The authors acknowledge a joint seed grant to Seyfferth and Tappero to support collaboration between the University of Delaware and Brookhaven National Laboratory. Parts of this research used the XFM (4-BM) Beamline of the National Synchrotron Light Source II, a U.S. Department of Energy (DOE) Office of Science User Facility operated for the DOE Office of Science by Brookhaven National Laboratory under Contract No. DE-SC0012704.

1. Preparation of slam-freezing equipment
<ol>
	<li>Place two copper blocks (~5 cm x 5 cm x 15 cm) horizontally inside of a clean cooler capable of holding liquid nitrogen and pour enough liquid nitrogen to submerge the blocks. Once the bubbling subsides, place two spacers on top of one copper block at each end. 
	NOTE: The spacer height determines the height of the sample to be frozen; this example uses a 2 cm spacer to create cubes approximately 3 cm x 3 cm x 2 cm. The volume of the liquid nitrogen will depend on the cooler size. This example uses approximately 1 L for approximately 5 cubes in series. 
	CAUTION: Use proper personal protective equipment and ventilation as liquid nitrogen is a cryogen and an asphyxiant.</li>
	<li>Using tongs and cryogenic gloves, stand up the other copper block on its end, to make retrieval easier when the sample is in place.</li>
</ol>
2. Sample collection and slam-freezing
<ol>
	<li>Extract the desired plant and rhizosphere from the wet soil using a shovel and ensure that the dug hole is much larger than the desired root volume. Place the soil and plant into a container and place it on a benchtop. 
	NOTE: The entire potted soil and plant from a pot study can also be used.</li>
	<li>Determine the desired soil location where roots are to be taken (i.e., depth and proximity to the shoot). Cut away excess soil using a steel blade, taking care not to disturb soil in the desired area. When the desired area is reached, cut a root &#34;cube&#34; approximately 3 cm x 3 cm x 2 cm and immediately place the cube between the two spacers on the horizontal copper block. Using cryogenic gloves, pick up the vertical copper block and place it on top of the spacers to slam-freeze the rhizosphere cube.</li>
	<li>After bubbling subsides (~5 min), retrieve the slam-frozen rhizosphere cube from the copper blocks and wrap inside of a pre-labeled aluminum foil square. Mark the orientation of the block on the foil if desired. Place in a second container of liquid nitrogen until storage in a -80 &#176;C freezer.</li>
	<li>Repeat as needed to obtain the desired number of root cubes from the field site or the experiment. Ensure both copper blocks are given time to cool between samples.</li>
</ol>
3. Freeze-drying and embedding rhizosphere cubes
<ol>
	<li>Prepare the freeze dryer according to the manufacturer&#39;s instructions. Take care to ensure it has obtained the proper vacuum pressure and temperature prior to removing samples from the -80 &#176;C freezer.</li>
	<li>When freeze dryer is ready to receive samples, place one frozen rhizosphere cube inside of a clean and acid-washed 50 mL tube and cover loosely with a clean disposable wipe. Secure the wipe with a rubber band. Repeat as needed to ensure one cube per tube. 
	NOTE: If the sample is too large for a tube, it can be placed directly into the freeze dryer vessel using the aluminum foil as a sample holder.</li>
	<li>Place tubes containing samples in freeze dryer vessels and freeze dry for several days. The exact drying time will depend on soil properties. 
	NOTE: Store dried samples in the freeze dryer or a desiccator to avoid rehydration.</li>
	<li>Use a steel blade to cut dried soil cubes to size so that they fit into the desired form (e.g., 25 mm diameter form is ideal for most applications). Label each form, place the soil cubes in the forms and place the forms inside a vacuum desiccator.</li>
	<li>Prepare epoxy according to manufacturer&#39;s instructions. Ensure that the chosen epoxy is not contaminated with and does not cause speciation changes of desired elements 20, 29, 30.</li>
	<li>Use a dropper to add epoxy to the form on one side of the soil, till it entirely covers the sample. The soil will darken in color as the epoxy wets the soil. 
	NOTE: Add the epoxy slowly to allow the air in the soil to escape.</li>
	<li>Once forms are filled with epoxy, close the vacuum desiccator and turn on the vacuum. Depending on the amount of air trapped in the soil, more epoxy may need to be added to the forms periodically. Check the level of epoxy every 30-90 min for the first 1-4 h and add epoxy as needed.</li>
	<li>Remove the sample from the form once the epoxy has hardened (~5 days).</li>
</ol>
4. Cutting and sectioning the rhizosphere cubes
<ol>
	<li>Cut the sample using a diamond blade precision wet saw. Cut the samples in different locations if no roots are obtained in the previous cut.</li>
	<li>Manually sand the cut samples with progressively finer sandpaper (e.g., 220, 500, 1000, and 1500 grit) on the cut side for ~30 s.</li>
	<li>Perform surface imaging of the samples using techniques such as LA-ICP-MS. 
	NOTE: To prepare thin sections for S-XRF, either send the samples out to a company capable of preparing the thin sections (single or double side polishing) or follow the steps 4.4 - 4.6 as described below.</li>
	<li>Glue the desired sample side to a quartz slide using super glue and allow to cure overnight.</li>
	<li>Using a thin sectioning machine, cut soil on slides to 2 mm thick and then grind to the desired thickness (typically 30 &#181;m). The sample surface can be polished if desired.</li>
	<li>Perform S-XRF imaging of the sections. Follow the appropriate steps at the desired synchrotron facility and beamline to apply for and utilize imaging time.</li>
</ol>

The authors have nothing to disclose.

This paper describes a protocol to obtain preserved bulk soil + rhizospheres of wetland plant roots using a slam-freezing technique that can be used for elemental imaging and/or chemical speciation mapping.
There are several benefits of this method over existing methods. First, this method allows the simultaneous investigation of roots and the surrounding rhizospheres. Methods currently exist to preserve and chemically image roots out of their soil environment by washing away the soil and pres...

Roots and their rhizospheres are dynamic, heterogeneous, and critically important for understanding how plants obtain mineral nutrients and contaminants1,2,3. Roots are the primary pathway by which nutrients (e.g., phosphorus) and contaminants (e.g., arsenic) move from soil to plants and thus understanding this process has implications for food quantity and quality, ecosystem functioning, and phytoremediation. However, roots are dynamic in space and time growing in response to nutrient acquisition needs and they often vary in function, diameter, and structure (e.g., lateral roots, adventitious roots, root hairs)2. Heterogeneity of root systems can be studied on spatial scales from cellular to ecosystem-level and on temporal scales from hourly to decadal. Thus, the dynamic and heterogeneous nature of roots and their surrounding soil, or rhizosphere, poses challenges for capturing rhizosphere chemistry over time. Despite this challenge, it is imperative to study roots in their soil environment to characterize this critical plant-soil relationship.
The rhizosphere chemistry of wetland plants is particularly challenging to investigate because of steep oxygen gradients that exist from bulk soil to the roots, which change in space and time. Because roots need oxygen to respire, wetland plants have adapted to the low oxygen conditions of wetland soils by creating aerenchyma4, 5. Aerenchyma are hollowed cortical tissues that extend from shoots to roots, allowing the diffusion of air through the plant into the roots. However, some of this air leaks into the rhizosphere in less suberized parts of the roots particularly near lateral root junctions, less mature root tips and elongation zones6,7,8,9. This radial oxygen loss creates an oxidized zone in the rhizosphere of wetland plants that affects rhizosphere (bio-geo)chemistry and is distinct from the reduced bulk soil10,11,12. To understand the fate and transport of nutrients and contaminants in wetland rhizospheres and roots, it is critical to preserve the chemically reduced bulk soil, the oxidized rhizosphere, and roots of wetland plants for analysis. However, because the bulk soil contains reduced soil constituents that are oxygen-sensitive, root and soil preservation methods must preserve root structures and minimize oxygen-sensitive reactions.
Methods exist to fix plant tissues and preserve the ultrastructure for imaging, but those methods cannot be applied to chemically preserve roots growing in wetland soil. For investigations where only the elemental distribution within plant cells is desired, plants are typically grown hydroponically and roots can be easily removed from solution, fixed under high-pressure freezing and freeze substitution and sectioned for a variety of imaging applications including high-resolution secondary ion mass spectrometry (nanoSIMS), electron microscopy, and synchrotron X-ray fluorescence (S-XRF) analysis13,14,15. To investigate Fe plaque on the outside of wetland roots, these hydroponic studies must artificially induce Fe plaque formation in solution16, which does not accurately represent the heterogeneity of the distribution and mineral composition of Fe plaque formation and associated elements in situ17,18,19,20. Methods exist to preserve wetland soil and associated microorganisms with freeze-coring21, but it is difficult to obtain roots with this technique. Current methods to visualize roots growing in soil and their rhizospheric chemistry consist of two primary measurement types: elemental fluxes and total elemental concentration (and speciation). The former is typically measured using diffusive gradients in thin films (DGT)22,23,24, in which soil is placed into rhizoboxes to support plant growth in a laboratory setting and labile elements in the soil diffuse through a gel into a binding layer. This binding layer can then be imaged to quantify the labile elements of interest. This technique can successfully illustrate relationships between roots and the rhizosphere24,25,26,27, but artefacts from root-bounding may exist by growing plants in rhizoboxes, and information on the root interior is not captured with DGT. The latter involves sampling of the roots and rhizosphere, preserving the sample, and directly analyzing elemental distribution on a sample section. For this environmental sampling of wetland plant roots and their surrounding rhizosphere, careful sample handling is required to avoid artefacts from sample preparation.
Here a protocol is described that effectively preserves root structures and rhizosphere chemistry of wetland plants by slam-freezing and freeze drying. Flash-freezing can drastically slow down transformations of oxygen sensitive solutes but may damage roots and may cause mobilization when samples dry out. However, slam-freezing where the sample is frozen between copper blocks pre-cooled with liquid nitrogen minimizes root damage and sample distortion28. The preserved samples are then embedded in an epoxy resin that preserves As speciation20, 29 and can be cut and polished for imaging of roots within their rhizosphere soil. The samples in this report were analyzed by S-XRF chemical speciation imaging after thin sectioning. However, other imaging techniques could also be used, including laser ablation-inductively coupled plasma mass spectrometry (LA-ICP-MS), particle induced x-ray emission (PIXE), secondary ion mass spectrometry (SIMS), and laser induced breakdown spectroscopy (LIBS) imaging.

<table><tbody><tr><td>Copper blocks</td><td>McMaster Carr</td><td>89275K42</td><td></td></tr><tr><td>Diamond blade</td><td>Buehler</td><td>15 LC, 102 mm x 0.3 mm</td><td>operation speed: 225 rpm</td></tr><tr><td>Epoxy forms</td><td>Struers</td><td>40300085</td><td>FixiForm</td></tr><tr><td>Epoxy</td><td>Epotek</td><td>301-2FL</td><td></td></tr><tr><td>Superglue</td><td>Loctite</td><td>404</td><td></td></tr><tr><td>Thin sectioning machine</td><td>Buehler</td><td>PetroThin</td><td></td></tr><tr><td>Wet saw</td><td>Buehler</td><td>IsoMet 1000</td><td></td></tr></tbody></table>

a method to preserve wetland roots and rhizospheres for elemental imaging

Roots extensively interact with their soil environment but visualizing such interactions between roots and the surrounding rhizosphere is challenging. The rhizosphere chemistry of wetland plants is particularly challenging to capture because of steep oxygen gradients from the roots to the bulk soil. Here a protocol is described that effectively preserves root structure and rhizosphere chemistry of wetland plants through slam-freezing and freeze drying. Slam-freezing, where the sample is frozen between copper blocks pre-cooled with liquid nitrogen, minimizes root damage and sample distortion that can occur with flash-freezing while still minimizing chemical speciation changes. While sample distortion is still possible, the ability to obtain multiple samples quickly and with minimal cost increases the potential to obtain satisfactory samples and optimizes imaging time. The data show that this method is successful in preserving reduced arsenic species in rice roots and rhizospheres associated with iron plaques. This method can be adopted for studies of plant-soil relationships in a wide variety of wetland environments that span concentration ranges from trace-element cycling to phytoremediation applications.

This method allows for preservation of roots and chemical species in the roots and rhizosphere of wetland plants and into the bulk soil. In this work, the method was used to evaluate As speciation and co-localization with Fe and Mn oxides and plant nutrients in the rhizosphere of rice (Oryza sativa L.). Rice was grown at the RICE Facility at the University of Delaware where 30 rice paddy mesocosms (2 m x 2 m, 49 plants each) are used to grow rice under various soil and water management conditions with the goal o...

Watch this Scientific Journal Video about A Method to Preserve Wetland Roots and Rhizospheres for Elemental Imaging at JoVE.com

A Method to Preserve Wetland Roots and Rhizospheres for Elemental Imaging

We describe a protocol to sample, preserve, and section intact roots and the surrounding rhizosphere soil from wetland environments using rice (Oryza sativa L.) as a model species. Once preserved, the sample can be analyzed using elemental imaging techniques, such as synchrotron X-ray fluorescence (XRF) chemical speciation imaging.

Roots extensively interact with their soil environment but visualizing such interactions between roots and the surrounding rhizosphere is challenging. The ...

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Universidad de Cartagena UDEC

Research

JoVE Journal

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3.3K Views.  University of Delaware. We describe a protocol to sample, preserve, and section intact roots and the surrounding rhizosphere soil from wetland environments using rice (Oryza sativa L.) as a model species. Once preserved, the sample can be analyzed using elemental imaging techniques, such as synchrotron X-ray fluorescence (XRF) chemical speciation imaging. 

Video: A Method to Preserve Wetland Roots and Rhizospheres for Elemental Imaging

Exploring the Root Microbiome: Extracting Bacterial Community Data from the Soil, Rhizosphere, and Root Endosphere

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to abiotic stresses and diseases. As the plant microbiome can be highly complex, low-cost, high-throughput methods such as amplicon-based sequencing of the 16S rRNA gene are often preferred for characterizing its microbial composition and diversity. However, the selection of appropriate methodology when conducting such experiments is critical for reducing biases that can make analysis and comparisons between samples and studies difficult. This protocol describes in detail a standardized methodology for the collection and extraction of DNA from soil, rhizosphere, and root samples. Additionally, we highlight a well-established 16S rRNA amplicon sequencing pipeline that allows for the exploration of the composition of bacterial communities in these samples, and can easily be adapted for other marker genes. This pipeline has been validated for a variety of plant species, including sorghum, maize, wheat, strawberry, and agave, and can help overcome issues associated with the contamination from plant organelles.

Here, we describe a protocol to obtain amplicon sequence data of soil, rhizosphere, and root endosphere microbiomes. This information can be used to investigate the composition and diversity of plant-associated microbial communities, and is suitable for the use with a wide range of plant species.

The intimate interaction between plant host and associated microorganisms is crucial in determining plant fitness, and can foster improved tolerance to ...

Genetics

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 ...

Ecosystem Fabrication (EcoFAB) Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A better mechanistic understanding of these complex plant-microbe interactions will be crucial to improving plant production as well as performing basic ecological studies investigating plant-soil-microbe interactions. Here, a detailed description for ecosystem fabrication is presented, using widely available 3D printing technologies, to create controlled laboratory habitats (EcoFABs) for mechanistic studies of plant-microbe interactions within specific environmental conditions. Two sizes of EcoFABs are described that are suited for the investigation of microbial interactions with various plant species, including Arabidopsis thaliana, Brachypodium distachyon, and Panicum virgatum. These flow-through devices allow for controlled manipulation and sampling of root microbiomes, root chemistry as well as imaging of root morphology and microbial localization. This protocol includes the details for maintaining sterile conditions inside EcoFABs and mounting independent LED light systems onto EcoFABs. Detailed methods for addition of different forms of media, including soils, sand, and liquid growth media coupled to the characterization of these systems using imaging and metabolomics are described. Together, these systems enable dynamic and detailed investigation of plant and plant-microbial consortia including the manipulation of microbiome composition (including mutants), the monitoring of plant growth, root morphology, exudate composition, and microbial localization under controlled environmental conditions. We anticipate that these detailed protocols will serve as an important starting point for other researchers, ideally helping create standardized experimental systems for investigating plant-microbe interactions.

Ecosystem Fabrication EcoFAB Protocols for The Construction of Laboratory Ecosystems Designed to Study Plant-microbe Interactions

This article describes detailed protocols for ecosystem fabrication of devices (EcoFABs) that enable the studies of plants and plant-microbe interactions in highly controlled laboratory conditions.

Beneficial plant-microbe interactions offer a sustainable biological solution with the potential to boost low-input food and bioenergy production. A ...

Isolation and Analysis of Microbial Communities in Soil, Rhizosphere, and Roots in Perennial Grass Experiments

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The purpose of this manuscript is to provide detail on how to rapidly sample soil, rhizosphere, and endosphere of replicated field trials and analyze changes that may occur in the microbial communities due to sample type, treatment, and plant genotype. The experiment used to demonstrate these methods consists of replicated field plots containing two, pure, warm-season grasses (Panicum virgatum and Andropogon gerardii) and a low-diversity grass mixture (A. gerardii, Sorghastrum nutans, and Bouteloua curtipendula). Briefly, plants are excavated, a variety of roots are cut and placed in phosphate buffer, and then shaken to collect the rhizosphere. Roots are brought to the laboratory on ice and surface sterilized with bleach and ethanol (EtOH). The rhizosphere is filtered and concentrated by centrifugation. Excavated soil from around the root ball is placed into plastic bags and brought to the lab where a small amount of soil is taken for DNA extractions. DNA is extracted from roots, soil, and rhizosphere and then amplified with primers for the V4 region of the 16S rRNA gene. Amplicons are sequenced, then analyzed with open access bioinformatics tools. These methods allow researchers to test how the microbial community diversity and composition varies due to sample type, treatment, and plant genotype. Using these methods along with statistical models, the representative results demonstrate there are significant differences in microbial communities of roots, rhizosphere, and soil. Methods presented here provide a complete set of steps for how to collect field samples, isolate, extract, quantify, amplify, and sequence DNA, and analyze microbial community diversity and composition in replicated field trials.

Excavation of plant roots from the field as well as processing of samples into endosphere, rhizosphere, and soil are described in detail, including DNA extraction and data analysis methods. This paper is designed to enable other laboratories to use these techniques for the study of soil, endosphere, and rhizosphere microbiomes.

Plant and soil microbiome studies are becoming increasingly important for understanding the roles microorganisms play in agricultural productivity. The ...

Measuring Stolons and Rhizomes of Turfgrasses Using a Digital Image Analysis System

Length and diameter of stolons or rhizomes are usually measured using simple rulers and calipers. This procedure is slow and laborious, so it is often used on a limited number of stolons or rhizomes. For this reason, these traits are limited in their use for morphological characterization of plants. The use of digital image analysis software technology may overcome measurement errors due to human mistakes, which tend to increase as the number and size of samples also increase. The protocol can be used for any kind of crop but is particularly suitable for forage or grasses, where plants are small and numerous. Turf samples consist of aboveground biomass and an upper soil layer to the depth of maximum rhizome development, depending on the species of interest. In studies, samples are washed from the soil, and stolons/rhizomes are cleaned by hand before analysis by digital image analysis software. The samples are further dried in a laboratory heating oven to measure dry weight; therefore, for each sample, the resultant data are total length, total dry weight, and average diameter. Scanned images can be corrected before analysis by excluding visible extraneous parts, such as remaining roots or leaves not removed with the cleaning process. Indeed, these fragments normally have much smaller diameters than stolons or rhizomes, so they can be easily excluded from analysis by fixing the minimum diameter below which objects are not considered. Stolon or rhizome density per unit area can then be calculated based on sample size. The advantage of this method is quick and efficient measurement of the length and average diameter of large sample numbers of stolons or rhizomes.

A software-based image analysis system provides an alternative method to study the morphology of stoloniferous and rhizomatous species. This protocol permits measurement of the length and diameter of stolons and rhizomes and can be applied to samples with a large amount of biomass and to a wide variety of species.

Length and diameter of stolons or rhizomes are usually measured using simple rulers and calipers. This procedure is slow and laborious, so it is often ...

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

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 ...

Investigation of Plant Interactions Across Common Mycorrhizal Networks Using Rotated Cores

Arbuscular mycorrhizal (AM) fungi influence plant mineral nutrient uptake and growth, hence, they have the potential to influence plant interactions. The power of their influence is in extraradical mycelia that spread beyond nutrient depletion zones found near roots to ultimately interconnect individuals within a common mycorrhizal network (CMN). Most experiments, however, have investigated the role of AM fungi in plant interactions by growing plants with versus without mycorrhizal fungi, a method that fails to explicitly address the role of CMNs. Here, we propose a method that manipulates CMNs to investigate their role in plant interactions. Our method uses modified containers with conical bottoms with a nylon mesh and/or hydrophobic material covering slotted openings, 15N fertilizer, and a nutrient-poor interstitial sand. CMNs are left either intact between interacting individuals, severed by rotation of containers, or prevented from forming by a solid barrier. Our findings suggest that rotating containers is sufficient to disrupt CMNs and prevent their effects on plant interactions across CMNs. Our approach is advantageous because it mimics aspects of nature, such as seedlings tapping into already established CMNs and the use of a suite of AM fungi that may provide diverse benefits. Although our experiment is limited to investigating plants at the seedling stage, plant interactions across CMNs can be detected using our approach which therefore can be applied to investigate biological questions about the functioning of CMNs in ecosystems.

Most plants within communities likely are interconnected by arbuscular mycorrhizal (AM) fungi, but mediation of plant interactions by them has been investigated primarily by growing plants with versus without mycorrhizas. We present a method to manipulate common mycorrhizal networks among mycorrhizal plants to investigate their consequences for plant interactions.

Arbuscular mycorrhizal (AM) fungi influence plant mineral nutrient uptake and growth, hence, they have the potential to influence plant interactions. The ...

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 ...

Light Sheet Fluorescence Microscopy of Plant Roots Growing on the Surface of a Gel

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the observation of the whole organ with a high spatial- as well as temporal resolution over prolonged periods of time, which may cause photo-toxic effects. This protocol shows a plant sample preparation method for light-sheet microscopy, which is characterized by mounting the plant vertically on the surface of a gel. The plant is mounted in such a way that the roots are submerged in a liquid medium while the leaves remain in the air. In order to ensure photosynthetic activity of the plant, a custom-made lighting system illuminates the leaves. To keep the roots in darkness the water surface is covered with sheets of black plastic foil. This method allows long-term imaging of plant organ development in standardized conditions.

This protocol shows a plant sample preparation method for light-sheet microscopy. The setup is characterized by mounting the plant vertically on the surface of a gel and letting it grow in controlled bright conditions. This allows long-term observation of plant organ development in standardized conditions.

One of the key questions in understanding plant development is how single cells behave in a larger context of the tissue. Therefore, it requires the ...