This work was supported by grants from DOE Great Lakes Bioenergy Research Center (DOE BER office of Science DE-FC02-07ER64494). We are grateful to Dr. Xudong Zhang and his group members for helpful technical discussions and invaluable comments on finalizing the protocol.

1. Sample Preparation and Acid Extraction
<ol>
	<li>Freeze-dry soil samples after field collection.</li>
	<li>Grind and homogenize soil samples using a ball mill, soil grinder, or a mortar and pestle.</li>
	<li>Weigh soil samples (containing &gt; 0.3 mg N) into a 25 mL hydrolysis flask.</li>
	<li>Add 10 mL 6M HCl into each hydrolysis flask, fill N2 gas in the flasks, and cap tightly.</li>
	<li>Hydrolyze at 105 &deg;C in an incubator for 8 hours using an auto timer switch.</li>
</ol>
2. Sample Purification
<ol>
	<li>Remove the flasks from the incubator; cool to room temperature.</li>
	<li>Add 100 &mu;L internal standard myo-inositol (1 mg mL-1 in water) to each flask; mix by swirling.</li>
	<li>Set up plastic funnels draining into 200 mL pear shaped flasks with ST/NS 24/40 joints, set on plastic cups for stability.</li>
	<li>Fold Whatman #2 Qualitative Circles (11 cm diameter) into quarters and set into funnels.</li>
	<li>Swirl each hydrolysis flask, and pour slurry into funnel to filter (you may further rinse each flask with ~3 mL of water).</li>
	<li>Dry the filtrate using a rotary evaporator at ~45 &deg;C, applying vacuum.</li>
	<li>Resuspend the dried residue from each pear flask with 3~5 mL water (use ultrasonic bath if desired), and pour into a 40 mL Teflon tube; rinse the flask with a second aliquot of water.</li>
	<li>Adjust the pH to 6.6 - 6.8 using 1M KOH solution to precipitate metal ions and other organic molecules.</li>
	<li>Remove the precipitates by centrifugation at 2000 rcf for 10 minutes.</li>
	<li>Pour the supernatant into a 40 mL glass tube, cover the tube opening with parafilm, freeze at -20 &deg;C, then poke holes in the parafilm.</li>
	<li>Freeze-dry the frozen supernatant to remove all liquid.</li>
	<li>Dissolve the residue with 3 mL dry methanol, vortexing thoroughly (use ultrasonic bath if desired); cap the tube, and then centrifuge at 2000 rcf for 10 minutes to settle out salts.</li>
	<li>Transfer the supernatant to a 3 mL conical reaction vial; evaporate to dryness by RapidVap machine at 45 &deg;C (or under a gentle stream of dry nitrogen gas if desired).</li>
	<li>To each vial, add 100 &mu;L recovery standard N-methylglucamine (1 mg mL-1 in water) and 1 mL H2O, cover the vial mouths with parafilm, freeze at -20 &deg;C, perforate the parafilm, and then freeze dry.</li>
	<li>Make standards: add 100 &mu;L muramic acid (0.5 mg mL-1 in methanol), 100 &mu;L glucosamine (1 mg mL-1 in water), 100 &mu;L myo-inositol (1 mg mL-1 in water), 100 &mu;L N-methylglucamine (1 mg mL-1 in water), and 1 mL H2O, parafilm each vial, freeze at -20 &deg;C, perforate the parafilm, then freeze-dry.</li>
</ol>
3. Derivatization
<ol>
	<li>Prepare derivatization reagent containing 32 mg mL-1 hydroxylamine hydrochloride and 40 mg mL-1 4-dimethylamino-pyridine in pyridine-methanol (4:1 v/v).</li>
	<li>Add 300 &mu;L of the derivatization reagent to each of the reactivials, cap tightly, and vortex thoroughly.</li>
	<li>Put the vials in 75-80 &deg;C water bath for 35 minutes (well sealed the vials to avoid allowing any water to enter the vials).</li>
	<li>Remove vials from the water bath, and cool to room temperature.</li>
	<li>Add 1 mL acetic anhydride to each of the 3 mL reactivials, cap tightly, and vortex thoroughly, then reheat in 75-80 &deg;C water bath for 25 minutes.</li>
	<li>Remove vials from the water bath, and cool to room temperature.</li>
</ol>
4. Separation and Measurement
<ol>
	<li>Add 1.5 mL dichloromethane to each vial, cap tightly, and vortex thoroughly.</li>
	<li>Add 1 mL 1M HCl to each vial, cap tightly and vortex thoroughly to allow the solutions to sit undisturbed until the two phrases separate, aspirate and discard the top (aqueous) phase using 1000 &mu;L pipettor.</li>
	<li>In the same fashion, extract the organic phase 3 times but with 1 mL H2O (with the last washing step, take special care to ensure that all of the aqueous top phase has been removed).</li>
	<li>Dry the final solution using RapidVap at 45 &deg;C (or dry using nitrogen gas if desired).</li>
	<li>Dissolve in 300 &mu;L ethyl acetate-hexane (1:1 v/v) and transfer to 2 mL amber screw-cap vials with a small-volume insert and cap tightly.</li>
	<li>For quantification, analyze by GC-FID using a fused silica nonpolar capillary column: 30 m long, 0.25 mm i.d., 0.25 um film thickness; stationary phase 5% phenyl-, 95% methyl-polysiloxane (DB-5 or equivalent) with hydrogen or helium as the carrier gas, at 0.5 mL min-1 constant flow. The chromatography is better on premium "inert" phases and hydrogen carrier gas, but is acceptable on the more common varieties and with helium. The detector settings are 300 &deg;C; 400 mL min-1 air and 30 mL min-1 for both the nitrogen and hydrogen makeup gases. The injection and oven parameters are as follows: 1 &mu;L split injection (30:1) with the GC inlet set at 250 &deg;C; initial oven temperature, 120 &deg;C; hold 1 min.; increase the oven temperature at 10 &deg;C min-1 to 250 &deg;C; hold 2.5 min.; ramp to 270 &deg;C at 20 &deg;C min-1; hold 2 minutes; ramp to 290 &deg;C at 20 &deg;C min-1, hold 5 min. Adjust the carrier gas flow rate so that the inositol, glucosamine and muramic acid derivatives elute at 250 &deg;C, and the N-methylglucamine elutes at 270 &deg;C.</li>
</ol>
5. Derivative Validation
<ol>
	<li>Use soft ionization LC-ESI-TOF-MS to identify the molecular ion and determine the molecular weight of the derivative.</li>
	<li>Use GC-EI-MS-SIM or GC-EI-MSMS for sensitivity enhancement to investigate targeted ions of the derivative.</li>
	<li>Use multiple isotopically labeled reagents for preparation of the derivatives, and then use the mass shift of those isotopically labeled derivatives in MS to investigate molecular ion and ion fragments.</li>
</ol>
6. Representative Results
The protocol of the method mainly comprises four steps: acid digestion, sample purification, derivatization and GC determination (Figure 1). An example of the analysis for glucosamine and muramic acid from standard stocks and from a soil sample is shown in Figure 2. Besides glucosamine and muramic acid, mannosamine and galactosamine (two isomers of glucosamine) can also be determined simultaneously using the method. Based on response factors of standards with respect to the internal standard myo-inositol, we can quantify these biomarkers in soil samples. The recovery standard has been used to qualitatively monitor the derivatization process. The schemes for formation of the aldononitrile acetate derivatized glucosamine and muramic acid are shown in Figure 3.
The proposed structures of the derivatives were determined by GC-EIMS-SIM for sensitivity enhancement, or soft ionization LC-ESI-TOF-MS8; the proposed structures of the ion fragments formed upon electron ionization were studied by comparing the ion spectra of the samples prepared with various isotope incorporations11. The Figure 4 shows the mass shift of the dominant ion m/z 187 of aldononitrile acetate derivatized glucosamine in three scenarios, prepared with non-labeled agents, D-acetic anhydrite, and U-13C-glucosamine. Other detailed information and explanations can be referred to our recent publications8, 11. This strategy could serve as a model to study derivative chemistry.
<img alt="Figure 1" src="/files/ftp_upload/3767/3767fig1.jpg" /> 
Figure 1. Measurement flow chart of the protocol for analysis of glucosamine and muramic acid in soil samples. The protocol mainly contains 4 steps: Acid digestion, Purification, Derivatization and GC determination.
<img alt="Figure 2" src="/files/ftp_upload/3767/3767fig2.jpg" /> 
Figure 2. GC-FID chromatograms of aldononitrile acetates of inositol, glucosamine, muramic acid, mannosamine, galactosamine and methylglucamine for standards and soil.
<img alt="Figure 3" src="/files/ftp_upload/3767/3767fig3.jpg" /> 
Figure 3. Schemes for formation of the aldononitrile acetate derivatized glucosamine and muramic acid. The number 1 represents nitrile reaction. The number 2 represents acetylation.
<img alt="Figure 4" src="/files/ftp_upload/3767/3767fig4.jpg" /> 
Figure 4. Mass spectra of aldononitrile acetate derivatized glucosamine associated with the dominant ion structures under EI mode prepared with (A) non-labeled agents, (B) D-acetic anhydrite, (C) U-13C-glucosamine. Star represents heavy isotope atom or isotope group. Please <a href="https://www.jove.com/files/ftp_upload/3767/3767fig4_lg.jpg">click here</a> to see a larger version of this figure.

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	<li>Zelles, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fatty+acid+patterns+of+phospholipids+and+lipopolysaccharides+in+the+characterisation+of+microbial+communities+in+soil:+a+review.">Fatty acid patterns of phospholipids and lipopolysaccharides in the characterisation of microbial communities in soil: a review.</a> Biology and Fertility of Soils. 29, 111-129 (1999).</li><li>Kirk, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+of+studying+soil+microbial+diversity.">Methods of studying soil microbial diversity.</a> Journal of Microbiological Methods. 58, 169-188 (2004).</li><li>Joergensen, R. G., Emmerling, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+evaluating+human+impact+on+soil+microorganisms+based+on+their+activity,+biomass,+and+diversity+in+agricultural+soils.">Methods for evaluating human impact on soil microorganisms based on their activity, biomass, and diversity in agricultural soils.</a> Journal of Plant Nutrition and Soil Science. 169, 295-309 (2006).</li><li>Guggenberger, G., Frey, S. D., Six, J., Paustian, K., Elliott, E. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bacterial+and+fungal+cell-wall+residues+in+conventional+and+no-tillage+agroecosystems.">Bacterial and fungal cell-wall residues in conventional and no-tillage agroecosystems.</a> Soil Science Society of America Journal. 63, 1188-1198 (1999).</li><li>Amelung, W., Lal, R., Kimble, J. 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C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Mass+spectrometric+characterization+of+amino+sugar+aldononitrile+acetate+derivatives+used+for+isotope+enrichment+assessment+of+microbial+residues.">Mass spectrometric characterization of amino sugar aldononitrile acetate derivatives used for isotope enrichment assessment of microbial residues.</a> Soil Biology and Biochemistry. 42, 904-909 (2010).</li><li>Liang, C., Pedersen, J. A., Balser, T. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aminoglycoside+antibiotics+may+interfere+with+microbial+amino+sugar+analysis.">Aminoglycoside antibiotics may interfere with microbial amino sugar analysis.</a> Journal of Chromatography A. 1216, 5296-5301 (2009).</li></ol>

We have nothing to disclose.

The presented GC-based method for the analysis of aldononitrile acetate derivatized glucosamine and muramic acid provides a relatively rapid method to quantify these amino sugars, extracted from soil. The derivatives are chemically stable, and can be determined in one analysis. The method is not restricted to soil samples, and can be simplified for samples from non-soil matrix.
The vacuum pump used in this method is built to be resistant to acid. We further suggest setting up a base trap to p...

<table border="1">
 <tbody>
 <tr>
 <td>Name of the reagent</td>
 <td>Company</td>
 <td>Catalogue number</td>
 </tr>
 <tr>
 <td>Muramic acid</td>
 <td>Sigma-Aldrich</td>
 <td>M2503-25MG</td>
 </tr>
 <tr>
 <td>D-(+)-glucosamine hydrochloride</td>
 <td>Sigma-Aldrich</td>
 <td>G1514-100G</td>
 </tr>
 <tr>
 <td>N-methyl-D-glucamine</td>
 <td>Sigma-Aldrich</td>
 <td>M2004-500G</td>
 </tr>
 <tr>
 <td>Myo-inositol</td>
 <td>Fisher Scientific</td>
 <td>A307003G025</td>
 </tr>
 <tr>
 <td>Methanol (dry)</td>
 <td>Acros organics</td>
 <td>AC326950010</td>
 </tr>
 <tr>
 <td>4-dimethylamino-pyridine</td>
 <td>Acros organics</td>
 <td>AC148270050</td>
 </tr>
 <tr>
 <td>Ethyl acetate</td>
 <td>VWR International</td>
 <td>BJGC100-4</td>
 </tr>
 <tr>
 <td>Hydroxlamine hydrochloride </td>
 <td>Fisher Scientific</td>
 <td>H330-100</td>
 </tr>
 <tr>
 <td>Pyridine</td>
 <td>Fisher Scientific</td>
 <td>P368-500</td>
 </tr>
 <tr>
 <td>Acetic anhydride</td>
 <td>Fisher Scientific</td>
 <td>A10-100</td>
 </tr>
 <tr>
 <td>Dichloromethane (Methylene chloride)</td>
 <td>Fisher Scientific</td>
 <td>D37-500</td>
 </tr>
 <tr>
 <td>Hexane</td>
 <td>Fisher Scientific</td>
 <td>H303-4</td>
 </tr>
 <tr>
 <td>Hydrochloric acid (6M)</td>
 <td>Fisher Scientific</td>
 <td>S456-4</td>
 </tr>
 <tr>
 <td>Hydroxylamine hydrochloride-15N</td>
 <td>Icon services</td>
 <td>IN5280</td>
 </tr>
 <tr>
 <td>Acetic anhydride-2H (D6C4O3)</td>
 <td>Acros organics</td>
 <td>AC174670050</td>
 </tr>
 <tr>
 <td>D-glucose-U-13C</td>
 <td>Cambridge isotope lab</td>
 <td>CLM-1396-1</td>
 </tr>
 <tr>
 <td>Ammonium sufate-15N </td>
 <td>Cambridge isotope lab</td>
 <td>NLM-713-1</td>
 </tr>
 <tr>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 <td>&nbsp;</td>
 </tr>
 <tr>
 <td>Name of the equipment</td>
 <td>Company</td>
 <td>Type</td>
 </tr>
 <tr>
 <td>Rapid-Vap</td>
 <td>LabConco</td>
 <td>790002 </td>
 </tr>
 <tr>
 <td>Vacum pump</td>
 <td>KNF Neuberger</td>
 <td>D-79112</td>
 </tr>
 <tr>
 <td>Hydrolysis flask</td>
 <td>Fisher Scientific </td>
 <td>06 423A</td>
 </tr>
 <tr>
 <td>Derivatization microvial</td>
 <td>Fisher Scientific</td>
 <td>06-100E</td>
 </tr>
 <tr>
 <td>GC</td>
 <td>Hewlett-Packard</td>
 <td>6890</td>
 </tr>
 <tr>
 <td>MS</td>
 <td>Hewlett-Packard</td>
 <td>5972</td>
 </tr>
 <tr>
 <td>LC-ESI-TOF-MS</td>
 <td>Agilent</td>
 <td>An Agilent 1200 series HPLC system coupled to an Agilent LC/MSD-TOF </td>
 </tr>
 </tbody>
</table>

gc-based detection of aldononitrile acetate derivatized glucosamine and muramic acid for microbial residue determination in soil

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. Current strategies for microbial analysis include both traditional laboratory culture-dependent techniques and those based on direct extraction and determination of certain biomarkers1, 2. Few among the diversity of microbial species inhabiting soil can be cultured, so culture-dependent methods introduce significant biases, a limitation absent in biomarker analysis.
The glucosamine, mannosamine, galactosamine and muramic acid have been well served as measures of both the living and dead microbial mass, of these the glucosamine (most abundant) and muramic acid (uniquely from bacterial cell) are most important constituents in the soil systems3, 4. However, the lack of knowledge on the analysis restricts the wide popularization among scientific peers. Among all existing analytical methods, derivatization to aldononitrile acetates followed by GC-based analysis has emerged as a good option with respect to optimally balancing precision, sensitivity, simplicity, good chromatographic separation, and stability upon sample storage5.
Here, we present a detailed protocol for a reliable and relatively simple analysis of glucosamine and muramic acid from soil after their conversion to aldononitrile acetates. The protocol mainly comprises four steps: acid digestion, sample purification, derivatization and GC determination. The step-by-step procedure is modified according to former publications6, 7. In addition, we present a strategy to structurally validate the molecular ion of the derivative and its ion fragments formed upon electron ionization. We applied GC-EI-MS-SIM, LC-ESI-TOF-MS and isotopically labeled reagents to determine the molecular weight of aldononitrile acetate derivatized glucosamine and muramic acid; we used the mass shift of isotope-labeled derivatives in the ion spectrum to investigate ion fragments of each derivatives8. In addition to the theoretical elucidation, the validation of molecular ion of the derivative and its ion fragments will be useful to researchers using &delta;13C or ion fragments of these biomarkers in biogeochemical studies9, 10.

Watch this Scientific Journal Video about GC-based Detection of Aldononitrile Acetate Derivatized Glucosamine and Muramic Acid for Microbial Residue Determination in Soil at JoVE.com

GC-based Detection of Aldononitrile Acetate Derivatized Glucosamine and Muramic Acid for Microbial Residue Determination in Soil

We describe a method protocol for the GC-based analysis of the aldonitrile acetate derivatives of glucosamine and muramic acid extracted from soil. For elucidation of the chemical mechanism, we also present a strategy to confirm the structure of the derivative and the ion fragments formed upon electron ionization.

Quantitative approaches to characterizing microorganisms are crucial for a broader understanding of the microbial status and function within ecosystems. ...

gc-based-detection-aldononitrile-acetate-derivatized-glucosamine

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16.8K Views.  University of Wisconsin, Madison. We describe a method protocol for the GC-based analysis of the aldonitrile acetate derivatives of glucosamine and muramic acid extracted from soil. For elucidation of the chemical mechanism, we also present a strategy to confirm the structure of the derivative and the ion fragments formed upon electron ionization.

Video: GC-based Detection of Aldononitrile Acetate Derivatized Glucosamine and Muramic Acid for Microbial Residue Determination in Soil

Microbial Communities in Nature and Laboratory - Interview

Direct Detection of the Acetate-forming Activity  of the Enzyme Acetate Kinase

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of acetate to acetyl phosphate utilizing ATP as a substrate. Acetate kinases are ubiquitous in the Bacteria, found in one genus of Archaea, and are also present in microbes of the Eukarya6. The most well characterized acetate kinase is that from the methane-producing archaeon Methanosarcina thermophila7-14. An acetate kinase which can only utilize PPi but not ATP in the acetyl phosphate-forming direction has been isolated from Entamoeba histolytica, the causative agent of amoebic dysentery, and has thus far only been found in this genus15,16.
In the direction of acetyl phosphate formation, acetate kinase activity is typically measured using the hydroxamate assay, first described by Lipmann17-20, a coupled assay in which conversion of ATP to ADP is coupled to oxidation of NADH to NAD+ by the enzymes pyruvate kinase and lactate dehydrogenase21,22, or an assay measuring release of inorganic phosphate after reaction of the acetyl phosphate product with hydroxylamine23. Activity in the opposite, acetate-forming direction is measured by coupling ATP formation from ADP to the reduction of NADP+ to NADPH by the enzymes hexokinase and glucose 6-phosphate dehydrogenase24.
Here we describe a method for the detection of acetate kinase activity in the direction of acetate formation that does not require coupling enzymes, but is instead based on direct determination of acetyl phosphate consumption. After the enzymatic reaction, remaining acetyl phosphate is converted to a ferric hydroxamate complex that can be measured spectrophotometrically, as for the hydroxamate assay. Thus, unlike the standard coupled assay for this direction that is dependent on the production of ATP from ADP, this direct assay can be used for acetate kinases that produce ATP or PPi.

A method for the determination of acetate kinase activity is described. This assay utilizes a direct reaction for determining enzyme activity and kinetics of acetate kinase in the acetate-forming direction with different phosphoryl acceptors. Furthermore, this method can be utilized for assaying other acetyl phosphate or acetyl-CoA utilizing enzymes.

Acetate kinase, a member of the acetate and sugar kinase-Hsp70-actin (ASKHA) enzyme superfamily1-5, is responsible for the reversible phosphorylation of ...

The Use of Chemostats in Microbial Systems Biology

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated including macromolecular synthesis, metabolism and ultimately, commitment to the cell division cycle. The chemostat, a method of experimentally controlling cell growth rate, provides a powerful means of systematically studying how growth rate impacts cellular processes - including gene expression and metabolism - and the regulatory networks that control the rate of cell growth. When maintained for hundreds of generations chemostats can be used to study adaptive evolution of microbes in environmental conditions that limit cell growth. We describe the principle of chemostat cultures, demonstrate their operation and provide examples of their various applications. Following a period of disuse after their introduction in the middle of the twentieth century, the convergence of genome-scale methodologies with a renewed interest in the regulation of cell growth and the molecular basis of adaptive evolution is stimulating a renaissance in the use of chemostats in biological research.

Cell growth rate is a regulated process and a primary determinant of cell physiology. Continuous culturing using chemostats enables extrinsic control of cell growth rate by nutrient limitation facilitating the study of molecular networks that control cell growth and how those networks evolve to optimize cell growth.

Cells regulate their rate of growth in response to signals from the external world. As the cell grows, diverse cellular processes must be coordinated ...

Isolation of Native Soil Microorganisms with Potential for Breaking Down Biodegradable Plastic Mulch Films Used in Agriculture

Fungi native to agricultural soils that colonized commercially available biodegradable mulch (BDM) films were isolated and assessed for potential to degrade plastics. Typically, when formulations of plastics are known and a source of the feedstock is available, powdered plastic can be suspended in agar-based media and degradation determined by visualization of clearing zones. However, this approach poorly mimics in situ degradation of BDMs. First, BDMs are not dispersed as small particles throughout the soil matrix. Secondly, BDMs are not sold commercially as pure polymers, but rather as films containing additives (e.g. fillers, plasticizers and dyes) that may affect microbial growth. The procedures described herein were used for isolates acquired from soil-buried mulch films. Fungal isolates acquired from excavated BDMs were tested individually for growth on pieces of new, disinfested BDMs laid atop defined medium containing no carbon source except agar. Isolates that grew on BDMs were further tested in liquid medium where BDMs were the sole added carbon source. After approximately ten weeks, fungal colonization and BDM degradation were assessed by scanning electron microscopy. Isolates were identified via analysis of ribosomal RNA gene sequences. This report describes methods for fungal isolation, but bacteria also were isolated using these methods by substituting media appropriate for bacteria. Our methodology should prove useful for studies investigating breakdown of intact plastic films or products for which plastic feedstocks are either unknown or not available. However our approach does not provide a quantitative method for comparing rates of BDM degradation.

Plastic films labeled "biodegradable" are commercially available for agricultural use as mulches. Tillage represents an attractive disposal method, but degradation under field conditions is poorly understood. The purpose of this study was to develop methods for isolating native soil fungi and bacteria that colonize plastic mulch films after field burial.

Fungi native to agricultural soils that colonized commercially available biodegradable mulch (BDM) films were isolated and assessed for potential to ...

Pyrosequencing for Microbial Identification and Characterization

Pyrosequencing is a versatile technique that facilitates microbial genome sequencing that can be used to identify bacterial species, discriminate bacterial strains and detect genetic mutations that confer resistance to anti-microbial agents. The advantages of pyrosequencing for microbiology applications include rapid and reliable high-throughput screening and accurate identification of microbes and microbial genome mutations. Pyrosequencing involves sequencing of DNA by synthesizing the complementary strand a single base at a time, while determining the specific nucleotide being incorporated during the synthesis reaction. The reaction occurs on immobilized single stranded template DNA where the four deoxyribonucleotides (dNTP) are added sequentially and the unincorporated dNTPs are enzymatically degraded before addition of the next dNTP to the synthesis reaction. Detection of the specific base incorporated into the template is monitored by generation of chemiluminescent signals. The order of dNTPs that produce the chemiluminescent signals determines the DNA sequence of the template. The real-time sequencing capability of pyrosequencing technology enables rapid microbial identification in a single assay. In addition, the pyrosequencing instrument, can analyze the full genetic diversity of anti-microbial drug resistance, including typing of SNPs, point mutations, insertions, and deletions, as well as quantification of multiple gene copies that may occur in some anti-microbial resistance patterns.

Pyrosequencing is a versatile technique that facilitates microbial genome sequencing that can be used to identify bacterial species, discriminate bacterial strains, and detect genetic mutations that confer resistance to anti-microbial agents. In this video, the procedure for microbial amplicon generation, amplicon pyrosequencing, and DNA sequence analysis will be demonstrated.

Pyrosequencing is a versatile technique that facilitates microbial genome sequencing that can be used to identify bacterial species, discriminate ...

Determination of Fatty Acid Oxidation and Lipogenesis in Mouse Primary Hepatocytes

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid oxidation, or alternatively, synthesize new lipid via de novo lipogenesis. The net sum of these pathways is dictated by a number of factors, which in certain disease states leads to fatty liver disease. Excess hepatic lipid accumulation is associated with whole body insulin resistance and coronary heart disease. Tools to study lipid metabolism in hepatocytes are useful to understand the role of hepatic lipid metabolism in certain metabolic disorders.
In the liver, hepatocytes regulate the breakdown and synthesis of fatty acids via &#946;-fatty oxidation and de novo lipogenesis, respectively. Quantifying metabolism in these pathways provides insight into hepatic lipid handling. Unlike in vitro quantification, using primary hepatocytes, making measurements in vivo is technically challenging and resource intensive. Hence, quantifying &#946;-fatty acid oxidation and de novo lipogenesis in cultured mouse hepatocytes provides a straight forward method to assess hepatocyte lipid handling. 
Here we describe a method for the isolation of primary mouse hepatocytes, and we demonstrate quantification of &#946;-fatty acid oxidation and de novo lipogenesis, using radiolabeled substrates.

De novo lipogenesis and &#946;-fatty acid oxidation constitute key metabolic pathways in hepatocyte, pathways that are perturbed in several metabolic disorders, including fatty liver disease. Here we demonstrate isolation of mouse primary hepatocytes and describe quantification of &#946;-fatty acid oxidation and lipogenesis.

Lipid metabolism in liver is complex. In addition to importing and exporting lipid via lipoproteins, hepatocytes can oxidize lipid via fatty acid ...

Isolation of Specific Genomic Regions and Identification of Associated Molecules by enChIP

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. To enable the non-biased identification of molecules interacting with a specific genomic region of interest, we recently developed the engineered DNA-binding molecule-mediated chromatin immunoprecipitation (enChIP) technique. Here, we describe how to use enChIP to isolate specific genomic regions and identify the associated proteins and RNAs. First, a genomic region of interest is tagged with a transcription activator-like (TAL) protein or a clustered regularly interspaced short palindromic repeats (CRISPR) complex consisting of a catalytically inactive form of Cas9 and a guide RNA. Subsequently, the chromatin is crosslinked and fragmented by sonication. The tagged locus is then immunoprecipitated and the crosslinking is reversed. Finally, the proteins or RNAs that are associated with the isolated chromatin are subjected to mass spectrometric or RNA sequencing analyses, respectively. This approach allows the successful identification of proteins and RNAs associated with a genomic region of interest.

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the functions of these regions. This protocol describes procedures to perform engineered DNA-binding molecule-mediated chromatin imunoprecipitation (enChIP) for identification of proteins and RNAs associated with a specific genomic region.

The identification of molecules associated with specific genomic regions of interest is required to understand the mechanisms of regulation of the ...

Methods for the Determination of Rates of Glucose and Fatty Acid Oxidation in the Isolated Working Rat Heart

The mammalian heart is a major consumer of ATP and requires a constant supply of energy substrates for contraction. Not surprisingly, alterations of myocardial metabolism have been linked to the development of contractile dysfunction and heart failure. Therefore, unraveling the link between metabolism and contraction should shed light on some of the mechanisms governing cardiac adaptation or maladaptation in disease states. The isolated working rat heart preparation can be used to follow, simultaneously and in real time, cardiac contractile function and flux of energy providing substrates into oxidative metabolic pathways. The present protocol aims to provide a detailed description of the methods used in the preparation and utilization of buffers for the quantitative measurement of the rates of oxidation for glucose and fatty acids, the main energy providing substrates of the heart. The methods used for sample analysis and data interpretation are also discussed. In brief, the technique is based on the supply of 14C- radiolabeled glucose and a 3H- radiolabeled long-chain fatty acid to an ex vivo beating heart via normothermic crystalloid perfusion. 14CO2 and 3H2O, end byproducts of the enzymatic reactions involved in the utilization of these energy providing substrates, are then quantitatively recovered from the coronary effluent. With knowledge of the specific activity of the radiolabeled substrates used, it is then possible to individually quantitate the flux of glucose and fatty acid in the oxidation pathways. Contractile function of the isolated heart can be determined in parallel with the appropriate recording equipment and directly correlated to metabolic flux values. The technique is extremely useful to study the metabolism/contraction relationship in response to various stress conditions such as alterations in pre and after load and ischemia, a drug or a circulating factor, or following the alteration in the expression of a gene product.

The following protocol describes the preparation and utilization of buffers for the quantitative measurement of rates of glucose and fatty acid oxidation in the isolated working rat heart. The methods used for sample analysis and data interpretation are also discussed.

The mammalian heart is a major consumer of ATP and requires a constant supply of energy substrates for contraction. Not surprisingly, alterations of ...

A Plate Competition Assay As a Quick Preliminary Assessment of Disease Suppression

The goal was to develop and optimize a simple, affordable, and effective bioassay to detect disease suppressive ability of a specific compost against soilborne fungus Rhizoctonia solani. R. solani is a pathogen of a wide range of plant hosts worldwide. The fungus survives in soils as a saprophyte and grows rapidly on simple water agar media. The plate assay is a rapid method to compare composts for their ability to slow the growth of R. solani. The assay also correlates well with suppression of other soilborne fungal pathogens that survive as saprophytes in soils such as Alternaria early blights, Fusarium wilt, Phytophthora root rot, and Pythium root rot.

Presented is a protocol for a plate competition assay to identify whether a specific compost is likely to contain bacteria and fungi that suppress growth of Rhizoctonia solani.

The goal was to develop and optimize a simple, affordable, and effective bioassay to detect disease suppressive ability of a specific compost against ...

Laser-Capture Microdissection RNA-Sequencing for Spatial and Temporal Tissue-Specific Gene Expression Analysis in Plants

The development of a complex multicellular organism is governed by distinct cell types that have different transcriptional profiles. To identify transcriptional regulatory networks that govern developmental processes it is necessary to measure the spatial and temporal gene expression profiles of these individual cell types. Therefore, insight into the spatio-temporal control of gene expression is essential to gain understanding of how biological and developmental processes are regulated. Here, we describe a laser-capture microdissection (LCM) method to isolate small number of cells from three barley embryo organs over a time-course during germination followed by transcript profiling. The method consists of tissue fixation, tissue processing, paraffin embedding, sectioning, LCM and RNA extraction followed by real-time PCR or RNA-seq. This method has enabled us to obtain spatial and temporal profiles of seed organ transcriptomes from varying numbers of cells (tens to hundreds), providing much greater tissue-specificity than typical bulk-tissue analyses. From these data we were able to define and compare transcriptional regulatory networks as well as predict candidate regulatory transcription factors for individual tissues. The method should be applicable to other plant tissues with minimal optimization.

Presented here is a protocol for laser-capture microdissection (LCM) of plant tissues. LCM is a microscopic technique for isolating areas of tissue in a contamination-free manner. The procedure includes tissue fixation, paraffin embedding, sectioning, LCM and RNA extraction. RNA is used in the downstream tissue-specific, temporally resolved analysis of transcriptomes.