Name: high-throughput siderophore screening from environmental samples: plant tissues, bulk soils, and rhizosphere soils
Uploaded: 2019-02-09T15:00:00.000+00:00
Duration: 12 min 36 s
Description: Watch this Scientific Journal Video about High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils at JoVE.com

The authors wish to thank Kalyani Muhunthan for assistance in laboratory procedures, Lee Opdahl for wheat genotype harvesting, the Washington State Concord Grape Research Council, and the Washington State University Center for Sustaining Agriculture and Natural Resources for a BIOAg grant to support this work. Additional funding was provided by the USDA/NIFA through Hatch project 1014527.

NOTE: Location of Field Site: Washington State University, Plant Pathology Farm (46&#176;46&#8217;38.0&#8221;N 117&#176;04&#8217;57.4&#8221;W). Seeds were sown using a mechanical planter on October 19, 2017. Each wheat genotype was planted in headrows, approximately 1 meter apart to avoid overlapping of root system. Plant and soil samples were collected on August 9, 2018, when plants were ready for harvest. Samples were gathered from three replicates of four wheat genotypes: PI561727, PI561725, Madsen, Lewjain.
1. Preparation of modified M9 medium
<ol>
	<li>Use Na2PO4&#8729;7H2O (12.8 g/200 mL), KH2PO4 (0.3 g/200 mL), NaCl (0.5 g/200 mL), and NH4Cl (1 g/20 mL) reagents to prepare the M9 salt solution.</li>
	<li>Use 18 g of MgSO4&#8729;7H2O in 100 mL of double deionized water (ddH2O) to prepare 0.75 M MgSO4&#8729;7H2O.</li>
	<li>Make 1 M of CaCl2&#8729;2H2O by adding 14.7 g of CaCl2&#8729;2H2O with 100 mL of ddH2O.</li>
	<li>Prepare the buffer solution by dissolving 6.048 g of PIPES into 156 mL of ddH2O with stirring. Adjust the pH to 6.8 with 5 M NaOH.</li>
	<li>Prepare 20% glucose (dextrose monohydrate) by dissolving 20 g of glucose into 100 mL of ddH2O.</li>
	<li>Prepare solutions and individually autoclave all but the glucose. Filter sterilize the glucose solution for addition to the M9 media after autoclaving (0.22 &#181;m).</li>
	<li>Prepare the modified M9 medium by mixing 156 mL of PIPES Buffer solution, 40 mL of M9 salts solution, 20 &#956;L of CaCl2&#183;2H2O solution, 266 &#956;L of MgSO4&#183;7H2O, and 4 mL of 20% glucose solutions together in the biosafety cabinet, using sterile technique.</li>
	<li>For preservation of the modified M9, seal the container, cover with aluminum foil to protect from UV, and place at 4 &#176;C.</li>
</ol>
2. Preparation of CAS-Fe-Agar medium
<ol>
	<li>Acid wash all glassware in 100 mM HCl/100 mM HNO3 for a minimum of 2 h prior to utilization in the CAS assay.</li>
	<li>Prepare an aluminum baking pan filled with laboratory grade sand and cover it with aluminum foil. Autoclave at 121 &#176;C for 30 min and set aside.</li>
	<li>Prepare HDTMA (hexadecyltrimethylammonium bromide) by adding 0.0365 g to 20 mL ddH2O and place the solution at 37 &#176;C to promote solubilization.</li>
	<li>Prepare 10 mM HCl and generate 1 mM FeCl3&#183;6H2O using 10 mM HCl as the solvent. Add 0.0302 g of CAS to 25 mL of ddH2O while gently stirring with a sterile magnetic stir bar. Then add 5 mL of 1 mM FeCl3&#183;6H2O (in 10 mM HCl) to the 25 mL of CAS solution while continuing to gently stir (the solution turns into dark reddish black color).</li>
	<li>Slowly add the 20 mL of HDTMA solution while gently stirring, into the Fe-CAS solution (this yields a dark blue solution).</li>
	<li>Prepare the buffer solution by dissolving 15.12 g of PIPES into 375 mL of ddH2O, with gentle stirring. Adjust the pH to 6.8 with 5 M NaOH. Add water to bring the volume to 450 mL. Add 5 g of agarose to the solution. Autoclave the PIPES buffer solution and the CAS-Fe solution at 121 &#176;C for 30 min.</li>
	<li>Carefully add the entirety of the CAS-Fe solution to the entirety of the PIPES Buffer in the biosafety cabinet after each of them is autoclaved.</li>
	<li>Place the mixed solution in a water bath at 50 &#176;C. 
	NOTE: Freshly prepare all the reagents in the CAS-Fe-Agar medium before each assay, as long-term storage at 50 &#176;C results in precipitation of the CAS-Fe complex, and cooling results in solidified medium.</li>
	<li>Place a sterile reagent boat in the sterile sand in the biosafety cabinet and heat to 50 &#176;C. Transfer the CAS-Fe-Agar to the boat, then quickly aliquot 100 &#181;L to each well in a clear, flat-bottom, sterile 96-well microplate.</li>
</ol>
3. Pyoverdine/EDTA standard preparation
<ol>
	<li>Pyoverdine preparation
	<ol>
		<li>Prepare 800 &#956;M pyoverdine standard (mixture of succinic acid, 2-hydroxy glutaramide, and succinaminde forms of pyoverdine &#8211; see Table of Materials), in previously prepared modified M9 medium.</li>
		<li>Dilute this solution into 400, 200, 100, 50, 25, 12.5, and 6.25 &#956;M solutions.</li>
	</ol>
	</li>
	<li>EDTA standard preparation
	<ol>
		<li>Add 0.594 g of disodium ethylenediaminetetraacetic acid (EDTA: C10H14N2Na2O8.2H2O), to 500 mL of previously prepared modified M9 medium to prepare 3200 &#956;M EDTA standard.</li>
		<li>Dilute this solution into 1600, 800, 400, 200, 100, 50, 25, 12.5, and 6.25 &#956;M solutions.</li>
	</ol>
	</li>
	<li>Standard curve generation
	<ol>
		<li>Add 100 &#956;L of each concentration of pyoverdine and EDTA to separate wells of a 96-well microplate containing 100 &#956;L of CAS-Fe Agar medium. Make duplicate technical replications of each concentration. Also, add blank wells with only M9 (no EDTA or pyoverdine).</li>
		<li>Using a microplate reader, measure absorbance (at 420 nm and 665 nm) after 1, 6, and 24 h incubation at 22 &#176;C, and use absorbance measurements to generate standard curves. 
		NOTE: For 420 nm measurements, subtract absorbance of blanks from the absorbance of pyoverdine or EDTA containing wells. For 665 nm, subtract absorbance of pyoverdine or EDTA containing wells from absorbance of blanks. Then, log10(&#181;M pyoverdine or EDTA) is regressed against the absorbance measurements. For ease of interpreting sample results, use absorbance as the x-axis and log10(&#181;M pyoverdine or EDTA) as the y-axis.</li>
	</ol>
	</li>
</ol>
4. Collection of environmental samples: soil and plant tissues
<ol>
	<li>Wash sampling equipment (shovels and scissors) with 0.22 &#181;m filtered ddH2O followed by 70% ethanol, and wipe with paper towels before sampling and in between samples to maintain sterile technique and reduce cross-contamination.</li>
	<li>Excise the plant tissues (grains and shoots) from plants in the field, and place them in a labeled plastic storage bag, leaving enough shoot stubble for ease in uprooting the desired soil and root samples.</li>
	<li>Excavate a small root ball approximately 15 cm deep and 23 cm wide and place it in a separate, labeled plastic bag for sample preparation in the laboratory environment. This step is similar to the methods of McPherson et al.27.</li>
	<li>Place all samples (grains, shoots, bulk soil, and root balls in separate bags) directly on ice and keep at 4 &#176;C until samples are processed for the siderophore production assay.</li>
	<li>Separate root associated soil samples into bulk, loosely-bound rhizosphere soil, and tightly-bound rhizosphere soil.
	<ol>
		<li>Take the root balls out of the bags. Gently shake off soil from the root ball. Shaken off soil, along with the soil left in the bag comprises the &#8220;bulk&#8221; soil.</li>
		<li>Use a rubber mallet to further remove soil from the root ball. This is the loosely-bound rhizosphere soil.</li>
		<li>Generate tightly-bound rhizosphere soil by taking roots with the tightly-bound sample and putting them in a centrifuge tube. Add 30 mL of ddH2O and vortex it for 2-3 minutes. Remove roots to get the tightly-bound rhizosphere soil slurry dilution.</li>
	</ol>
	</li>
</ol>
5. Preparation of siderophore enrichment cultures and CAS-Fe siderophore production assay
NOTE: All glassware should be acid washed prior to beginning the assays.
<ol>
	<li>Soil sample preparation (for each of the three soil sample types)
	<ol>
		<li>Homogenize each soil sample within the sample bag, by mixing and turning the soil as much as possible without opening the bag. 
		NOTE: This helps reduce natural soil spatial variability and aligns with normal soil sampling procedures. Other methods may be utilized to homogenize environmental samples, as appropriate, and depending on the experimental design.</li>
		<li>After each sample has been thoroughly mixed, aliquot and suspend 2.0 g of each soil in 20 mL of modified M9 medium, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
		<li>For tightly-bound rhizosphere samples, add 2 mL of the rhizosphere soil slurry to 20 mL of modified M9 medium, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
	</ol>
	</li>
	<li>Tissue sample (root, shoot and grain) preparation
	<ol>
		<li>Surface sterilize the sample with 70% ethanol. Macerate 2.0 g of fresh tissue in 20 mL of modified M9 medium using a blender on high for 30 seconds. Transfer the sample to a sterile 50 mL centrifuge tube, then dilute 10-3 to a total volume of 20 mL in a sterile 50 mL centrifuge tube with a sterile foam plug to allow aeration.</li>
	</ol>
	</li>
	<li>Enrichment of Siderophore Production Through Fe Limitation
	<ol>
		<li>Incubate 50 mL centrifuge tubes at room temperature and shake at 160 rpm.</li>
	</ol>
	</li>
</ol>
6. CAS-Fe agar assays for detection of siderophore production in environmental samples
<ol>
	<li>At 24, 48, and 72 h after initiating the enrichment culture, remove 1 mL subsamples from the enrichment tubes using sterile technique and centrifuge at 10,000 x g for 1 min in 2 mL centrifuge tubes to pelletize the cells.</li>
	<li>Collect the separated supernatant. Using sterile technique, add 100 &#956;L of the supernatant to 100 &#956;L solution of CAS-Fe-Agar in duplicate or triplicate in the microplate. Also add 100 &#181;L of sterile M9 medium (as blanks). Then incubate the plate at 28 &#176;C.</li>
	<li>Suspend the remaining supernatant and pellet for each sample (not added to microtiter plate) into its own, sterile 2 mL centrifuge tube. Add 400 &#956;L of sterile glycerol into each sample-supernatant tube and resuspend the pellet to create glycerol stocks. Freeze the stock at -80 &#176;C for later analyses. 
	NOTE: This step can be modified to generate glycerol stocks according to any preferred in-house protocol.</li>
	<li>Measure absorbance at 6, 24, 48, and 72 h, at 420 nm wavelength.</li>
	<li>Use standard curves generated from pyoverdine or EDTA to interpret sample absorbance measurements in terms of pyoverdine equivalents. 
	NOTE: Pyoverdine was determined to be a superior standard compared with EDTA (vide infra), so EDTA was not used to interpret results in the current study.</li>
</ol>

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The authors have no conflicts of interest to disclose.

The primary result of this work is the production of a new methodology that can be used to rapidly enrich for siderophore producing microbes while quantitatively measuring siderophore production/activity in the environmental sample. The methodology is quick, simple, and cost-effective, and the results show how it can be used to detect siderophore activity from complex and novel sample types (e.g., soil and plant tissue). The protocol also results in the production of glycerol stocks of the enrichment cultures, w...

Siderophores are important biomolecules involved primarily in iron-chelation for bioavailability, but with a wide array of additional purposes in terrestrial and aquatic ecosystems ranging from microbial quorum sensing, signaling to microbial plant-hosts, plant growth promotion, cooperation and competition within complex microbial communities1,2. Siderophores can be broadly classified according to their active sites and structural features, creating four basic types: carboxylate, hydroxamate, catecholate, and mixed types3,4. Many microorganisms are capable of excreting more than one type of siderophore5 and in complex communities, a vast majority of organisms biosynthesize the membrane receptors to allow the uptake of an even wider variety of siderophores1,6. Recent work indicates that siderophores are particularly important at the community level, and even in inter-kingdom communications and biogeochemical transfers7,8,9,10,11.
Chrome azurol sulphonate (CAS) has been used for over 30 years as a chelating agent to bind iron (Fe) in such a way that addition of ligands (i.e., siderophores) can result in dissociation of the CAS-Fe complex, creating an easily identifiable color change in the medium12. When the CAS is bound with Fe, the dye appears as a royal blue color, and as the CAS-Fe complex dissociates, the medium changes color according to the type of ligand used to scavenge the Fe13. The initial, liquid-based medium established by Schwyn and Neilands in 1987, has been modified in many ways to accommodate changing microbial targets14, growth habits and limitations15, as well as a variety of metals besides Fe, including aluminum, manganese, cobalt, cadmium nickel, lithium, zinc16, copper17, and even arsenic18.
Many human pathogens, as well as plant growth promoting microorganisms (PGPM) have been identified as siderophore-producing organisms3,19,20, and important rhizosphere and endophytic PGPM often test positive for siderophore-production4. The traditional Fe-based liquid method has been adapted to microtiter testing of isolates in cultivation for siderophore production21. However, these techniques fail to recognize the importance of the microbial community as a whole (the microbiome), in cooperation and potential regulation of siderophore production in soils and plant systems22. For that reason, we have developed a high-throughput community-level assessment of siderophore production from a given environment, based on the traditional CAS assay, but with replication, ease of measurement, reliability, and repeatability in a microplate assay.
In this study, a cost-effective, high-throughput CAS-Fe assay for detecting siderophore production is presented to assess the enrichment of siderophore production from complex samples (i.e., soil and plant tissue homogenates). Bulk, loosely-bound, and tightly-bound rhizosphere soil (in terms of how the soil was bound to the root) were obtained along with grain, shoot, and root tissues from four distinct wheat (Triticum aestivum L.) genotypes: Lewjain, Madsen, PI561725, and PI561727. It was hypothesized that fundamental differences in the wheat genotypes could result in differences in recruitment and selection of siderophore producing communities. Of particular interest is the difference between microbial communities associated with the PI561725 isogenic line, which is aluminum tolerant because it possesses ALMT1 (Aluminum-activated Malate Transporter 1), compared with the aluminum sensitive PI561727 isogenic line, which possesses a non-aluminum responsive form of the gene, almt123,24,25,26. The chief objective of the study was to develop a straightforward, rapid method of quantitatively assessing siderophore production in siderophore enrichment cultures of complex sample types while preserving the cultures for future work.

<table><tbody><tr><td>Agarose</td><td>Apex</td><td>LF451320014</td><td></td></tr><tr><td>Aluminum Baking Pan</td><td></td><td></td><td></td></tr><tr><td>Aluminum Foil</td><td></td><td></td><td></td></tr><tr><td>Ammonium chloride, granular</td><td>Fiesher Scientific</td><td>152315A</td><td></td></tr><tr><td>Autoclave and Sterilizer</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Calcium chloride dihydrate</td><td>Fiesher Scientific</td><td>171428</td><td></td></tr><tr><td>CAS (Chrome Azurol S)</td><td>Chem-Impex Int&#039;l Inc)</td><td>000331-27168</td><td></td></tr><tr><td>Dextrose Monohydrate (glucose), crystalline powder</td><td>Fiesher Scientific</td><td>1521754</td><td></td></tr><tr><td>EDTA, disodium salt, dihydrate, Crystal</td><td>J.T.Baker</td><td>JI2476</td><td></td></tr><tr><td>Glycerol, Anhydrous</td><td>Baker Analyzed</td><td>C22634</td><td></td></tr><tr><td>HDTMA (Cetyltrimethylammomonium Bromide</td><td>Reagent World</td><td>FZ0941</td><td></td></tr><tr><td>Hydrochloride acid</td><td>ACROS Organic</td><td>B0756767</td><td></td></tr><tr><td>Infinite M200 PRO plate reader</td><td>TECAN</td><td></td><td></td></tr><tr><td>Iron (III) chloride hexahydrate, 99%</td><td>ACROS Organic</td><td>A0342179</td><td></td></tr><tr><td>Laboratory Fume Hood</td><td>Thermo Scientific</td><td></td><td></td></tr><tr><td>Laboratory Incubator</td><td>VWR Scientific</td><td></td><td></td></tr><tr><td>Magnesium Sulfate</td><td>Fiesher Scientific</td><td>27855</td><td></td></tr><tr><td>Niric Acid, (69-70)%</td><td>J.T.Baker</td><td>72287</td><td></td></tr><tr><td>PIPES buffer, 98.5%</td><td>ACROS Organic</td><td>A0338723</td><td></td></tr><tr><td>Potassium phosphate, dibaisc,powder</td><td>J.T.Baker</td><td>J48594</td><td></td></tr><tr><td>Pyoverdine</td><td>SIGMA-ALDRICH</td><td>078M4094V</td><td></td></tr><tr><td>Sand</td><td></td><td></td><td></td></tr><tr><td>SI-600R Shaker</td><td>Lab Companion</td><td></td><td></td></tr><tr><td>Sodium chloride, granular</td><td>Fiesher Scientific</td><td>136539</td><td></td></tr><tr><td>Sodium hydroxide, pellets</td><td>J.T.Baker</td><td>G48K53</td><td></td></tr><tr><td>Sodium phosphate, dibasic heptahydrate, 99%</td><td>ACROS Organic</td><td>A0371705</td><td></td></tr></tbody></table>

high-throughput siderophore screening from environmental samples: plant tissues, bulk soils, and rhizosphere soils

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling in soils, to pathogen competition, plant growth promotion, and cross-kingdom signaling. Furthermore, siderophores are also of commercial interest in bioleaching and bioweathering of metal-bearing minerals and ores. A rapid, cost effective, and robust means of quantitatively assessing siderophore production in complex samples is key to identifying important aspects of the ecological ramifications of siderophore activity, including, novel siderophore producing microbes. The method presented here was developed to assess siderophore activity of in-tact microbiome communities, in environmental samples, such as soil or plant tissues. The samples were homogenized and diluted in a modified M9 medium (without Fe), and enrichment cultures were incubated for 3 days. Siderophore production was assessed in samples at 24, 48, and 72 hours (h) using a novel 96-well microplate CAS (Chrome azurol sulphonate)-Fe agar assay, an adaptation of the traditionally tedious and time-consuming colorimetric method of assessing siderophore activity, performed on individual cultivated microbial isolates. We applied our method to 4 different genotypes/Lines of wheat (Triticum aestivum L.), including Lewjain, Madsen, and PI561725, and PI561727 commonly grown in the inland Pacific Northwest. Siderophore production was clearly impacted by the genotype of wheat, and in the specific types of plant tissues observed. We successfully used our method to rapidly screen for the influence of plant genotype on siderophore production, a key function in terrestrial and aquatic ecosystems. We produced many technical replicates, yielding very reliable statistical differences in soils and within plant tissues. Importantly, the results show the proposed method can be used to rapidly examine siderophore production in complex samples with a high degree of reliability, in a manner that allows communities to be preserved for later work to identify taxa and functional genes.

A pyoverdine mixture biosynthesized by Pseudomonas fluorescens was used as a standard to interpret and quantify absorbance (at 420 nm) of samples in terms of pyoverdine equivalents in &#181;M. Figure 1 shows the relationship between absorbance (420 nm) and starting concentration of pyoverdine (Log10 molarity in &#181;M). EDTA did not provide an adequate standard because samples exhibited greater absorbance measurements than were attainable...

Watch this Scientific Journal Video about High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils at JoVE.com

High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils

We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

Siderophores (low-molecular weight metal chelating compounds) are important in various ecological phenomenon ranging from iron (Fe) biogeochemical cycling ...

high-throughput-siderophore-screening-from-environmental-samples

Universidad de Cartagena UDEC

Research

JoVE Journal

Environment

12.5K Views.  Washington State University. We present a protocol for rapid screening of environmental samples for siderophore potential contributing to micronutrient bioavailability and turnover in terrestrial systems.

Video: High-throughput Siderophore Screening from Environmental Samples: Plant Tissues, Bulk Soils, and Rhizosphere Soils

High-throughput Screening of Carbohydrate-degrading Enzymes Using Novel Insoluble Chromogenic Substrate Assay Kits

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper and food industries. A deeper understanding of CAZymes is important from both fundamental biology and industrial standpoints. Vast numbers of CAZymes exist in nature (especially in microorganisms) and hundreds of thousands have been cataloged and described in the carbohydrate active enzyme database (CAZy). However, the rate of discovery of putative enzymes has outstripped our ability to biochemically characterize their activities. One reason for this is that advances in genome and transcriptome sequencing, together with associated bioinformatics tools allow for rapid identification of candidate CAZymes, but technology for determining an enzyme's biochemical characteristics has advanced more slowly. To address this technology gap, a novel high-throughput assay kit based on insoluble chromogenic substrates is described here. Two distinct substrate types were produced: Chromogenic Polymer Hydrogel (CPH) substrates (made from purified polysaccharides and proteins) and Insoluble Chromogenic Biomass (ICB) substrates (made from complex biomass materials). Both CPH and ICB substrates are provided in a 96-well high-throughput assay system. The CPH substrates can be made in four different colors, enabling them to be mixed together and thus increasing assay throughput. The protocol describes a 96-well plate assay and illustrates how this assay can be used for screening the activities of enzymes, enzyme cocktails, and broths.

A high-throughput assay for enzyme screening is described. This multiplexed ready-to-use assay kit comprises of pre-chosen Chromogenic Polymer Hydrogel (CPH) substrates and complex Insoluble Chromogenic Biomass (ICB) substrates. Target enzymes are polysaccharide degrading endo-enzymes and proteases.

Carbohydrates active enzymes (CAZymes) have multiple roles in vivo and are widely used for industrial processing in the biofuel, textile, detergent, paper ...

Biochemistry

A Whole Cell Bioreporter Approach to Assess Transport and Bioavailability of Organic Contaminants in Water Unsaturated Systems

Bioavailability of contaminants is a prerequisite for their effective biodegradation in soil. The average bulk concentration of a contaminant, however, is not an appropriate measure for its availability; bioavailability rather depends on the dynamic interplay of potential mass transfer (flux) of a compound to a microbial cell and the capacity of the latter to degrade the compound. In water-unsaturated parts of the soil, mycelia have been shown to overcome bioavailability limitations by actively transporting and mobilizing organic compounds over the range of centimeters. Whereas the extent of mycelia-based transport can be quantified easily by chemical means, verification of the contaminant-bioavailability to bacterial cells requires a biological method. Addressing this constraint, we chose the PAH fluorene (FLU) as a model compound and developed a water unsaturated model microcosm linking a spatially separated FLU point source and the FLU degrading bioreporter bacterium Burkholderia sartisoli RP037-mChe by a mycelial network of Pythium ultimum. Since the bioreporter expresses eGFP in response of the PAH flux to the cell, bacterial FLU exposure and degradation could be monitored directly in the microcosms via confocal laser scanning microscopy (CLSM). CLSM and image analyses revealed a significant increase of the eGFP expression in the presence of P. ultimum compared to controls without mycelia or FLU thus indicating FLU bioavailability to bacteria after mycelia-mediated transport. CLSM results were supported by chemical analyses in identical microcosms. The developed microcosm proved suitable to investigate contaminant bioavailability and to concomitantly visualize the involved bacteria-mycelial interactions.

A whole cell bioreporter assay with Burkholderia sartisoli RP037-mChe was developed to detect fractions of an organic contaminant (i.e., fluorene) available for bacterial degradation after active transport by mycelia bridging air-filled pores in a water unsaturated model system.

Bioavailability of contaminants is a prerequisite for their effective biodegradation in soil. The average bulk concentration of a contaminant, however, is ...

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

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 ...

Single-throughput Complementary High-resolution Analytical Techniques for Characterizing Complex Natural Organic Matter Mixtures

Natural organic matter (NOM) is composed of a highly complex mixture of thousands of organic compounds which, historically, proved difficult to characterize. However, to understand the thermodynamic and kinetic controls on greenhouse gas (carbon dioxide [CO2] and methane [CH4]) production resulting from the decomposition of NOM, a molecular-level characterization coupled with microbial proteome analyses is necessary. Further, climate and environmental changes are expected to perturb natural ecosystems, potentially upsetting complex interactions that influence both the supply of organic matter substrates and the microorganisms performing the transformations. A detailed molecular characterization of the organic matter, microbial proteomics, and the pathways and transformations by which organic matter is decomposed will be necessary to predict the direction and magnitude of the effects of environmental changes. This article describes a methodological throughput for comprehensive metabolite characterization in a single sample by direct injection Fourier transform ion cyclotron resonance mass spectrometry (FTICR-MS), gas chromatography mass spectrometry (GC-MS), nuclear magnetic resonance (NMR) spectroscopy, liquid chromatography mass spectrometry (LC-MS), and proteomics analysis. This approach results in a fully-paired dataset which improves statistical confidence for inferring pathways of organic matter decomposition, the resulting CO2 and CH4 production rates, and their responses to environmental perturbation. Herein we present results of applying this method to NOM samples collected from peatlands; however, the protocol is applicable to any NOM sample (e.g., peat, forested soils, marine sediments, etc.).

This protocol describes a single throughput for complementary analytical and omics techniques culminating in a fully-paired characterization of natural organic matter and microbial proteomics in different ecosystems. This approach permits robust comparisons for identifying metabolic pathways and transformations important for describing greenhouse gas production and predicting responses to environmental change.

Natural organic matter (NOM) is composed of a highly complex mixture of thousands of organic compounds which, historically, proved difficult to ...

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 ...

Bioprospecting of Extremophilic Microorganisms to Address Environmental Pollution

Geothermal springs are rich in various metal ions due to the interaction between rock and water that takes place in the deep aquifer. Moreover, due to seasonality variation in pH and temperature, fluctuation in element composition is periodically observed within these extreme environments, influencing the environmental microbial communities. Extremophilic microorganisms that thrive in volcanic thermal vents have developed resistance mechanisms to handle several metal ions present in the environment, thus taking part to complex metal biogeochemical cycles. Moreover, extremophiles and their products have found an extensive foothold in the market, and this holds true especially for their enzymes. In this context, their characterization is functional to the development of biosystems and bioprocesses for environmental monitoring and bioremediation. To date, the isolation and cultivation under laboratory conditions of extremophilic microorganisms still represent a bottleneck for fully exploiting their biotechnological potential. This work describes a streamlined protocol for the isolation of thermophilic microorganisms from hot springs as well as their genotypical and phenotypical identification through the following steps: (1) Sampling of microorganisms from geothermal sites ("Pisciarelli", a volcanic area of Campi Flegrei in Naples, Italy); (2) Isolation of heavy metal resistant microorganisms; (3) Identification of microbial isolates; (4) Phenotypical characterization of the isolates. The methodologies described in this work might be generally applied also for the isolation of microorganisms from other extreme environments.

The isolation of heavy metal-resistant microbes from geothermal springs is a hot topic for the development of bioremediation and environmental monitoring biosystems. This study provides a methodological approach for isolating and identifying heavy metal tolerant bacteria from hot springs.

Geothermal springs are rich in various metal ions due to the interaction between rock and water that takes place in the deep aquifer. Moreover, due to ...

High Throughput Co-culture Assays for the Investigation of Microbial Interactions

The study of interactions between microorganisms has led to numerous discoveries, from novel antimicrobials to insights in microbial ecology. Many approaches used for the study of microbial interactions require specialized equipment and are expensive and time intensive. This paper presents a protocol for co-culture interaction assays that are inexpensive, scalable to large sample numbers, and easily adaptable to numerous experimental designs. Microorganisms are cultured together, with each well representing one pairwise combination of microorganisms. A test organism is cultured on one side of each well and first incubated in monoculture. Subsequently, target organisms are simultaneously inoculated onto the opposite side of each well using a 3D-printed inoculation stamp. After co-culture, the completed assays are scored for visual phenotypes, such as growth or inhibition. These assays can be used to confirm phenotypes or identify patterns among isolates of interest. Using this simple and effective method, users can analyze combinations of microorganisms rapidly and efficiently. This co-culture approach is applicable to antibiotic discovery as well as culture-based microbiome research and has already been successfully applied to both applications.

The co-culture interaction assays presented in this protocol are inexpensive, high throughput, and simple. These assays can be used to observe microbial interactions in co-culture, identify interaction patterns, and characterize the inhibitory potential of a microbial strain of interest against human and environmental pathogens.

The study of interactions between microorganisms has led to numerous discoveries, from novel antimicrobials to insights in microbial ecology. Many ...

Immunology and Infection

Using Coculture to Detect Chemically Mediated Interspecies Interactions

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by secreting metabolites. These metabolites have the potential to modulate the physiology and differentiation of their microbial neighbors and are likely important factors in the establishment and maintenance of complex microbial communities. We have developed a fluorescence-based coculture screen to identify such chemically mediated microbial interactions. The screen involves combining a fluorescent transcriptional reporter strain with environmental microbes on solid media and allowing the colonies to grow in coculture. The fluorescent transcriptional reporter is designed so that the chosen bacterial strain fluoresces when it is expressing a particular phenotype of interest (i.e. biofilm formation, sporulation, virulence factor production, etc.) Screening is performed under growth conditions where this phenotype is not expressed (and therefore the reporter strain is typically nonfluorescent). When an environmental microbe secretes a metabolite that activates this phenotype, it diffuses through the agar and activates the fluorescent reporter construct. This allows the inducing-metabolite-producing microbe to be detected: they are the nonfluorescent colonies most proximal to the fluorescent colonies. Thus, this screen allows the identification of environmental microbes that produce diffusible metabolites that activate a particular physiological response in a reporter strain. This publication discusses how to: a) select appropriate coculture screening conditions, b) prepare the reporter and environmental microbes for screening, c) perform the coculture screen, d) isolate putative inducing organisms, and e) confirm their activity in a secondary screen. We developed this method to screen for soil organisms that activate biofilm matrix-production in Bacillus subtilis; however, we also discuss considerations for applying this approach to other genetically tractable bacteria.

Bacteria produce secreted compounds that have the potential to affect the physiology of their microbial neighbors. Here we describe a coculture screen that allows detection of such chemically mediated interspecies interactions by mixing soil microbes with fluorescent transcriptional reporter strains of Bacillus subtilis on solid media.

In nature, bacteria rarely exist in isolation; they are instead surrounded by a diverse array of other microorganisms that alter the local environment by ...

Biology

Isolation and Screening from Soil Biodiversity for Fungi Involved in the Degradation of Recalcitrant Materials

Environmental pollution is an increasing problem, and identifying fungi involved in the bioremediation process is an essential task. Soil hosts an incredible diversity of microbial life and can be a good source of these bioremediative fungi. This work aims to search for soil fungi with bioremediation potential by using different screening tests. Mineral culture media supplemented with recalcitrant substances as the sole carbon source were used as growth tests. First, soil dilutions were plated on Petri dishes with mineral medium amended with humic acids or lignocellulose. The growing fungal colonies were isolated and tested on different substrates, such as complex mixtures of hydrocarbons (petrolatum and used motor oil) and powders of different plastic polymers (PET, PP, PS, PUR, PVC). Qualitative enzymatic tests were associated with the growth tests to investigate the production of esterases, laccases, peroxidases, and proteases. These enzymes are involved in the main degradation processes of recalcitrant material, and their constitutive secretion by the examined fungal strains could have the potential to be exploited for bioremediation. More than 100 strains were isolated and tested, and several isolates with good bioremediation potential were found. In conclusion, the described screening tests are an easy and low-cost method to identify fungal strains with bioremediation potential from the soil. In addition, it is possible to tailor the screening tests for different pollutants, according to requirements, by adding other recalcitrant substances to minimal culture media.

Here, we present a protocol for screening soil biodiversity to look for fungal strains involved in the degradation of recalcitrant materials. First, fungal strains able to grow on humic acids or lignocellulose are isolated. Their activity is then tested both in enzymatic assays and on pollutants such as hydrocarbons and plastics.