The authors would like to acknowledge anonymous reviewers for their feedback, as well as J.C. Cahill and Tan Bao for initial guidance on developing the rhizobox protocol. Funding was provided by the Foundation for Food and Agriculture Research, the US Department of Agriculture (USDA) National Institute of Food and Agriculture, Agricultural Experiment Station Project CA-D-PLS-2332-H, to A.G. and by the UC Davis Department of Plant Sciences through a fellowship to J.S.

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The authors have nothing to disclose.

The rhizoboxes described in this protocol can be used to answer varied questions in root and rhizosphere science, and have found diverse uses elsewhere10,20,21,22,23,24,25. Other researchers have captured time-lapse images of rhizoboxes21,25,26, some using automated systems22,27. These approaches may be used for quantitative analyses of root length and architecture not possible with tracing methods. Rhizoboxes have also been used to visualize microbial communities with techniques such as fluorescent in situ hybridization (FISH) and micro-autoradiography (MAR)21,22, or to capture spatially explicit patterns of water and nutrient resources with RGB imaging24 or extracellular enzyme activity with zymography11,30. The rhizoboxes presented here are unique from previous designs in that they are relatively large, making it possible to study species with extensive root systems; have a simple single-compartment design; use readily available, inexpensive materials; and are specially designed to study localized patches. The versatility of this rhizobox protocol could allow it to be customized for a range of other applications in root plasticity and rhizosphere interactions. Other nutrients could replace nitrogen in the patches. Immobile nutrients such as phosphorus would subject to lesser leaching, likely making them a good fit for this approach. The rhizoboxes are also well-suited to comparisons of bulk and rhizosphere soil, as the zone of root influence (a long-standing definition for the rhizosphere15) can be more clearly delineated than in pot studies and separated from bulk soil with a razor at harvest. Adapting this method to study rhizosphere processes opens up a broad new range of ways to extend the protocol, including study of both abiotic and biotic interactions23.
The rhizobox method presented here is well-suited to measuring relative differences among genotypes or species in root growth early in development, characterizing relationships among root traits, and exploring the effects of soil characteristics on root growth. Certain steps of the protocol are particularly critical because they affect factors with disproportionate influence on root growth: soil moisture, bulk density, and slope. Watering evenly and equally across boxes is critical given the influence of soil moisture on root growth patterns1,31. Pilot experiments showed that water delivered through drip emitters became evenly distributed after 24-48 h due to capillary action; however, variability among emitters in the volume of water delivered during a given irrigation period was too high to recommend this method using our arrangement. Hand watering slowly and evenly was the best of the techniques tested, but other irrigation methods are certainly possible. Watering to a previously calculated weight ensures that all boxes maintain the same soil moisture content, preventing variability in root growth due to water stress32. The weight of empty rhizoboxes can vary significantly, so calculating ideal weights for individual boxes is important.
As with moisture, establishing even bulk density of the substrate throughout the rhizobox is critical to measure root proliferation responses. Roots grow more extensively in less-dense soil33 and compaction may create channels for preferential water flow, further affecting resource distribution and root growth patterns34. Drying and sieving field soil, thoroughly mixing sand and soil, and moving the funnel back and forth slowly and evenly help create a homogeneous matrix for root growth.
A number of components within this experiment will depend on the research questions of interest as well as the soil type and plant species utilized within the study. The ratio of sand to soil may need to be adjusted for soils of different texture, to ensure that the substrate wets evenly without clumping. Pilot experiments showed that a 1:1 (v/v) soil : sand mixture was superior to 1:2 or 1:3 mixtures for the soil used in this experiment, a very fine sandy loam. The dimensions of the rhizoboxes and duration of the experiment can be adjusted depending on the research question, root traits, and plant species of interest. Maize is a relatively fast growing plant species; we therefore selected larger rhizobox dimensions compared to other rhizobox studies20,35 to provide a sufficient volume of soil that allows the experiment to continue for the optimal duration as plants become increasingly rootbound over time. Finally, the fitting of statistical models to visible and traced root growth must be determined for each study and species of interest, perhaps with pilot experiments. Visible root growth may follow a saturating curve rather than a linear model25,36, and the slope of the regression of visible on harvested roots varies by plant species21.
Roots have an inherent tendency to pursue gravity and avoid light37, but care must be taken to ensure that gravitropism and light avoidance do not bias root growth data. If greenhouse benches are tilted slightly, for example, roots will grow down the slope rather than responding to nutrient patches. Similarly, light gradients in the greenhouse could cause shifts in shoot growth and roots to grow preferentially on one side if the rhizoboxes are not kept completely dark. The light deprivation cases presented in the protocol are an effective means of reducing incident light, but other methods such as wrapping the rhizoboxes in aluminum foil may also work.
The non-destructive rhizobox method presented here facilitates the tracking of roots in situ over time38 and can be implemented by a graduate student with a basic working knowledge of power tools. As such, it offers advantages over destructive harvesting methods such as shovelomics6, which are better suited to studying mature root architecture but do not allow repeated measurements of the same root system over time, and MRI or X-ray computed tomography39,40, which allow high-quality imaging of root-substrate interactions but requires expensive equipment. However, it is not without limitations. Construction of the rhizoboxes can be time-consuming, and ensuring that factors such as moisture, bulk density, slope, and light are as uniform as possible, as described above, is non-trivial. Nonetheless, given sufficient time and attention to detail, the rhizoboxes deliver reliable results and can be reused for many experiments.

Although often overlooked compared to their aboveground counterparts, roots play a critical role in plant nutrient acquisition. Given the substantial carbon cost of root construction and maintenance, plants have evolved mechanisms to develop roots only where foraging is worth the investment. Root systems can thus efficiently and dynamically mine resource patches by proliferating in hotspots, upregulating rates of uptake, and rapidly translocating nutrients to the phloem for further transport1. Plasticity responses can vary widely among plant species or genotypes2,3 and depending on the chemical form of the nutrient involved4,5. Variation in root plasticity should be explored further, as understanding complex root responses to heterogeneous soil resources could inform breeding and management strategies to increase nutrient use efficiency in agriculture.
Despite its necessity and relevance for understanding plant systems, visualizing and quantifying root plasticity at relevant scales poses technical challenges. Excavating the root crown from the soil (&#34;shovelomics&#34;6) is a common method, but fine roots exploit small pores between soil aggregates, and excavation inevitably leads to some degree of loss of these fragile roots. Furthermore, destructive harvest makes it impossible to follow changes in one root system over time. In situ imaging methods such as X-ray computed tomography allow direct visualization of roots and soil resources at high spatial resolution7, but are expensive and require specialized equipment. Hydroponic experiments avoid constraints associated with extracting roots from soil, but root morphology and architecture differ in aqueous media as compared to the mechanical constraints and biophysical complexity of soils8,9. Finally, rhizosphere processes and functions cannot be integrated with developmental plasticity in these artificial media.
We present a protocol for the construction and use of rhizoboxes (narrow, clear-sided rectangular containers) as a low-cost, customizable method to characterize root growth in soil over time. Specially designed frames encourage roots to grow preferentially against the back&#160;panel due to gravitropism, increasing the accuracy of root length measurements. Rhizoboxes are commonly used to study root growth and rhizosphere interactions10,11,12, but the method presented here offers an advantage in simplicity with its single-compartment design and inexpensive materials, and is designed to study root responses to localized nutrients. However, the method could also be adapted to study a range of other root and rhizosphere processes such as intra/interspecies competition, spatial distribution of chemical compounds, microbes, or enzyme activity. Here, we investigate genotypic differences among maize hybrids in response to patches of 15N-labeled legume residue and highlight representative results to validate the rhizobox method.

<table><tbody><tr><td>1.27 cm diameter PVC pipe</td><td>JM Eagle</td><td>530048</td><td>305 cm per box, cut into lengths as specified in the protocol</td></tr><tr><td>PVC side elbows</td><td>Lasco</td><td>315498</td><td>2 per box</td></tr><tr><td>PVC 90-degree elbows</td><td>Charlotte</td><td>PVC 02300 0600</td><td>4 per box</td></tr><tr><td>PVC T joints</td><td>Charlotte</td><td>PVC 02402 0600</td><td>4 per box</td></tr><tr><td>Extruded acrylic panes</td><td>TAP Plastics</td><td>N/A</td><td>2 per box, 0.64 cm thick x 40.5 cm wide x 61 cm long</td></tr><tr><td>HDPE spacers (sides)</td><td>TAP Plastics</td><td>N/A</td><td>2 per box, 0.64 cm thick x 2.5 cm wide x 57 cm long</td></tr><tr><td>HDPE spacers (bottom)</td><td>TAP Plastics</td><td>N/A</td><td>1 per box, 0.64 cm thick x 2.5 cm wide x 40.5 cm long</td></tr><tr><td>HDPE spacers (patch)</td><td>TAP Plastics</td><td>N/A</td><td>2 per box, 0.64 cm thick x 3.8 cm wide x 28 cm long</td></tr><tr><td>Polyester batting</td><td>Fairfield</td><td>#A-X90</td><td>2.5 cm x 40.5 cm strip per box</td></tr><tr><td>20-thread screws</td><td>N/A</td><td>N/A</td><td>3.2 cm long, 0.64 cm diameter</td></tr><tr><td>Washers</td><td>N/A</td><td>N/A</td><td>0.64 cm internal diameter</td></tr><tr><td>Hex nuts</td><td>N/A</td><td>N/A</td><td>sized to fit the screws</td></tr><tr><td>Light deprivation fabric</td><td>Americover, Inc.</td><td>Bold 8WB26.5</td><td>1 piece 95 cm wide and 69 cm long per box</td></tr><tr><td>Sand</td><td>Quikrete</td><td>No. 1113</td><td></td></tr><tr><td>Field soil</td><td>N/A</td><td>N/A</td><td></td></tr><tr><td>Transparencies for tracing</td><td>FXN</td><td>FXNT1319100S</td><td>One per side of the box to be traced</td></tr></tbody></table>

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.

Roots grew preferentially against the back of the box, as anticipated. Total traced root length on the back of the box ranged from 400 to 1,956 cm, as compared to 93-758 cm on the front of the box. Pairwise Pearson correlation coefficients were calculated between scanned root length and traced root length on the front of the box, back of the box, and the sum of front and back was used to determine whether tracing accurately reflected total root length (n = 23, as the plant in one box died during the experiment). Scanned total root length was significantly correlated with traced root length on the back of the box (Figure 5A, p = 0.0059), front of the box (Figure 5B, p = 0.022), and sum of back and front (Figure 5C, p = 0.0036). Tracing only the back of the box is thus validated as giving a representative measure of root growth while halving the time required to trace roots. It should be noted, however, that tracing captures only 21.6-54.6% of total root length. While roots do grow preferentially against the surface of the rhizobox, fine lateral roots in particular may not be visible for tracing. Tracing is well-suited to relative comparisons of root length over time, especially early in development, but harvesting and scanning root systems is preferable if the goal is to accurately quantify total root length.
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58674/58674fig5v2.jpg" /> 
Figure 5: Correlations between traced and scanned root length data. A) Traced root length was significantly correlated with scanned root length when only the front of the box was traced. B) Tracing roots on the back of the box, where the majority of roots were visible, improved the R2 value of the regression against scanned root length over tracing the front of the box; the correlation was again significant. C) The most accurate method is to trace roots on both sides of the box, as shown by the highest R2 value of the three methods and the significant correlation with scanned root length. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58674/58674fig5v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Root growth rates over time were similar among boxes, as shown by consistent slopes when plotting the natural log of total root length against time (Figure 6). While slight variability is to be expected, consistent growth rates indicate that experimental conditions were uniform for all boxes. Dramatically different slopes would indicate that plants were growing at different rates, suggesting the need to check for differences in variables such as temperature or moisture.
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58674/58674fig6.jpg" /> 
Figure 6: Root growth rates over time. Similar slopes of root length vs. time among rhizoboxes indicate that roots grew at equal rates. Non-uniform slopes could indicate that experimental conditions vary among rhizoboxes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58674/58674fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Roots of all maize genotypes proliferated in patches containing 15N-labeled cover crop residue. Two-way ANOVA with patch type and genotype as fixed factors (n = 23) revealed that root length density was higher in treatment than control patches using scanned root data (Figure 7a, p = 0.013) as well as traced root data (Figure 7b, p = 0.005). RLD was not significantly different among genotypes in either case.
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58674/58674fig7.jpg" /> 
Figure 7: Root length density by genotype and root data type. a) Scanned root data showed that all genotypes (A-F) proliferated in the treatment (T) patch, and genotypic differences were not significant. b) Harvested and scanned root data confirmed the significant effect of legume residue, but not genotype (A-F), on root length density in patches. Letters A-F represent six different genotypes and error bars represent standard error. C: control; T: treatment. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58674/58674fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Root diameter can be used to make inferences about root function and turnover. Fine roots are more likely to be lateral roots that rapidly develop and proliferate in response to resource hotspots, while larger coarse roots are more likely to be long-lived, slow-to-respond axial roots. Scanned root systems were analyzed for the proportion of roots in each diameter class: &#60;0.2 mm, 0.2-0.4 mm, 0.4-0.8 mm, 0.8-1.6 mm, and &#62;1.6 mm, and each size class was tested for genotypic differences. Genotypes with more fine roots in treatment patches might indicate a more effective proliferation response. One-way ANOVA with genotype as a fixed factor (n = 23) revealed that genotypes did not differ in root length in each size class for the root system overall (Figure 8a), treatment patches (Figure 8b), or control patches (Figure 8c). A majority of the roots were fine roots (&#60;0.2 mm).
<img alt="Figure 8" class="xfigimg" src="/files/ftp_upload/58674/58674fig8.jpg" /> 
Figure 8: Proportions of roots in different diameter classes by genotype and location. a) In each rhizobox (excluding treatment and control patches), the majority of roots were fine (&#60;0.2mm in diameter). Genotypes did not differ in the proportions of roots in each diameter class. b) In treatment patches, root length per size class likewise decreased with increasing diameter across genotypes. c) Control patches were characterized by the same patterns. Letters A-F represent six different genotypes and error bars represent standard error. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58674/58674fig8large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Label N was higher in shoot than root samples across genotypes according to two-way ANOVA with sample type and genotype as fixed factors (n = 23, Figure 9), showing that 77-81% of N taken up from the treatment patch was translocated from roots to shoots during the experiment. One-way ANOVA (n = 23) showed that &#948;15N of root and shoot samples did not vary by genotype.
<img alt="Figure 9" class="xfigimg" src="/files/ftp_upload/58674/58674fig9.jpg" /> 
Figure 9: Nitrogen obtained from legume residue in roots and shoots at harvest. All genotypes were equally effective at taking up N from the patch containing 15N-labeled legume residue. The majority of N taken up from the patch was translocated from roots to shoots. Letters A-F represent six different genotypes and error bars represent standard error. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58674/58674fig9large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about An Optimized Rhizobox Protocol to Visualize Root Growth and Responsiveness to Localized Nutrients at JoVE.com

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

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 ...

an-optimized-rhizobox-protocol-to-visualize-root-growth

Nanyang Technological University

Research

JoVE Journal

Environment

15.8K Views.  University of California at Davis. Deze methode kan helpen bij het beantwoorden van belangrijke vragen op het gebied van Root en Rhizosphere Science, zoals hoe wortels reageren op gelokaliseerde plekken van voedingsstoffen. De belangrijkste voordelen van deze techniek zijn dat het mogelijk maakt herhaalde, niet-invasieve metingen van een enkel wortelstelsel in de bodem, maar het vereist geen gespecialiseerde apparatuur en het is relatief goedkoop. Verzamel de onderdelen voor de Rhizobox voor montage.Deze omvatten voor-e...

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

Single-plant, Sterile Microcosms for Nodulation and Growth of the Legume Plant Medicago truncatula with the Rhizobial Symbiont Sinorhizobium meliloti

Rhizobial bacteria form symbiotic, nitrogen-fixing nodules on the roots of compatible host legume plants. One of the most well-developed model systems for studying these interactions is the plant Medicago truncatula cv. Jemalong A17 and the rhizobial bacterium Sinorhizobium meliloti 1021. Repeated imaging of plant roots and scoring of symbiotic phenotypes requires methods that are non-destructive to either plants or bacteria. The symbiotic phenotypes of some plant and bacterial mutants become apparent after relatively short periods of growth, and do not require long-term observation of the host/symbiont interaction. However, subtle differences in symbiotic efficiency and nodule senescence phenotypes that are not apparent in the early stages of the nodulation process require relatively long growth periods before they can be scored. Several methods have been developed for long-term growth and observation of this host/symbiont pair. However, many of these methods require repeated watering, which increases the possibility of contamination by other microbes. Other methods require a relatively large space for growth of large numbers of plants. The method described here, symbiotic growth of M. truncatula/S. meliloti in sterile, single-plant microcosms, has several advantages. Plants in these microcosms have sufficient moisture and nutrients to ensure that watering is not required for up to 9 weeks, preventing cross-contamination during watering. This allows phenotypes to be quantified that might be missed in short-term growth systems, such as subtle delays in nodule development and early nodule senescence. Also, the roots and nodules in the microcosm are easily viewed through the plate lid, so up-rooting of the plants for observation is not required.

Single-plant, Sterile Microcosms for Nodulation and Growth of the Legume Plant Medicago truncatula with the Rhizobial Symbiont Sinorhizobium meliloti

Growth of Medicago truncatula plants in symbiosis with the nitrogen-fixing bacteria Sinorhizobium meliloti in individual, sterile microcosms made from standard laboratory plates permits frequent examination of root systems and nodules without compromising sterility. Plants can be maintained in these growth chambers for up to 9 weeks.

Rhizobial  bacteria form symbiotic, nitrogen-fixing nodules on the roots of compatible host  legume plants.  One  of the most well-developed model systems ...

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 ...

A Simple Chamber for Long-term Confocal Imaging of Root and Hypocotyl Development

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and subcellular processes, high resolution time-lapse microscopy strategies that preserve physiological conditions are required. Plant tissues must have adequate nutrient and water supply with sustained gaseous exchange but, when submerged and immobilized under a coverslip, they are particularly susceptible to anoxia. One strategy that has been successfully employed is the use of a perfusion system to maintain a constant supply of oxygen and nutrients. However, such arrangements can be complicated, cumbersome, and require specialized equipment. Presented here is an alternative strategy for a simple imaging system using perfluorodecalin as an immersion medium. This system is easy to set up, requires minimal equipment, and is easily mounted on a microscope stage, allowing several imaging chambers to be set up and imaged in parallel. In this system, lateral root growth rates are indistinguishable from growth rates under standard conditions on agar plates for the first two days, and lateral root growth continues at reduced rates for at least another day. Plant tissues are supplied with nutrients via an agar slab that can be used also to administer a range of pharmacological compounds. The system was established to monitor lateral root development but is readily adaptable to image other plant organs such as hypocotyls and primary roots.

Presented here is a simple technique for high-resolution confocal time-lapse imaging of root and hypocotyl development for up to 3 days using high numerical-aperture objectives and perfluorodecalin as an immersion medium.

Several aspects of plant development, such as lateral root morphogenesis, occur on time spans of several days. To study underlying cellular and ...

Developmental Biology

Two-Dimensional Visualization and Quantification of Labile, Inorganic Plant Nutrients and Contaminants in Soil

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic nutrient (e.g., P, Fe, Mn) and contaminant (e.g., As, Cd, Pb) solute species in the soil adjacent to plant roots (the &#8216;rhizosphere&#8217;) at sub-millimeter (~100 &#181;m) spatial resolution. The method combines sink-based solute sampling by the diffusive gradients in thin films (DGT) technique with spatially resolved chemical analysis by laser ablation inductively coupled plasma mass spectrometry (LA-ICP-MS). The DGT technique is based on thin hydrogels with homogeneously distributed analyte-selective binding phases. The variety of available binding phases allows for the preparation of different DGT gel types following simple gel fabrication procedures. For DGT gel deployment in the rhizosphere, plants are grown in flat, transparent growth containers (rhizotrons), which enable minimal invasive access to a soil-grown root system. After a pre-growth period, DGT gels are applied to selected regions of interest for in situ solute sampling in the rhizosphere. Afterwards, DGT gels are retrieved and prepared for subsequent chemical analysis of the bound solutes using LA-ICP-MS line-scan imaging. Application of internal normalization using 13C and external calibration using matrix-matched gel standards further allows for the quantification of the 2D solute fluxes. This method is unique in its capability to generate quantitative, sub-mm scale 2D images of multi-element solute fluxes in soil-plant environments, exceeding the achievable spatial resolution of other methods for measuring solute gradients in the rhizosphere substantially. We present the application and evaluation of the method for imaging multiple cationic and anionic solute species in the rhizosphere of terrestrial plants and highlight the possibility of combining this method with complementary solute imaging techniques.

This protocol presents a workflow for sub-mm 2D visualization of multiple labile inorganic nutrient and contaminant solute species using diffusive gradients in thin films (DGT) combined with mass spectrometry imaging. Solute sampling and high-resolution chemical analysis are described in detail for quantitative mapping of solutes in the rhizosphere of terrestrial plants.

We describe a method for two-dimensional (2D) visualization and quantification of the distribution of labile (i.e., reversibly adsorbed) inorganic ...

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 ...

Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System

Bacteria form complex rhizosphere microbiomes shaped by interacting microbes, larger organisms, and the abiotic environment. Under laboratory conditions, rhizosphere colonization by plant growth-promoting bacteria (PGPB) can increase the health or the development of host plants relative to uncolonized plants. However, in field settings, bacterial treatments with PGPB often do not provide substantial benefits to crops. One explanation is that this may be due to loss of the PGPB during interactions with endogenous soil microbes over the lifespan of the plant. This possibility has been difficult to confirm, since most studies focus on the initial colonization rather than maintenance of PGPB within rhizosphere communities. It is hypothesized here that the assembly, coexistence, and maintenance of bacterial communities are shaped by deterministic features of the rhizosphere microenvironment, and that these interactions may impact PGPB survival in native settings. To study these behaviors, a hydroponic plant-growth assay is optimized using Arabidopsis thaliana to quantify and visualize the spatial distribution of bacteria during initial colonization of plant roots and after transfer to different growth environments. This system&#8217;s reproducibility and utility are then validated with the well-studied PGPB Pseudomonas simiae. To investigate how the presence of multiple bacterial species may affect colonization and maintenance dynamics on the plant root, a model community from three bacterial strains (an Arthrobacter, Curtobacterium, and Microbacterium species) originally isolated from the A. thaliana rhizosphere is constructed. It is shown that the presence of these diverse bacterial species can be measured using this hydroponic plant-maintanence assay, which provides an alternative to sequencing-based bacterial community studies. Future studies using this system may improve the understanding of bacterial behavior in multispecies plant microbiomes over time and in changing environmental conditions.

Monitoring Bacterial Colonization and Maintenance on Arabidopsis thaliana Roots in a Floating Hydroponic System

Here we describe a hydroponic plant growth assay to quantify species presence and visualize the spatial distribution of bacteria during initial colonization of plant roots and after their transfer into different growth environments.

Bacteria form complex rhizosphere microbiomes shaped by interacting microbes, larger organisms, and the abiotic environment. Under laboratory conditions, ...

Immunology and Infection

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 ...

Biology

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 ...

Plant Promoter Analysis: Identification and Characterization of Root Nodule Specific Promoter in the Common Bean

The upstream sequences of gene coding sequences are termed as promoter sequences. Studying the expression patterns of promoters are very significant in understanding the gene regulation and spatiotemporal expression patterns of target genes. On the other hand, it is also critical to establish promoter evaluation tools and genetic transformation techniques that are fast, efficient, and reproducible. In this study, we investigated the spatiotemporal expression pattern of the rhizobial symbiosis-specific nodule inception (NIN) promoter of Phaseolus vulgaris in the transgenic hairy roots. Using plant genome databases and analysis tools we identified, isolated, and cloned the P. vulgaris NIN promoter in a transcriptional fusion to the chimeric reporter &#946;-glucuronidase (GUS) GUS-enhanced::GFP. Further, this protocol describes a rapid and versatile system of genetic transformation in the P. vulgaris using Agrobacterium rhizogenes induced hairy roots. This system generates &#8805;2 cm hairy roots at 10 to 12 days after transformation. Next, we assessed the spatiotemporal expression of NIN promoter in Rhizobium inoculated hairy roots at periodic intervals of post-inoculation. Our results depicted by GUS activity show that the NIN promoter was active during the process of nodulation. Together, the present protocol demonstrates how to identify, isolate, clone, and characterize a plant promoter in the common bean hairy roots. Moreover, this protocol is easy to use in non-specialized laboratories.

Promoter expression analyses are crucial to improving the understanding of gene regulation and the spatiotemporal expression of target genes. Herein we present a protocol to identify, isolate, and clone a plant promoter. Further, we describe the characterization of the nodule-specific promoter in the common bean hairy roots.

The upstream sequences of gene coding sequences are termed as promoter sequences. Studying the expression patterns of promoters are very significant in ...

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

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