We acknowledge the U.S. Department of Agriculture (Grant 58-6406-1-017) for supporting this research. We also acknowledge the WKU Biotechnology Centre, Western Kentucky University, Bowling Green, KY, USA, and the Director, CSIR Central Institute of Medicinal and Aromatic Plants, Lucknow, India, for providing the instrument facilities and support (CSIR CIMAP manuscript communication no. CIMAP/PUB/2022/103).&#160;SS acknowledges the financial support from Saint Joseph&#39;s University, Philadelphia, USA.

The whole protocol is summarized schematically in Figure 1, showing all the essential steps involved in revealing the root system architecture (RSA) of plantlets. Protocol steps are given in detail below:
1. Arabidopsis seed surface sterilization
<ol>
	<li>Transfer a tiny scoop (approximately 100 seeds = approximately 2.5 mg) of seeds to a microfuge tube, and soak for 30 min in distilled water at room temperature (RT). This entire procedure is carried out in the aseptic condition.</li>
	<li>Briefly centrifuge the microfuge tube containing seeds at 500 x g for 5 s, using any tabletop centrifuge at RT to let the seeds settle down.</li>
	<li>Decant the water, add 700 &#181;L of 70% (v/v) ethanol, vortex for a few seconds, and spin. Repeat vortexing and spinning if required, but ensure the treatment time of 70% ethanol remains at 3 min.</li>
	<li>After 3 min, immediately rinse once with sterile water. Keep the ethanol washing step as timely as possible, as prolonged ethanol exposure decreases germination.</li>
	<li>Treat the seeds with diluted commercial bleach (4% v/v) with a drop of Tween-20 for 7 min. Mix the seeds with bleach solution by inverting the tubes rapidly 8-12 times, followed by a brief centrifuge (500 x g for 5 s at RT). Froth is seen appearing in the tube.</li>
	<li>Decant the supernatant using a 1 mL pipette and rinse the seeds with at least five washes with sterile water, following the same vortexing procedure.</li>
	<li>Leave the surface sterilized seeds in water and incubate for 2-3 days at 4 &#176;C for stratification10.</li>
</ol>
2. Setting a hydroponic system for seed germination
<ol>
	<li>Half-fill a standard magenta box with distilled water and autoclave it. Autoclave the polycarbonate sheet (clear color and smooth texture) and cut 4 cm x 8 cm rectangles, with a midpoint notched more than halfway through the rectangle so that two rectangles may slot together to form an X shape10. Use this setup to hold the polypropylene mesh (6 cm x 6 cm squares of 250 &#181;m pore size, or depending upon the requirement) cut from 12x 24 inch sheets10. 
	NOTE: Polypropylene is highly resistant to acids, alkalis, and other chemicals; therefore, it has been opted. Autoclaving tends to distort polypropylene mesh; hence, it is recommended to be carried separately wrapped in aluminum foil. Typical autoclaving conditions of 16 min, 121 &#176;C, 15 psi, or 775 mm Hg are recommended.</li>
	<li>Add sterile half-MS basal media with vitamins + 1.5% (w/v) sucrose, as described by Shukla et al.1, to each box to reach the bottom edge of the polypropylene mesh in a laminar flow. All the procedures are carried out under aseptic conditions.</li>
	<li>Sow the surface-sterilized seeds on the mesh (250 &#181;m pore size) hydroponically and allow them to grow for 3 days.</li>
	<li>After 3 days, transfer the seedlings onto a mesh (500 &#181;m pore size) and allow them to grow for 2 days.</li>
	<li>After 2 days (total of 5 days), transfer the seedlings onto the control media (i.e., modified MS nutrient media1 containing 2.0 mM NH4NO3, 1.9 mM KNO3, 0.15 mM MgSO4&#183;7H2O, 0.1 mM MnSO4&#183;H2O, 3.0 &#181;M ZnSO4&#183;7H2O, 0.1 &#181;M CuSO4&#183;5H2O, 0.3 mM CaCl2&#183;2H2O, 5.0 &#181;M KI, 0.1 &#181;M CoCl2&#183;6H2O, 0.1 mM FeSO4&#183;7H2O, 0.1 mM Na2EDTA&#183;2H2O, 1.25 mM KH2PO4, 100 &#181;M H3BO3, 1 &#181;M Na2MoO4&#183;2H2O, 1.5% sucrose, 1.25 mM MES, pH 5.7 adjusted with 0.1 M MES [pH 6.1]) and to the experimental media (e.g., P- [0 mM] treatment; KH2PO4 is replaced with 0.62 mM K2SO4 from the control media composition as mentioned above1. For excess Pi treatments, the concentration of KH2PO4 is increased in modified MS medium [2.5, 5.0, 10.0, 20.0 mM]1) and let the seeds grow for 7 days. 
	&#8203;NOTE: A larger mesh pore size (500 &#181;m) facilitates the smooth picking of entire seedlings out without any damage or need of cutting at the hypocotyl. Plantlets grow under standard growth conditions (i.e., 16 h light/8 h dark photoperiod, 150 &#181;mol&#183;m-2&#183;s-1 light intensity, 60%-70% humidity) at 23 &#176;C.</li>
</ol>
3. Examination of RSA
<ol>
	<li>Prepare agar (1.1%) plates for root spreading (Petri plate size: 150 mm x 15 mm).</li>
	<li>Add 10-20 mL of autoclaved filtered tap water to the Petri plate, as mentioned above. Gently pull out the seedlings from the mesh (500 &#181;m) and submerge them in water on the plates.</li>
	<li>Gently spread each plantlets&#39; root in the water-filled plate with the help of a round watercolor art brush (sizes: no. 14, 16, 18, and 20). 
	NOTE: While carrying out the spreading of the root system, first get hold of the primary root and spread it into a straight line, as it serves as an axis. Then, spread the LRs symmetrically on each side of the primary root, wherever possible. After that, spread the second-order LR linked to the first-order LR. This spreading process is a kind of art; do it gently, slowly, like an artist drawing an image of the RSA.</li>
	<li>Tilt the plate slightly to remove the water. 
	&#8203;NOTE: At this point, the procedure can be paused by putting these spread plates at 4 &#176;C. Later, when image processing is required, take out the plates and place them at RT for a while. Wipe out the condensed water, and then the image can be processed conveniently.</li>
</ol>
4. Recording images for RSA
<ol>
	<li>Scan or photograph these Petri plates appropriately. 
	NOTE: For obtaining high-quality photographs, the 600 dpi resolution is recommended for scanning, and at least a 12 megapixel camera is recommended for photography.</li>
	<li>Measure the root system architecture traits using freely available ImageJ software (https://imagej.nih.gov/ij/index.html). To quickly follow the steps to measure the root length using ImageJ software, please refer to the example &#34;measuring DNA contour length&#34;&#8203;17. 
	NOTE: These steps are followed to measure the root lengths on pictures captured using a high-resolution scanner or camera.
	<ol>
		<li>Use a given distance of length to set the scale. The known distance of the scale bar in Figure 3 is 2 cm. Select the Straight Line tool from the ImageJ toolbar (fifth tool from the left). Use the Straight-Line tool to create a line selection that outlines the scale bar. Finish outlining by right-clicking, double-clicking, or clicking in the box at the beginning.</li>
		<li>Measure the length of the known scale bar in pixels using the Analyze &#62; Measure toolbar. Make a note of the pixel length.</li>
		<li>Open the Set Scale dialogue box by clicking the Set Scale tab in the Analyze tab. In the Distance in Pixels field, enter the pixel length (as noted above). Next, in the Known Distance field, enter the value, as shown by the scale bar (here, it is 20 mm). Set the Unit of Length as mm. The pixel-aspect ratio is 1.0. Now, the scale is specified by the x number of pixels per millimeter. To lock the scale for this particular image, click on OK.</li>
		<li>Create a line selection that outlines the root length using the Segmented Line tool. Finish outlining by right-clicking, double-clicking, or clicking in the box at the beginning. Click and drag the small black and white &#34;handles&#34; along the outline to adjust the line selection as needed.</li>
		<li>Use the Measure command under the Analyze tab of ImageJ to quantify the length of the root. To transfer the measured data to a spreadsheet, right-click on the Results window, select Copy All from the popup menu, switch to the spreadsheet, and then paste the data. 
		NOTE: As described above, set the scale using the known distance of the scale bar in the ImageJ set scale option. This gives the number of pixels per unit length. It is required to freshly set the scale every time, whenever a new image is being analyzed.</li>
	</ol>
	</li>
	<li>Measurement and calculation of RSA traits
	<ol>
		<li>Measure the primary root length between the hypocotyl junction to the root tip&#39;s end.</li>
		<li>Measure the first- and second-order LR length.</li>
		<li>Measure the branching zone (BZ) of the primary root. The branching zone of the primary root (BZPR) spans the first LR emerging point to the last LR emerging point.</li>
		<li>Record the number of LRs, which is the number of LRs originating within the boundary of the BZPR.</li>
		<li>Measure the average length of the first- and higher-order LRs. Derive the average length of the first-order LR (1&#176; LR) (centimeter per root) by dividing the total length of the 1&#176; LR by the total number of 1&#176; LRs.</li>
		<li>Measure the average length of the second-order LR. Calculate the average length of the second-order LR (2&#176; LR) by dividing the total length of the 2&#176; LR by the total number of 2&#176; LRs.</li>
		<li>Measure the 1&#176; LR density. Calculate the 1&#176; LR density (number of 1&#176; LRs per unit length of BZPR) by dividing the number of 1&#176; LRs by the length of the BZPR.</li>
		<li>Measure the 2&#176; LR density. Calculate the 2&#176; LR density by dividing the number of 2&#176; LRs by the length of the BZ of 1&#176; LRs (number of 2&#176; LRs per unit length of the BZ of 1&#176; lateral roots).</li>
		<li>Measure the total root length (TRL). This is the aggregate of the primary root, 1&#176; LR, and 2&#176; LR (and more if present) lengths.</li>
	</ol>
	</li>
</ol>
5. Root hair measurement
NOTE: Although the hydroponic system is not good at promoting root hair growth and development, despite being as robust as it is in solid growth media, it is still important to study it in the present context. Follow the steps below to analyze root hair development in a 5 mm section from the tip of the primary root of the seedlings.
<ol>
	<li>Chop off a 2 cm section of the primary root from the root tip.</li>
	<li>Mount the root section on a slide using 10% glycerol as a mounting medium.</li>
	<li>Place the slide under a stereo microscope.</li>
	<li>Use the axial carrier to visualize and capture images of the root hairs.</li>
	<li>Analyze the images to study the structure and characteristics of the root hairs using ImageJ software as earlier described.</li>
</ol>

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The authors declare no conflict of interest.

This work demonstrated mapping RSA utilizing simple laboratory equipment. Using this method, phenotypic alterations are recorded at the refined level. The benefit of this strategy is that the shoot portion never comes in contact with the media, so the phenotype of the plantlets is original. This method involves setting up a hydroponic system to grow plantlets as described in the protocol. Then, each plantlet is taken out intact and placed on an agar-filled Petri plate. The root system is then allowed to spread manually u...

The root system architecture (RSA), which is underground, is a vital organ for plant growth and productivity1,2,3. After the embryonic stage, plants undergo their most significant morphological changes. The way in which the roots grow in the soil greatly affects the growth of plant parts above ground. Root growth is the first step in germination. It is an informative trait as it uniquely responds to different available nutrients1,2,3,4. The RSA exhibits a high degree of developmental plasticity, which means that the environment is always used to make decisions about development2,5. Changes in the environment have made crop production more difficult in the present scenario. On a continuous basis, the RSA incorporates environmental signals into developmental choices5. As a result, a thorough understanding of the principles behind root development is essential for learning how plants respond to changing environments2,5.
The RSA senses varying nutrient concentrations and renders phenotypic alterations4,6,7,8,9,10,11,12. Studies suggest that root morphology/RSA is highly plastic compared to shoot morphology1,3. RSA trait mapping is highly effective in recording the effect of changing the surrounding soil environment1,11,12.
In general, discrepancies in the effect of various nutrient deficiencies on the root phenotype have been reported in many earlier studies3,11,13,14,15. For example, there are several contrasting reports on phosphate (Pi) starvation-induced changes in the number, length, and density of lateral roots (LRs). An increase in LR density has been reported under the Pi deficient condition6,8. In contrast, a decrease in LR density under Pi deficient conditions has also been reported by other authors3,13,16. One of the prominent causes of these inconsistencies is the use of the elemental contamination-prone gelling medium, which agar often contains10. Researchers typically grow their experimental plants on an agar-based plate system and record the root traits. Numerous RSA traits are frequently concealed or entrenched within the agar material and cannot be documented. Experiments linked to inducing nutrient deficiency, in which users often exclude one component totally from the medium, cannot be performed in elemental contamination-prone gelling medium11,14,15. Numerous nutrients are frequently present in significant amounts in the agar media, including P, Zn, Fe, and many more11,14,15. Furthermore, RSA growth is slower in agar-based media than in non-agar-based liquid medium. As a result, there is a need to establish an alternate non-agar-based approach for quantifying and qualitatively recording the phenotype of RSA. Consequently, the current method has been developed, in which plantlets are raised in a magenta box-based hydroponic system atop a polypropylene mesh supported by polycarbonate wedges1,10,11.
This study presents a detailed improvised version of the earlier method described by Jain et al.10. This strategy has been tuned for current demands in plant root biology and can also be used for plants like Alfalfa, other than model plants. The protocol is the primary way to measure the changes in RSA, and it only requires simple equipment. The present protocol illustrates how to phenotype several root features, such as primary and lateral roots in normal and modified medium (Pi deficient). Step-by-step directions and other helpful hints gleaned from the author&#39;s experiences are provided to help the researchers follow along with the methodologies offered in this method. The present study aims to provide a simple and effective method for revealing the entire root system of plants, including higher-order LRs. This method involves manually spreading the root system with a round watercolor art brush, allowing for precise control over the exposure of the roots1,10,11,12. It does not require expensive equipment or complicated software. This method has improved nutrient uptake and growth rate; plants have a nutrient-rich solution easily absorbed by their roots. The present method is suitable for researchers who wish to map the traits of a plant&#39;s root system in detail, particularly during early development (10-15 days after germination). It is suitable for small root systems, model plants like Arabidopsis and tobacco, and non-conventional plants like Alfalfa until their root system fits in the magenta boxes.
The steps for phenotypic analysis of RSA development in Arabidopsis are outlined in this protocol as follows: (1) the method of seed surface sterilization for plants (Arabidopsis), (2) the steps to set up the hydroponic system, followed by seed sowing on a medium, (3) procedure for taking out the complete seedings and spreading on the Petri plate for RSA analysis, (4) how to record the images for RSA, and (5) calculate important RSA parameters using ImageJ software.

<table><tbody><tr><td>Arabidospsis thaliana (Col 0)</td><td>Lehle Seeds</td><td>WT-02</td><td>Columbia (Col-0**, no markers)*</td></tr><tr><td>Art brushes</td><td>Amazon or any other vendor</td><td></td><td>Water color round brush size no. 14 (8 mm), 16 (9.5 mm), 18 (12 mm), and 20 (14.2 mm)</td></tr><tr><td>Automated Microscope with digital camera</td><td>Leica Microsystems</td><td></td><td>LAS version 4.12.0, Leica Microsystems</td></tr><tr><td>Imaging Software</td><td>ImageJ</td><td></td><td>ImageJ V&lt;br/&gt; &amp;nbsp;1.8.0</td></tr><tr><td>Magenta box GA-7</td><td>Fisher Scientific&amp;nbsp;</td><td>50-255-176</td><td></td></tr><tr><td>Medicago sativa</td><td>Johnny&#039;s Seeds</td><td></td><td></td></tr><tr><td>Petri-plate (150 mm x 15 mm)</td><td>USA Scientific</td><td>8609-0215</td><td>150 mm x 15 mm PS Petri Dish (https://www.usascientific.com)</td></tr><tr><td>Photo camera</td><td>Cannon or Nikon</td><td></td><td>Any high mega pixel (atleast 12 mega pixel per inch) camera on macro mode</td></tr><tr><td>Plant-Agar</td><td>Sigma-Aldrich</td><td>A3301</td><td>Agargel&amp;nbsp; Suitable for plant tissue culture</td></tr><tr><td>Polycarbonate Sheets</td><td>Amazon</td><td></td><td>1 mm&amp;nbsp; thick</td></tr><tr><td>Polypropylene Mesh</td><td>Amazon</td><td></td><td>Pore size 250 &amp;micro;m, 500 &amp;micro;m and 1000 &amp;micro;m</td></tr><tr><td>Scanner</td><td>Epson</td><td></td><td>Epson Perfection V700 Photo (Scan at 600 dpi)</td></tr></tbody></table>

a simple protocol for mapping the plant root system architecture traits

Comprehensive knowledge of plant root system architecture (RSA) development is critical for improving nutrient use efficiency and increasing crop cultivar tolerance to environmental challenges. An experimental protocol is presented for setting up the hydroponic system, plantlet growth, RSA spreading, and imaging. The approach used a magenta box-based hydroponic system containing polypropylene mesh supported by polycarbonate wedges. Experimental settings are exemplified by assessing the RSA of the plantlets under varying nutrient (phosphate [Pi]) supply. The system was established to examine the RSA of Arabidopsis, but it is readily adaptable to study other plants like Medicago sativa (Alfalfa). Arabidopsis thaliana (Col-0) plantlets are used in this investigation as an example to understand the plant RSA. Seeds are surface sterilized by treating ethanol and diluted commercial bleach, and kept at 4 &#176;C for stratification. The seeds are germinated and grown on a liquid half-MS medium on a polypropylene mesh supported by polycarbonate wedges. The plantlets are grown under standard growth conditions for the desired number days, gently picked out from the mesh, and submersed in water-containing agar plates. Each root system of the plantlets is spread gently on the water-filled plate with the help of a round art brush. These Petri plates are photographed or scanned at high resolution to document the RSA traits. The root traits, such as primary root, lateral roots, and branching zone, are measured using the freely available ImageJ software. This study provides techniques for measuring plant root characteristics in controlled environmental settings. We discuss how to (1) grow the plantlets, and collect and spread root samples, (2) obtain pictures of spread RSA samples, (3) capture the images, and (4) use image analysis software to quantify root attributes. The advantage of the present method is the versatile, easy, and efficient measurement of the RSA traits.

The different morphometric traits of root system architecture (RSA) are measured using simple laboratory tools, and the steps are depicted schematically in Figure 1. The details of the hydroponic setup demonstrate the protocol&#39;s potential in measuring the RSA (Figure 1 and Figure 2).
Given the observed differences in gelling agents, we used a hydroponic growing system to conduct all the studies<sup cl...

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A Simple Protocol for Mapping the Plant Root System Architecture Traits

We use simple laboratory tools to examine the root system architecture (RSA) of Arabidopsis and Medicago. The plantlets are grown hydroponically over mesh and spread using an art brush to reveal the RSA. Images are taken using scanning or a high-resolution camera, then analyzed with ImageJ to map traits.

Comprehensive knowledge of plant root system architecture (RSA) development is critical for improving nutrient use efficiency and increasing crop cultivar ...

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Research

JoVE Journal

Biology

2.6K Views.  Western Kentucky University. We use simple laboratory tools to examine the root system architecture (RSA) of Arabidopsis and Medicago. The plantlets are grown hydroponically over mesh and spread using an art brush to reveal the RSA. Images are taken using scanning or a high-resolution camera, then analyzed with ImageJ to map traits.

Video: A Simple Protocol for Mapping the Plant Root System Architecture Traits

Extracting Metrics for Three-dimensional Root Systems: Volume and Surface Analysis from In-soil X-ray Computed Tomography Data

Plant roots play a critical role in plant-soil-microbe interactions that occur in the rhizosphere, as well as processes with important implications to climate change and crop management. Quantitative size information on roots in their native environment is invaluable for studying root growth and environmental processes involving plants. X-ray computed tomography (XCT) has been demonstrated to be an effective tool for in situ root scanning and analysis. We aimed to develop a costless and efficient tool that approximates the surface and volume of the root regardless of its shape from three-dimensional (3D) tomography data. The root structure of a Prairie dropseed (Sporobolus heterolepis) specimen was imaged using XCT. The root was reconstructed, and the primary root structure was extracted from the data using a combination of licensed and open-source software. An isosurface polygonal mesh was then created for ease of analysis. We have developed the standalone application imeshJ, generated in MATLAB1, to calculate root volume and surface area from the mesh. The outputs of imeshJ are surface area (in mm2) and the volume (in mm3). The process, utilizing a unique combination of tools from imaging to quantitative root analysis, is described. A combination of XCT and open-source software proved to be a powerful combination to noninvasively image plant root samples, segment root data, and extract quantitative information from the 3D data. This methodology of processing 3D data should be applicable to other material/sample systems where there is connectivity between components of similar X-ray attenuation and difficulties arise with segmentation.

A methodology for obtaining visual and quantitative root structure information from X-ray computed tomography data acquired in-soil is presented.

Plant roots play a critical role in plant-soil-microbe interactions that occur in the rhizosphere, as well as processes with important implications to ...

Environment

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, phosphorus, sulfate and trace elements that plants acquire from the soil. If we want to develop sustainable approaches to producing high crop yield, we need to better understand how the root develops, takes up a wide spectrum of nutrients, and interacts with symbiotic and pathogenic organisms. To accomplish these goals, we need to be able to explore roots in microscopic detail over time periods ranging from minutes to days.
We developed the RootChip, a polydimethylsiloxane (PDMS)- based microfluidic device, which allows us to grow and image roots from Arabidopsis seedlings while avoiding any physical stress to roots during preparation for imaging1 (Figure 1). The device contains a bifurcated channel structure featuring micromechanical valves to guide the fluid flow from solution inlets to each of the eight observation chambers2. This perfusion system allows the root microenvironment to be controlled and modified with precision and speed. The volume of the chambers is approximately 400 nl, thus requiring only minimal amounts of test solution.
Here we provide a detailed protocol for studying root biology on the RootChip using imaging-based approaches with real time resolution. Roots can be analyzed over several days using time lapse microscopy. Roots can be perfused with nutrient solutions or inhibitors, and up to eight seedlings can be analyzed in parallel. This system has the potential for a wide range of applications, including analysis of root growth in the presence or absence of chemicals, fluorescence-based analysis of gene expression, and the analysis of biosensors, e.g. FRET nanosensors3.

This article provides a protocol for cultivation of Arabidopsis seedlings in the RootChip, a microfluidic imaging platform that combines automated control of growth conditions with microscopic root monitoring and FRET-based measurement of intracellular metabolite levels.

The root functions as the physical anchor of the plant and is the organ responsible for uptake of water and mineral nutrients such as nitrogen, ...

Bioengineering

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 ...

An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive harvest or expensive equipment. We present a customizable and affordable rhizobox method that allows the non-destructive visualization of root growth over time and is particularly well-suited to studying root plasticity in response to localized resource patches. The method was validated by assessing maize genotypic variation in plasticity responses to patches containing 15N-labeled legume residue. Methods are described to obtain representative developmental measurements over time, measure root length density in resource-containing and control patches, calculate root growth rates, and determine 15N recovery by plant roots and shoots. Advantages, caveats, and potential future applications of the method are also discussed. Although care must be taken to ensure that experimental conditions do not bias root growth data, the rhizobox protocol presented here yields reliable results if carried out with sufficient attention to detail.

Visualizing and measuring root growth in situ is extremely challenging. We present a customizable rhizobox method to track root development and proliferation over time in response to nutrient enrichment. This method is used to analyze maize genotypic differences in root plasticity in response to an organic nitrogen source.

Roots are notoriously difficult to study. Soil is both a visual and mechanical barrier, making it difficult to track roots in situ without destructive ...

Translating Ribosome Affinity Purification (TRAP) to Investigate Arabidopsis thaliana Root Development at a Cell Type-Specific Scale

In this article, we give hands-on instructions to obtain translatome data from different Arabidopsis thaliana root cell types via the translating ribosome affinity purification (TRAP) method and consecutive optimized low-input library preparation.
As starting material, we employ plant lines that express GFP-tagged ribosomal protein RPL18 in a cell type-specific manner by use of adequate promoters. Prior to immunopurification and RNA extraction, the tissue is snap frozen, which preserves tissue integrity and simultaneously allows execution of time series studies with high temporal resolution. Notably, cell wall structures remain intact, which is a major drawback in alternative procedures such as fluorescence-activated cell sorting-based approaches that rely on tissue protoplasting to isolate distinct cell populations. Additionally, no tissue fixation is necessary as in laser capture microdissection-based techniques, which allows high-quality RNA to be obtained.
However, sampling from subpopulations of cells and only isolating polysome-associated RNA severely limits RNA yields. It is, therefore, necessary to apply sufficiently sensitive library preparation methods for successful data acquisition by RNA-seq.
TRAP offers an ideal tool for plant research as many developmental processes involve cell wall-related and mechanical signaling pathways. The use of promoters to target specific cell populations is bridging the gap between organ and single-cell level that in turn suffer from little resolution or very high costs. Here, we apply TRAP to study cell-cell communication in lateral root formation.

Translating Ribosome Affinity Purification TRAP to Investigate Arabidopsis thaliana Root Development at a Cell Type-Specific Scale

Translating ribosome affinity purification (TRAP) offers the possibility to dissect developmental programs with minimal processing of organs and tissues. The protocol yields high-quality RNA from cells targeted with a green fluorescent protein (GFP)-labeled ribosomal subunit. Downstream analysis tools, such as qRT-PCR or RNA-seq, reveal tissue and cell type-specific expression profiles.

In this article, we give hands-on instructions to obtain translatome data from different Arabidopsis thaliana root cell types via the translating ribosome ...

Developmental Biology

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by their natural environment, root hair morphology and function are difficult to study and often excluded from plant research. In recent years, microfluidic platforms have offered a way to visualize root systems at high resolution without disturbing the roots during transfer to an imaging system. The microfluidic platform presented here builds on previous plant-on-a-chip research by incorporating a two-layer device to confine the Arabidopsis thaliana main root to the same optical plane as the root hairs. This design enables the quantification of root hairs on a cellular and organelle level and also prevents z-axis drifting during the addition of experimental treatments. We describe how to store the devices in a contained and hydrated environment, without the need for fluidic pumps, while maintaining a gnotobiotic environment for the seedling. After the optical imaging experiment, the device may be disassembled and used as a substrate for atomic force or scanning electron microscopy while keeping fine root structures intact.

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

This article demonstrates how to culture Arabidopsis thaliana seedlings in a two-layer microfluidic platform that confines the main root and root hairs to a single optical plane. This platform can be used for real-time optical imaging of fine root morphology as well as for high-resolution imaging by other means.

Root hairs increase root surface area for better water uptake and nutrient absorption by the plant. Because they are small in size and often obscured by ...

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories can face is the development and attainment of these methods. Observation of phenotypes in a process of interest is a typical objective of research labs studying gene function and this is often achieved through image capture. A particular process that is amenable to observation using imaging approaches is the corrective growth of a seedling root that has been displaced from alignment with the gravity vector. Imaging platforms used to measure the root gravitropic response can be expensive, relatively low in throughput, and/or labor intensive. These issues have been addressed by developing a high-throughput image capture method using inexpensive, yet high-resolution, flatbed scanners. Using this method, images can be captured every few minutes at 4,800 dpi. The current setup enables collection of 216 individual responses per day. The image data collected is of ample quality for image analysis applications.

This protocol describes a process for rapid collection of images of Arabidopsis seedlings responding to a gravity stimulus using commercially-available flatbed scanners. The method allows for inexpensive, high-volume capture of high-resolution images amenable for downstream analysis algorithms.

Research efforts in biology increasingly require use of methodologies that enable high-volume collection of high-resolution data. A challenge laboratories ...

Lateral Root Inducible System in Arabidopsis and Maize

Lateral root development contributes significantly to the root system, and hence is crucial for plant growth. The study of lateral root initiation is however tedious, because it occurs only in a few cells inside the root and in an unpredictable manner. To circumvent this problem, a Lateral Root Inducible System (LRIS) has been developed. By treating seedlings consecutively with an auxin transport inhibitor and a synthetic auxin, highly controlled lateral root initiation occurs synchronously in the primary root, allowing abundant sampling of a desired developmental stage. The LRIS has first been developed for Arabidopsis thaliana, but can be applied to other plants as well. Accordingly, it has been adapted for use in maize (Zea mays). A detailed overview of the different steps of the LRIS in both plants is given. The combination of this system with comparative transcriptomics made it possible to identify functional homologs of Arabidopsis lateral root initiation genes in other species as illustrated here for the CYCLIN B1;1 (CYCB1;1) cell cycle gene in maize. Finally, the principles that need to be taken into account when an LRIS is developed for other plant species are discussed.

Lateral Root Inducible System in Arabidopsis and Maize

The Lateral Root Inducible System (LRIS) allows for synchronous induction of lateral roots and is presented for Arabidopsis thaliana and maize.

Lateral root development contributes significantly to the root system, and hence is crucial for plant growth. The study of lateral root initiation is ...

Multipronged Phenotyping Approaches to Characterize Sugarcane Root Systems

Roots are the primary conductors of water and nutrients and play a vital role in sustaining growth and yield under stressful environments. The study of plant roots poses methodological difficulties in in situ assessment and sampling, which is especially true for sugarcane (Saccharum spp.). Traditional methods during the 1920s documented the genotypic variation in sugarcane root systems, after which few studies were reported on sugarcane root traits per se until recently. In addition to morphology, rhizosphere characteristics, including allelopathic effects and/or affinity for microbial symbiosis, determine plant establishment and survival.
Ultimately, root systems define the above-ground productivity of sugarcane. With the impetus for climate-resilient varieties, it is becoming more relevant to explore and utilize the variability in root system traits of sugarcane. This paper describes multipronged approaches for sugarcane root phenotyping, including field excavation by trench sampling, the use of a root core sampler, raised platforms for root sampling, and raising plants under hydroponic culture, employed by a team of scientists at the Indian Council of Agricultural Research-Sugarcane Breeding Institute (ICAR-SBI).
Field excavation by trench sampling is imperative to assess the plant roots in their natural growing environment. The use of raised platforms simulating field conditions and a root core sampler are alternative approaches, with a considerable reduction in time, uniform sample size, and less loss of root material. Hydroponic plant culture allows the study of morphology, anatomical features, and rhizosphere biology, including the exudation of organic compounds and microbial interactions. Data generated from different experiments using diverse sampling methods add to the wealth of information on the root system traits of sugarcane.

Characterizing root system traits is one of the areas of research that is still in its infancy, particularly in sugarcane. Integrating multiple approaches to precisely phenotype sugarcane roots leads to comprehensive and holistic results, enabling the utilization of identified traits and mechanisms for conventional and molecular breeding.

Roots are the primary conductors of water and nutrients and play a vital role in sustaining growth and yield under stressful environments. The study of ...

Mycorrhizal Maps as a Tool to Explore Colonization Patterns and Fungal Strategies in the Roots of Festuca rubra and Zea mays

Arbuscular mycorrhizal fungi are symbionts in the roots of plants. Their role is to sustain host development and maintain the nutritional equilibrium in the ecosystems. The colonization process is dependent on several factors like soil ecology, the genetic diversity of the fungi and host, and agronomic practices. Their synchronized action leads to the development of a complex hyphal network and leads to the secondary development of vesicles and arbuscules in the root cells. The aim of this research was to analyze the efficiency of the mycorrhizal patterns (MycoPatt) method for the positioning of fungal structures in the roots of Festuca rubra and Zea mays. Another objective was to explore the fungal colonization strategy as revealed by mycorrhizal maps of each species. The acquisition and assemblage of multiple microscopic images allow mycorrhizal colonization assessment in both corn and red fescue plants to provide information on the realistic position of the developed structures. The observed mycorrhizal patterns highlight the variable efficiency of each plant in terms of developing connections with soil symbiotic fungi, caused by applied treatments and growth stage. Mycorrhizal detailed maps obtained through the MycoPatt method are useful for the early detection of plant efficiency in symbiotic acquisition from the soil.

Mycorrhizal Maps as a Tool to Explore Colonization Patterns and Fungal Strategies in the Roots of Festuca rubra and Zea mays

The protocol here describes the methods for the assessment of the arbuscular mycorrhizal colonization patterns and strategy in roots for two species: Zea mays and Festuca rubra. The use of the MycoPatt method permits the calculation of parameters, the conversion of mycorrhizal structures into digital data, and the mapping of their real position in roots.

Arbuscular mycorrhizal fungi are symbionts in the roots of plants. Their role is to sustain host development and maintain the nutritional equilibrium in ...

A Simple Protocol for Mapping the Plant Root System Architecture Traits

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Chapters in this video

Extracting Metrics for Three-dimensional Root Systems: Volume and Surface Analysis from In-soil X-ray Computed Tomography Data

Time-lapse Fluorescence Imaging of Arabidopsis Root Growth with Rapid Manipulation of The Root Environment Using The RootChip

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

An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients

Translating Ribosome Affinity Purification (TRAP) to Investigate Arabidopsis thaliana Root Development at a Cell Type-Specific Scale

Imaging the Root Hair Morphology of Arabidopsis Seedlings in a Two-layer Microfluidic Platform

Using Flatbed Scanners to Collect High-resolution Time-lapsed Images of the Arabidopsis Root Gravitropic Response

Lateral Root Inducible System in Arabidopsis and Maize

Multipronged Phenotyping Approaches to Characterize Sugarcane Root Systems

Mycorrhizal Maps as a Tool to Explore Colonization Patterns and Fungal Strategies in the Roots of Festuca rubra and Zea mays