat Embedding PTFE Mold with Metal Frame, 16 cavities</td><td>PELCO, US</td><td>10501</td><td></td></tr><tr><td>Glass vial 2 ml</td><td>VWR Life Science, US</td><td>548-0045</td><td></td></tr><tr><td>Glutaraldehyde 50% solution</td><td>VWR Life Science, US</td><td>23H2856331</td><td>Danger: Fatal if inhaled. Toxic if swallowed. Causes severe skin burns and eye damage. May cause respiratory irritation. Wear protective gloves, protective clothing, eyes and face protection.</td></tr><tr><td>Histo diamond knife</td><td>Diatome, US</td><td></td><td></td></tr><tr><td>LEICA EM UC7</td><td>Leica Vienna, AT</td><td></td><td></td></tr><tr><td>LR White resin hard grade</td><td>Electron Microscopy Sciences, US</td><td>14383</td><td>Danger: Causes skin irritation. Causes severe eye irritation May cause respiratory irritation. May cause drowsiness or dizziness Wear protective gloves, protective clothing, eyes and face protection.</td></tr><tr><td>Microscope slides</td><td>Normax, PT</td><td>5470308A</td><td></td></tr><tr><td>Nikon Eclipse E600 and Nikon DS0Fi1 5 MP</td><td>Nikon Corporation, JP</td><td></td><td></td></tr><tr><td>Osmium Tetroxide (OsO&lt;sub&gt;4&lt;/sub&gt;)</td><td>Agar Scientific Ltd, GB</td><td>R1019</td><td>Danger: Fatal if swallowed, in contact with skin or if inhaled. Causes severe skin burns and eye damage Wear double protective gloves, protective clothing, eyes and face protection.</td></tr><tr><td>Pipette and pipette tips</td><td>Thermo Scientific, FI</td><td>KJ16047</td><td></td></tr><tr><td>Sodium dihydrogen phosphate dihydrate (NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4 &lt;/sub&gt;&lt;sup&gt;.&lt;/sup&gt; 2H&lt;sub&gt;2&lt;/sub&gt;O) pure</td><td>PENTA, CZ</td><td>13472-35-0</td><td></td></tr><tr><td>Syringe 10 ml</td><td>Ecoject, DE</td><td>20010</td><td></td></tr><tr><td>Toluidine blue, general purpose grade</td><td>Fisher Scientific, GB</td><td>2045836</td><td></td></tr></tbody></table>

assessing structural traits in triticum aestivum and zea mays for c3 and c4 photosynthetic differentiation using free-hand and semi-thin sections

The enhanced efficiency of C4 photosynthesis, compared to the C3 mechanism, arises from its ability to concentrate CO2 in bundle sheath cells. The effectiveness of C4 photosynthesis and intrinsic water use efficiency are directly linked to the share of mesophyll and bundle leaf cells,&#160;size and density of bundle sheaths, and size, density, and cell wall thickness of bundle sheath cells. Rapid microscopical analysis of these traits can be performed on free-hand and semi-thin sections using conventional light microscopy, providing valuable information about photosynthetic efficiency in C4 crops by means of identifying and examining specific cell types. Additionally, errors in freehand and semi-thin section preparation that affect anatomical measurements and cell type diagnoses are shown, as well as how to avoid these errors. This microscopical approach offers an efficient means of gathering insights into photosynthetic acclimation to environmental variation and aids in the rapid screening of crops for future climates.

Figure 1A shows the correct orientation for sectioning the leaf for both fresh sectioning and light microscopy. The method for cutting fresh sections using a single-sided razor and a dental wax sheet can be seen in Figure 1B. The resulting sections are shown in Figure 1C.
Figure 2 shows free-hand sections of representative leaves of the C3 plant T. aestivum</em...

Author Spotlight: Innovative Approaches to Understanding Plant Structure-Function Relationships for Climate-Resilient Crops

Assessing the anatomical differences between C3 and C4 leaf cross sections helps understand photosynthesis efficiency. This paper describes free-hand and semi-thin leaf cross sections preparation and examination, along with the caveats in the preparation for the crop species Triticum aestivum and Zea mays.

Assessing Structural Traits in Triticum aestivum and Zea mays for C3 and C4 Photosynthetic Differentiation Using Free-hand and Semi-thin Sections

The enhanced efficiency of C4 photosynthesis, compared to the C3 mechanism, arises from its ability to concentrate CO2 in bundle sheath cells. The ...

assessing-structural-traits-triticum-aestivum-zea-mays-for-c3-c4

Research

JoVE Journal

Biology

842 Views.  Estonian University of Life Sciences. Ons onderzoek richt zich op plantstructuur, functierelaties. We willen een beter inzicht krijgen in de beperkingen van fotosynthese tussen functionele planttypen en hoe deze kunnen worden verbeterd voor klimaatbestendige gewassen. Nieuwe technologie en software zijn duur en vaak ontoegankelijk, wat een zekere mate van proef en innovatie vereist om dezelfde resoluties te bereiken bij het afbeelden van verschillende soorten fabrieksfuncties.De identifica...

Video: Auteur in de schijnwerpers: innovatieve benaderingen voor het begrijpen van plantstructuur-functierelaties voor klimaatbestendige gewassen

RGB and Spectral Root Imaging for Plant Phenotyping and Physiological Research: Experimental Setup and Imaging Protocols

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop cultivars against environmental stresses. An experimental protocol is presented for RGB and hyperspectral imaging of root systems. The approach uses rhizoboxes where plants grow in natural soil over a longer time to observe fully developed root systems. Experimental settings are exemplified for assessing rhizobox plants under water stress and studying the role of roots. An RGB imaging setup is described for cheap and quick quantification of root development over time. Hyperspectral imaging improves root segmentation from the soil background compared to RGB color based thresholding. The particular strength of hyperspectral imaging is the acquisition of chemometric information on the root-soil system for functional understanding. This is demonstrated with high resolution water content mapping. Spectral imaging however is more complex in image acquisition, processing and analysis compared to the RGB approach. A combination of both methods can optimize a comprehensive assessment of the root system. Application examples integrating root and aboveground traits are given for the context of plant phenotyping and plant physiological research. Further improvement of root imaging can be obtained by optimizing RGB image quality with better illumination using different light sources and by extension of image analysis methods to infer on root zone properties from spectral data.

An experimental protocol is presented for assessment of soil grown plant root systems with RGB and hyperspectral imaging. Combination of RGB image time series with chemometric information from hyperspectral scans optimizes insights into plant root dynamics.

Better understanding of plant root dynamics is essential to improve resource use efficiency of agricultural systems and increase the resistance of crop ...

Environment

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is needed to open up the structure of the plant cell wall, increasing the accessibility of the cell wall carbohydrates. Lignin, a polyphenolic material present in many cell wall types, is known to be a significant hindrance to enzyme access. Reduction in lignin content to a level that does not interfere with the structural integrity and defense system of the plant might be a valuable step to reduce the costs of bioethanol production. In this study, we have genetically down-regulated one of the lignin biosynthesis-related genes, cinnamoyl-CoA reductase (ZmCCR1) via a double stranded RNA interference technique. The ZmCCR1_RNAi construct was integrated into the maize genome using the particle bombardment method. Transgenic maize plants grew normally as compared to the wild-type control plants without interfering with biomass growth or defense mechanisms, with the exception of displaying of brown-coloration in transgenic plants leaf mid-ribs, husks, and stems. The microscopic analyses, in conjunction with the histological assay, revealed that the leaf sclerenchyma fibers were thinned but the structure and size of other major vascular system components was not altered. The lignin content in the transgenic maize was reduced by 7-8.7%, the crystalline cellulose content was increased in response to lignin reduction, and hemicelluloses remained unchanged. The analyses may indicate that carbon flow might have been shifted from lignin biosynthesis to cellulose biosynthesis. This article delineates the procedures used to down-regulate the lignin content in maize via RNAi technology, and the cell wall compositional analyses used to verify the effect of the modifications on the cell wall structure.

Lignin Down-regulation of Zea mays via dsRNAi and Klason Lignin Analysis

A double stranded RNA interference (dsRNAi) technique is employed to down-regulate the maize cinnamoyl coenzyme A reductase (ZmCCR1) gene to lower plant lignin content. Lignin down-regulation from the cell wall is visualized by microscopic analyses and quantified by the Klason method. Compositional changes in hemicellulose and crystalline cellulose are analyzed.

To facilitate the use of lignocellulosic biomass as an alternative bioenergy resource, during biological conversion processes, a pretreatment step is ...

Bioengineering

Transverse Sectioning of Mature Rice (Oryza sativa L.) Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm phenotyping can be used to classify genotypes based on SG morphotype using scanning electron microscopic (SEM) analysis. SGs can be visualized using SEM by slicing through the kernel (pericarp, aleurone layers, and endosperm) and exposing the organellar contents. Current methods require the rice kernel to be embedded in plastic resin and sectioned using a microtome or embedded in a truncated pipette tip and sectioned by hand using a razor blade. The former method requires specialized equipment and is time-consuming, while the latter introduces a new host of problems depending on rice genotype. Chalky rice varieties, particularly, pose a problem for this type of sectioning due to the friable nature of their endosperm tissue. Presented here is a technique for preparing translucent and chalky rice kernel sections for microscopy, requiring only pipette tips and a scalpel blade. Preparing the sections within the confines of a pipette tip prevents rice kernel endosperm from shattering (for translucent or &#8216;vitreous&#8217; phenotypes) and crumbling (for chalky phenotypes). Using this technique, endosperm cell patterning and the structure of intact SGs can be observed.

Transverse Sectioning of Mature Rice Oryza sativa L. Kernels for Scanning Electron Microscopy Imaging Using Pipette Tips as Immobilization Support

This protocol allows for the preparation of transverse sections of cereal seeds (e.g., rice) for the analysis of endosperm and starch granule morphology using scanning electron microscopy.

Starch granules (SGs) exhibit different morphologies depending on the plant species, especially in the endosperm of the Poaceae family. Endosperm ...

Biochemistry

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 ...

Hybrid-Cut: An Improved Sectioning Method for Recalcitrant Plant Tissue Samples

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some plant cells have rigid cell walls, tough fibers and crystals (calcium oxalate, silica, etc.), and high water content that often disrupt tissue integrity during plant tissue sectioning.
This study establishes a simple Hybrid-Cut tissue sectioning method. This protocol modifies a paraffin-based sectioning technique and improves the integrity of tissue sections from different plants. Plant tissues were embedded in paraffin before sectioning in a cryostat at -16 &#176;C. Sectioning under low temperature hardened the paraffin blocks, reduced tearing and scratching, and improved tissue integrity significantly. This protocol was successfully applied to calcium oxalate-rich Phalaenopsis orchid tissues as well as recalcitrant tissues such as reproductive organs and leaves of rice, maize, and wheat. In addition, the high quality of tissue sections from Hybrid-Cut could be used in combination with in situ hybridization (ISH) to provide spatial expression patterns of genes of interest. In conclusion, this protocol is particularly useful for recalcitrant plant tissue containing high crystal or silica content. Good quality tissue sections enable morphological and other biological studies.

This protocol describes a simple Hybrid-Cut tissue sectioning method that is useful for recalcitrant plant tissues. Good quality tissue sections enable anatomical studies and other biological studies including in situ hybridization (ISH).

Maintaining plant section integrity is essential for studying detailed anatomical structures at the cellular, tissue, or even organ level. However, some ...

Micron-scale Phenotyping Techniques of Maize Vascular Bundles Based on X-ray Microcomputed Tomography

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a 'sample preparation protocol' for maize materials (i.e., stem, leaf, and root) suitable for ordinary microcomputed tomography (micro-CT) scanning. Based on high-resolution CT images of maize stem, leaf, and root, we describe two protocols for the phenotypic analysis of vascular bundles: (1) based on the CT image of maize stem and leaf, we developed a specific image analysis pipeline to automatically extract 31 and 33 phenotypic traits of vascular bundles; (2) based on the CT image series of maize root, we set up an image processing scheme for the three-dimensional (3-D) segmentation of metaxylem vessels, and extracted two-dimensional (2-D) and 3-D phenotypic traits, such as volume, surface area of metaxylem vessels, etc. Compared with traditional manual measurement of vascular bundles of maize materials, the proposed protocols significantly improve the efficiency and accuracy of micron-scale phenotypic quantification.

We provide a novel method to improve the X-ray absorption contrast of maize tissue suitable for ordinary microcomputed tomography scanning. Based on CT images, we introduce a set of image-processing workflows for different maize materials to effectively extract microscopic phenotypes of vascular bundles of maize.

It is necessary to accurately quantify the anatomical structures of maize materials based on high-throughput image analysis techniques. Here, we provide a ...

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 ...

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 ...

Improved Methods for Imaging Maize Leaf Primordia

Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study early leaf development. Here, we describe methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging. The first method uses a wire stripper to remove the upper portions of older leaves, exposing the tip of the leaf primordium and allowing its measurement for more accurate transverse section sampling. The second method uses clear, double-sided nano tape to unroll and mount whole-leaf primordia for imaging. We show the utility of the two methods in visualizing and analyzing fluorescent protein reporters in maize. These methods provide a solution to the challenges presented by the distinctive morphology of maize leaf primordia and will be useful for visualizing and quantifying leaf anatomical and developmental traits in maize and other grass species.

Author Spotlight: Improved Methods for Preparing Transverse Sections and Unrolled Whole Mounts of Maize Leaf Primordia for Fluorescence and Confocal Imaging  

Maize leaf primordia are deeply ensheathed and rolled, making them difficult to study. Here, we present methods for preparing transverse sections and unrolled whole mounts of maize leaf primordia for fluorescence and confocal imaging.

In maize (Zea mays) and other grasses (Poaceae), the leaf primordia are deeply ensheathed and rolled within the leaf whorl, making it difficult to study ...

Assessing Structural Traits in Triticum aestivum and Zea mays for C₃ and C₄ Photosynthetic Differentiation Using Free-hand and Semi-thin Sections

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