This paper uses data resulting from two Ph.D. studies in the thematic area of &#34;Corn Mycorrhizal Patterns Driven by Agronomic Inputs&#34;, conducted by Victoria Pop-Moldovan, and &#34;Mycorrhizal Status and Development of Colonization in Mountain Grassland Dominant Species&#34;, conducted by Larisa Corcoz, under the coordination of Prof. Dr. Roxana Vidican.

1. Selection of biological material, root sampling, and storage
<ol>
	<li>Collect the entire root of plants with a shovel (Figure 1A) separately for each variant and replicate. Remove gently, by hand, the large soil aggregates from the roots. Wash the entire root system and measure it on a scale with 1 cm x 1 cm cells (Figure 1B). Cut the roots separately for each plant, and place them into a plastic bag.</li>
	<li>Collect all the clean roots from each plant in a plastic bag, and collect all samples from one variant in one bigger bag. Write on each bag the stage/variant name and the sampling date. Store the roots in a fridge or a freezer at a temperature between &#8722;4 &#176;C and &#8722;20 &#176;C until processing.</li>
</ol>
2. Root processing, clearing, and staining for microscopy
NOTE: Use gloves, a mask, and a microbiological/chemical hood for this step of the protocol.
<ol>
	<li>Ensure that the root thawing process is done slowly at room temperature. For all the steps of processing, use small jars (30-50 mL) to reduce the amount of necessary agents.</li>
	<li>Perform the following four steps of the slow clearing and staining procedure15. Do all the steps at room temperature. This method allows the processing of a large number of samples at the same time without the use of a water bath for boiling.
	<ol>
		<li>Root clearing: Place all roots from one plant in a jar. Prepare a 10% NaOH solution with tap water and pour it into each jar until it completely covers the roots. Shake the jars vigorously for 1 min or 2 min to produce a homogenous dispersion of clearing solution in the roots. Repeat this procedure after 24 h and leave the roots in the clearing solution for at least 48 h. 
		NOTE: Clean roots have a pale yellow (up to white) aspect, and the consistency is soft (they can be easily crushed by pressing with a tweezer).</li>
		<li>Root rinsing: Pass the content of one jar at a time through a sieve. Recycle the clearing solution. Rinse the roots several times in tap water until the clearing solution is completely removed. 
		NOTE: If the clearing solution is not completely removed, it will affect the quality of the staining procedure.</li>
		<li>Root staining: Place the rinsed roots in a clean jar. Prepare a 5%:5% ink-vinegar solution with tap water (5 mL of blue ink + 5 mL of 9% acetic acid + 90 mL of tap water). Pour the solution into each jar until it completely covers the roots. Shake the jars vigorously for 1 min or 2 min to produce a homogenous dispersion of staining solution in the roots. Repeat this procedure after 24 h and leave the roots in this solution for 48 h. 
		NOTE: Stained roots have an intense blue color.</li>
		<li>Root partial destaining: Rinse the stained roots in tap water for 1-2 min. Shake the jars vigorously to remove the extra staining solution. Repeat the procedure if the staining is too intense and does not permit a clear microscopic evaluation. 
		NOTE: Stained roots can be kept in tap water for up to 1 week at room temperature without altering the staining quality (Figure 2). For longer periods, roots can be maintained for up to 2-3 months in a 5% commercial apple vinegar solution (5% acetic acid).</li>
	</ol>
	</li>
</ol>
3. Root processing for microscopy
<ol>
	<li>Root segmentation: Place the stained roots from each sample on a scaled cutting board (Figure 3A). Cut the roots into 1 cm segments (Figure 3B). Choose 15 segments for each variant.</li>
	<li>Gentle crushing method for segment preparation: Spread the roots on a slide. Use a laminating pouch for covering the roots and gently crush them starting from an edge (Figure 3C,D). Use a soft plastic tool, e.g., tweezers, scalpel handle, pen, or a pencil with an eraser, to slowly display the roots on the slide. Carefully remove the laminating pouch and cover the sample with a coverslip (Figure 3E). 
	NOTE: Roots have a tubular form, so it is necessary to separate them in a bi-dimensional plane. This action presumes the separation of roots on the middle point, leading to the display of the two parts of the internal diameter. The use of laminating pouches in the gentle crushing procedure permits the display of the roots, which have a cylinder form, in two pieces-one on the left and one on the right-toward their middle point. In this way, the entire root is analyzed in-depth, and colonization degree is the parameter that shows the volumetric colonization (described in the original work on MycoPatt11). Basically, we cut a cylinder in half, and after that, we rebuild it mathematically.</li>
	<li>Add water to a corner of the slide with a pipette and let the water slowly spread on the slide (Figure 3F). Remove the extra water with a paper towel.</li>
</ol>
4. Microscopic analysis of the root samples
<ol>
	<li>Use a microscope equipped with a good resolution camera.</li>
	<li>Analyze the slides starting from an extremity. Capture each microscopic field. Rename each image captured with a code that will permit the real post-assemblage of root parts. For thick roots, use the 10x or 40x magnification, and for thin roots, use the 40x magnification. Use the same objective and magnification for the entire set of roots from a species.</li>
</ol>
5. Post-microscopy image assemblage
<ol>
	<li>Use presentation software to design a drawing board for image assemblage. Set the width 2-3 cm wider than the image width. Add all captured images from one segment in the order of their capture and reconstruct the entire length of the root segment (Figure 4A).
	<ol>
		<li>In brief, collect a total of 15 pictures for each 1 cm segment and organize them vertically, starting with 1 to 15, in the presentation software to reconstruct the segment.</li>
	</ol>
	</li>
	<li>Align the images in the center. Use the vertical alignment to ensure that each image is following the previous one. On all pictures, place a grid of 10 cells x 150 cells to cover the entire root segment.
	<ol>
		<li>Additionally, on each individual picture, place a 10 x 10 grid, and in every cell of this grid, insert a number from one to six if an AM structure is visible or leave blank if no AM structure is present. In this way, the accuracy of the process is maximal with no errors in the location of the AM structures being observed.</li>
	</ol>
	</li>
	<li>Add a table for a grid of 10 cells in width and 150 cells in length (15 squares of 10 cells x 10 cells). Change the table width dimensions to the width of the images. Change the length of the table to comprise all the images (Figure 4B).</li>
</ol>
6. Scoring of mycorrhizal colonization
<ol>
	<li>Use the unique number to score each type of structure as described in the mycorrhizal patterns method11: 1 for hyphae; 2 for arbuscules; 3 for vesicles; 4 for spores; 5 for auxiliary cells; and 6 for entry points (Figure 4C). Score each observed mycorrhizal structure from each cell of the previously applied grids (Figure 4D).</li>
</ol>
7. Raw data analysis and result extraction
<ol>
	<li>Insert all the obtained scores in the MycoPatt spreadsheet11. Use the copy/paste function to transfer all the scores from the presentation into the first sheet named as rawdata (Figure 5).</li>
	<li>Primary analysis of results: Use the third sheet named parameters in the MycoPatt spreadsheet tool to visualize the results as percentages (%) separately in three forms (Figure 6A-C). Use columns A to K to analyze the horizontal image of colonization; columns M to W to analyze the vertical image of colonization; and columns Y to AI to analyze the transversal (average) colonization for each of the 15 10 x 10 squares (lines 2-17) and the final average colonization (lines 19-20). 
	NOTE: The transversal average colonization is used for the calculation of the real colonization parameters, related to both horizontal and vertical analysis. In this way, no errors can be made as compared to if only the horizontal or vertical analysis is used (described in detail in the original work11). Also, this set of parameters is calculated for the entire surface of the microscopic field.
	<ol>
		<li>Use the definitions and formulas, specific for each parameter, to analyze the results11. Use the following colonization parameters: frequency of colonization (%), intensity of colonization (%), arbuscules (%) and vesicles (%), spores (%) and auxiliary cells (%), entry points (%), the percentage of non-mycorrhizal areas (%), overall colonization degree (%), and the report of mycorrhizal/non-mycorrhizal areas. 
		NOTE: If arbuscules, vesicles, spores, auxiliary cells, and entry points are missing from analyzed samples, the MycoPatt spreadsheet will score them as zero (0).</li>
	</ol>
	</li>
	<li>Production and extraction of mycorrhizal maps: Visualize the image obtained from the conversion of mycorrhizal structures code into colors in the second sheet of the MycoPatt named graphs (Figure 7A). Export the resultant image in the graphs sheet as an image (Figure 7B). Use the color code in the legend for the analysis of mycorrhizal patterns.</li>
	<li>Mycorrhizal map analysis: Identify the most important structures and their assemblage on mycorrhizal maps. Describe the mycorrhizal colonization pattern observed in the analyzed roots. Describe the mycorrhizal colonization strategy based on observed structural development in the root, the branching pattern, and arbuscule/vesicle development.</li>
</ol>

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The authors declare no conflicts of interest.

Studies on mycorrhizal colonization are vital for new strategy development in the agronomic field. The potential of multiple cultivated plants to form a symbiotic association with arbuscular mycorrhizas made them an important component of the agroecosystem&#39;s sustainable development and the maintenance of its health16,17,18,19,20. Thus, there is a need for ...

Arbuscular mycorrhiza (AM) fungi are a category of soil-borne endophytes that are constantly an area of interest for researchers. Their presence in the roots of most plants and their involvement in nutrient cycles makes them vital components in the stability of every ecosystem where herbaceous plants are present1,2. Through their extra-radicular mycelium, AM act as a fungal extension for plant roots, especially in hard-to-reach areas3. The main activity is in host plant roots, where AM develops large hyphae networks and specific intracellular structures called arbuscules. The lack of host specificity allows the symbiont to colonize multiple species at the same time. This ability provides AM with the role of resource allocation and nutrient regulation in the ecosystem; the fungus also provides support in plant survival and aids in plant performance4,5,6,7. The reaction of AM species to host roots is visible in the extension and location of the intra-radicular mycelium and the presence and shape of the arbuscules developed intracellularly. The intracellular arbuscules act as an interchange point between the two symbionts and represent areas characterized by fast transfer processes. The structures that the AM produce are species-dependent, and, in addition to arbuscules, in the roots, they also develop vesicles, spores, and auxiliary cells.
There are many challenges in the assessment of AM symbionts in plant roots8,9. The first one is their constant development during the entire vegetation period of hosts, which leads to multiple changes in the hyphal arbuscular structure. The different stages of arbuscular growth, up to their collapse, are clearly present in roots, but the senescent AM structures are sometimes digested, which makes them only partially visible10. The second challenge is represented by the staining method and protocol, the large diversity of root systems, the dimension of their cells, and the differences in thickness, which make it hard to propose a unified method. The last challenge is represented by the assessment and scoring of AM colonization. There are numerous methods that score AM with different degrees of objectivity, and most of them are still restricted to microscopy techniques. The simple ones are based on the presence/absence of structures in the root cortex, while the more complex ones are based on visual scoring and the use of colonization classes, with the integration of the frequency and intensity of the colonization phenomenon. A lot of data have been produced in the last decades on the mycorrhizal status of multiple species, but most of the methods are restricted to the observed value of colonization without pointing to the real position of each structure in the root cortex. As a response to the necessity of more accurate results on AM colonization, a method based on microscopic analysis of mycorrhizal patterns (MycoPatt) in roots was developed to assemble, in a digital form, the detailed mycorrhizal maps11. Also, the method allows the objective calculation of colonization parameters and the determination of the actual position of each structure in the root.
The position of the AM fungal structures can be important in answering the following two questions. The first one is related to the analysis of the colonization in one specific moment from the vegetation cycle of a plant. In this context, it is very useful to observe the arbuscular/vesicle abundance, report how are they located in the root, and provide a very clear colonization image and parameters. The second one is related to the detection of fungal strategy and its orientation and even the forecast of its future development. One application of the MycoPatt can be for plants analyzed daily, every 2-3 days, weekly, or during various growth stages. In this context, the location of the vesicles/arbuscules is important to better understand the biological mechanism of AM colonization. These parameters and observations are very useful to supplement the mathematical parameters.
The aim of this article is to demonstrate the ability of the MycoPatt system to explore the native AM fungi colonization potential and strategy in Zea mays (corn) roots during different development stages and in Festuca rubra (red fescue) roots under different long-term fertilization conditions. To fulfill the aim, two large databases from two experiments were analyzed. The corn experiment was established at Cojocna (46&#176;44&#8242;56&#8243; lat. N and 23&#176;50&#8242;0&#8243; long. E), in the Experimental Didactic Farm of the University of Agricultural Sciences and Veterinary Medicine Cluj on a phaeoziom with a loamy texture soil12. The red fescue experiment is a part of a larger experimental site established in 2001 in Ghe&#539;ari, Apuseni Mountains (46&#176;49&#39;064&#34; lat. N and 22&#176;81&#39;418&#39;&#39; long. E), on a preluvosol (terra rossa) soil type13,14. Corn was collected in five different growth phenophases12: B1 = 2-4 leaves (as a control point for the start of mycorrhizal colonization); B2 = 6 leaves; B3 = 8-10 leaves; B4 = cob formation; B5 = physiological maturity. Starting from the 2-4 leaves stage (A0), an organic treatment was applied, which resulted in a two-graduation factor (A1 = control and A2 = treated). Roots of red fescue were collected at flowering from an experiment with five long-term fertilizations13,14: V1 = control, non-fertilized; V2 = 10 t&#183;ha-1 manure; V3 = 10 t&#183;ha-1 manure + N&#160;50 kg&#183;ha-1, P2O5 25 kg&#183;ha-1, K2O 25 kg&#183;ha-1; V4 = N 100 kg&#183;ha-1, P2O5 50 kg&#183;ha-1, K2O 50 kg&#183;ha-1; V5 = 10 t&#183;ha-1 manure + N 100 kg&#183;ha-1, P2O5 50 kg&#183;ha-1, K2O 50 kg&#183;ha-1. Five plants were collected in each development stage from every fertilization variant. The staining protocols and their performance in terms of sample processing time and quality of staining were analyzed. The relation between AM hyphae development and the presence of its structures in roots was analyzed separately for each species and continued with the identification of the most permissive roots for colonization. The specific colonization patterns of each root system were analyzed based on colonization maps and the value of AM parameters.
Corn is an annual plant, which implies continuous growth of the roots, and that was the main reason to apply the MycoPatt in the growing stages. Red fescue is a perennial plant from a grassland treated for a long time with different fertilizers. Its roots have a shorter development of 1 year, and the anthesis is considered as the vegetation point when the plant changes its metabolism from vegetative to generative. To catch these plants during these intense activity periods, the abovementioned time points were chosen. Sampling during the vegetation period is difficult for this species when grown in natural grasslands.

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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.

The correct use of the gentle crushing method of the roots after the staining procedures provides good details of mycorrhizal structures, both for Zea mays (Figure 8A-C) and Festuca rubra (Figure 9A-E), good contrast between mycorrhizal structures and root cells, and a confirmation of the stele due to the blue color. If the clearing and staining procedures fail to succeed, root...

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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.

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 ...

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Research

JoVE Journal

Biology

2.7K Views.  University of Agricultural Sciences and Veterinary Medicine Cluj-Napoca. MycoPatt-methode maakt een diepgaande verkenning van arbusculaire mycorrhiza-kolonisatie in wortels mogelijk. Mycorrhiza-kaarten wijzen op schimmeluitdijing en presenteren de werkelijke positie van elke structuur als een specifiek patroon. Mycorrhiza-kolonisatie wordt objectief geanalyseerd met een resolutie van 1%.Elk rootsegment wordt automatisch omgezet naar een digitale matrix en genereert een uitgebreide database met kolonisatieparameters. Verwijder om te beginnen de wortels uit d...

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

Quantification of Fungal Colonization, Sporogenesis, and Production of Mycotoxins Using Kernel Bioassays

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks to human and animal health. Among cereal production, maize is arguably the most affected crop, due to pathogen-induced losses in grain integrity and mycotoxin seed contamination. The two most prevalent and problematic mycotoxins for maize growers and food and feed processors are aflatoxin and fumonisin, produced by Aspergillus flavus and Fusarium verticillioides, respectively.
Recent studies in molecular plant-pathogen interactions have demonstrated promise in understanding specific mechanisms associated with plant responses to fungal infection and mycotoxin contamination1,2,3,4,5,6. Because many labs are using kernel assays to study plant-pathogen interactions, there is a need for a standardized method for quantifying different biological parameters, so results from different laboratories can be cross-interpreted. For a robust and reproducible means for quantitative analyses on seeds, we have developed in-lab kernel assays and subsequent methods to quantify fungal growth, biomass, and mycotoxin contamination. Four sterilized maize kernels are inoculated in glass vials with a fungal suspension (106) and incubated for a predetermined period. Sample vials are then selected for enumeration of conidia by hemocytometer, ergosterol-based biomass analysis by high performance liquid chromatography (HPLC), aflatoxin quantification using an AflaTest fluorometer method, and fumonisin quantification by HPLC.

The devastation of cereal crops by seed-infecting fungi has prompted numerous research efforts to better understand plant-pathogen interactions. To study seed-fungal interactions in a laboratory setting, we developed a robust method for the quantification of fungal reproduction, biomass, and mycotoxin contamination using kernel bioassays.

The rotting of grains by seed-infecting fungi poses one of the greatest economic challenges to cereal production worldwide, not to mention serious risks ...

Immunology and Infection

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 ...

Environment

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 ...

Culturing Arbuscular Mycorrhizal Fungi Using SAP-AS for Enhanced Observation

Inoculating and Observing Arbuscular Mycorrhizal Cultures on Superabsorbent Polymer-Based Autotrophic Systems

Arbuscular mycorrhizal (AM) fungi are difficult to manipulate and observe due to their permanent association with plant roots and propagation in the rhizosphere. Typically, AM fungi are cultured under in vivo conditions in pot culture with an autotrophic host or under in vitro conditions with Ri Transfer-DNA transformed roots (heterotrophic host) in a Petri dish. Additionally, the cultivation of AM fungi in pot culture occurs in an opaque and non-sterile environment. In contrast, in vitro culture involves the propagation of AM fungi in a sterile, transparent environment. The superabsorbent polymer-based autotrophic system (SAP-AS) has recently been developed and shown to combine the advantages of both methods while avoiding their respective limitations (opacity and heterotrophic host, sterility). Here, we present a detailed protocol for easy preparation, single spore inoculation, and observation of AM fungi in SAP-AS. By modifying the Petri dishes, high-resolution photographic and video observations were possible on living specimens, which would have been difficult or impossible with current in vivo and in vitro techniques.

Author Spotlight: Enhancing AM Fungi Research with SAP-AS &#8212; A Novel Technique for Single-Spore Cultures

Here, we describe a simple and inexpensive technique for inoculating and observing arbuscular mycorrhizal fungi in superabsorbent polymer-based autotrophic systems.

Arbuscular mycorrhizal (AM) fungi are difficult to manipulate and observe due to their permanent association with plant roots and propagation in the ...

Microfluidic Tools for Probing Fungal-Microbial Interactions at the Cellular Level

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic matter, as well as regulation of nutrient levels. There they also find numerous opportunities to interact with a variety of other microbes such as bacteria or other fungi. Studying fungal interactions at the cellular level, however, can be challenging owing to the black box-like nature of soil. New microfluidic tools are being developed for the study of fungal interactions; two platforms designed to study bacterial-fungal and fungal-fungal interactions are highlighted. Within these microchannels, fungal-microbial interactions can be monitored in controlled physico-chemical environments at higher temporal and spatial resolution than previously possible. Application of these tools have yielded numerous novel biological insights, such as the observation of bacterial polar attachment to hyphae or revealing uncharacterised fungal-fungal antagonisms. A key feature of these methodologies regards the ease of use of this tool by non-experts, yielding highly translatable technologies for use in microbiology labs.

Owing to the opacity of soil, interactions between its constituent microbes cannot easily be visualised with cellular resolution. Here, two microfluidic tools are presented, which offer new opportunities for investigating fungal-microbial interactions. The devices are versatile and simple to use, enabling high spatiotemporal control and high-resolution imaging at the cellular level.

Filamentous fungi are successful inhabitants of soil and play a major role in soil ecosystems, such as in the decomposition of organic and inorganic ...

Bioengineering

Isolation of Culturable Yeasts and Molds from Soils to Investigate Fungal Population Structure

Soil is host to an incredible amount of microbial life, with each gram containing up to billions of bacterial, archaeal, and fungal cells. Multicellular fungi such as molds and unicellular fungi, broadly defined as yeasts, fulfill essential roles in soil ecosystems as decomposers of organic material and as food sources for other soil dwellers. Fungal species diversity in soil is dependent on a multitude of climatic factors such as rainfall and temperature, as well as soil properties including organic matter, pH, and moisture. Lack of adequate environmental sampling, especially in regions of Asia, Africa, South America, and Central America, hinders the characterization of soil fungal communities and the discovery of novel species. 
We characterized soil fungal communities in nine countries across six continents using ~4,000 soil samples and a protocol developed in the laboratory for the isolation of yeasts and molds. This protocol begins with separate selective enrichment for yeasts and the medically relevant mold Aspergillus fumigatus, in liquid media while inhibiting bacterial growth. Resulting colonies are then transferred to solid media and further processed to obtain pure cultures, followed by downstream genetic characterization. Yeast species identity is established via sequencing of their internal transcribed spacer (ITS) region of the nuclear ribosomal RNA gene cluster, while global population structure of A. fumigatus is explored via microsatellite marker analysis. 
The protocol was successfully applied to isolate and characterize soil yeast and A. fumigatus populations in Cameroon, Canada, China, Costa Rica, Iceland, Peru, New Zealand, and Saudi Arabia. These findings revealed much-needed insights on global patterns in soil yeast diversity, as well as global population structure and antifungal resistance profiles of A. fumigatus. This paper presents the method of isolating both yeasts and A. fumigatus from international soil samples.

This protocol is an effective, speedy method of culturing yeasts and the mold Aspergillus fumigatus from large sets of soil samples in as little as 7 days. The methods can be easily modified to accommodate a range of incubation media and temperatures as needed for experiments.

Soil is host to an incredible amount of microbial life, with each gram containing up to billions of bacterial, archaeal, and fungal cells. Multicellular ...

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

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

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

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

Visualization of Cellular Interactions Between the Maize Sheaths and Fungi

Detached Maize Sheaths for Live-Cell Imaging of Infection by Fungal Foliar Maize Pathogens

We have optimized a protocol to inoculate maize leaf sheaths with hemibiotrophic and necrotrophic foliar pathogenic fungi. The method is modified from one originally applied to rice leaf sheaths and allows direct microscopic observation of fungal growth and development in living plant cells. Leaf sheaths collected from maize seedlings with two fully emerged leaf collars are inoculated with 20 &#181;L drops of 5 x 105 spores/mL fungal spore suspensions and incubated in humidity chambers at 23 &#176;C under continuous fluorescent light. After 24-72 h, excess tissue is removed with a razor blade to leave a single layer of epidermal cells, an optically clear sample that can be imaged directly without the necessity for chemical fixation or clearing. Plant and fungal cells remain alive for the duration of the experiment and interactions can be visualized in real-time. Sheaths can be stained or subjected to plasmolysis to study the developmental cytology and viability of host and pathogen cells during infection and colonization. Fungal strains transformed to express fluorescent proteins can be inoculated or co-inoculated on the sheaths for increased resolution and to facilitate the evaluation of competitive or synergistic interactions. Fungal strains expressing fluorescent fusion proteins can be used to track and quantify the production and targeting of these individual proteins in planta. Inoculated sheath tissues can be extracted to characterize nucleic acids, proteins, or metabolites. The use of these sheath assays has greatly advanced the detailed studies of the mechanisms of fungal pathogenicity in maize and also of fungal protein effectors and secondary metabolites contributing to pathogenicity.

Author Spotlight: Investigating Fungal Pathogenicity Mechanisms in Maize 

This manuscript details an optimized inoculation protocol that uses detached maize leaf sheaths for reproducible cytological, physiological, and molecular studies of maize interactions with fungal plant pathogens. The leaf sheaths facilitate real-time observation of cellular interactions between the living plant and fungus in unfixed tissues.

We have optimized a protocol to inoculate maize leaf sheaths with hemibiotrophic and necrotrophic foliar pathogenic fungi. The method is modified from one ...

Quantification of Arbuscular Mycorrhizal Fungi Colonization Rate in the Study of Invasive Alien Plants

Arbuscular mycorrhizal fungi (AMF) are widely distributed soil fungi in ecosystems and can form symbiotic associations (mycorrhizae) with the roots of most terrestrial plants. Plants provide carbon sources to AMF through mycorrhizal associations, while AMF hyphae can expand the range of nutrient absorption by roots and promote plant nutrient uptake. There are many different species of AMF, and the symbiotic relationships between different species of AMF and different plants vary. Invasive plants can enrich AMF species with better symbiotic capabilities through root exudates, promoting their growth and thereby increasing their colonization in invasive plant roots. At the same time, invasive plants can also disrupt the symbiotic relationship between AMF and native plants, affecting the local plant community, which is one of the mechanisms for successful plant invasion. The colonization rate of AMF in the roots of invasive and native plants indirectly reflects the role of AMF in the process of invasive plant invasion. In this method, collected plant roots can be processed directly or saved in a fixative for later batch processing. Through decolorization, acidification, staining, and destaining treatment of roots, the hyphae, spores, and arbuscular structures of AMF in the root system can be clearly observed. This method can be completed in a basic laboratory to observe and calculate the colonization rate of AMF in the root systems of invasive plants.

This protocol provides a simple and easy-to-use approach for determining the colonization rate of Arbuscular mycorrhizal fungi (AMF) in the roots of invasive plants.

Arbuscular mycorrhizal fungi (AMF) are widely distributed soil fungi in ecosystems and can form symbiotic associations (mycorrhizae) with the roots of ...

Production of Arbuscular Mycorrhizal (AM) Fungal Inoculum and Phenotypic Evaluation of Rice and AM Symbiosis Under Saline Conditions

Rice (Oryza sativa L.) is a vital food crop for more than half of the global population. However, its growth is severely impacted by saline soils, which present a significant challenge to crop production worldwide. Arbuscular mycorrhizal (AM) fungi, which form mutualistic symbiotic relationships with over 90% of agricultural plants and 80% of terrestrial plant species, have been shown to enhance the salt tolerance of rice plants. AM fungi are obligate symbionts that cannot complete their life cycle without a host root. Therefore, effectively utilizing plants to produce AM fungal inoculum is crucial for advancing research in this field. In this study, we present a series of robust methods that begin with generating sand inoculum containing spores of Rhizophagus irregularis using Allium tuberosum L. These methods include inoculating rice seedlings with the sand inoculum, analyzing the growth phenotype of mycorrhizal rice, and quantifying fungal colonization levels using trypan blue staining under salt stress. These approaches can efficiently generate AM fungal inoculum for further investigation into how AM symbiosis enhances the salinity tolerance of rice.

This article describes a protocol for generating arbuscular mycorrhizal (AM) fungi inoculum to investigate AM-enhanced salt stress tolerance in rice.

Rice (Oryza sativa L.) is a vital food crop for more than half of the global population. However, its growth is severely impacted by saline soils, which ...

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

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