We are grateful to Dr. Neil C. Gudmestad at North Dakota State University for providing S. subterranea plasmid DNA, Dr. Hanu Pappu at Washington State University (WSU) for providing PMTV RNA, and Dr. Debra Inglis at WSU for providing R. solani AG3. Special thanks to Dr. Jeremy Jewell at WSU for providing comments on the manuscript and WSU CAHNRS Communications for videography. This research was supported by the Northwest Potato Research Consortium and the Washington State Department of Agriculture - Specialty Crop Block Grant Program (grant no. K1764). PPNS No. 0741, Department of Plant Pathology, College of Agriculture, Human and Natural Resource Sciences, Agricultural Research Center, Hatch Project No. WNP00833, Washington State University, Pullman, WA, USA.

1. On-Site Molecular Detection of Pathogens using a Portable Real-Time PCR System
Note: See Figure 1.
<ol>
	<li>Magnetic bead-based DNA extraction 
	Note: A magnetic bead-based DNA extraction kit (e.g. from Primerdesign) was used according to the manufacturer&#39;s instructions. All reagents should be stored at room temperature (18-25 &#176;C). Once the lyophilized Proteinase K (Bottle No.1) is suspended (using Bottle No.1a), store at -20 &#176;C.
	<ol>
		<li>Mix 20-50 mg of soil sample with 500 &#181;L of Sample Prep Solution in a microtube. 
		Note: The ratio of soil:Prep Solution is important as mixing them in other ratios may cause a failure of downstream experiments (e.g., contamination by inhibitors of PCR).</li>
		<li>Grind the soil on the bottom of the tube using a small sterile pestle until the solution is cloudy. Further suspend soil particles in the solution by shaking the microtube and let it stand, undisturbed, to let the soil particles settle completely (typically between 5 to 10 min).</li>
		<li>Transfer 200 &#181;L of supernatant into a fresh microtube and add 200 &#181;L of Lysis Buffer (Bottle No. 2: Guanidine Hydrochloride solution) and 20 &#181;L of Proteinase K (Bottle No.1).</li>
		<li>Mix the lysate thoroughly by inverting the tube and incubate at ambient temperature for 15 min. 
		Note: If the lysate is found on the microtube lid, tap the tube or use a centrifuge, if available to remove from the lid.</li>
		<li>Add 500 &#181;L of the binding buffer/magnetic bead mix (Bottle No.3) to the lysed sample. Mix well by pipetting up and down and incubate at ambient temperature for 5 min away from the magnetic tube rack. 
		Note: Make sure to mix the bead solution well before use to ensure that the beads are aliquoted evenly from the storage bottle.</li>
		<li>Place the microtube on the magnetic tube rack. Wait at least 2 min or until all the beads in the microtube attach to the magnetic-side wall. Then, remove and discard all of the supernatant by pipetting. 
		Note: Do not disturb the magnetized beads while removing and aspirating the supernatant. DNA has now been captured by the magnetic beads.</li>
		<li>Remove the microtube from the magnetic tube rack, add 500 &#181;L of Wash Buffer-1 (Bottle No. 4: sodium perchlorate/ethanol solution) and re-suspend the beads by repeated pipetting until the beads are uniformly dispersed. Perform this washing step to remove protein and salt from the sample. Let the mixture sit for 30 s.</li>
		<li>Repeat step 1.1.6.</li>
		<li>Remove the microtube from the magnetic tube rack, add 500 &#181;L of Wash Buffer-2 (Bottle No. 5: sodium perchlorate/ethanol solution) and re-suspend the beads by repeated pipetting until the beads are uniformly dispersed. Let the mixture sit for 30 s.</li>
		<li>Repeat step 1.1.6.</li>
		<li>Remove the microtube from the magnetic tube rack and then add 500 &#181;L of 80% ethanol (Bottle No.6). 
		Note: This step is necessary for the removal of residual salts from the sample.
		<ol>
			<li>Re-suspend the beads by repeated pipetting until the beads are uniformly dispersed. Let this stand for 10 min with occasional mixing by inversion.</li>
		</ol>
		</li>
		<li>Repeat step 1.1.6.</li>
		<li>Air dry the magnetic bead pellet for 10 min at ambient temperature with the microtube lid open. 
		Note: The beads should be free from any visible residual ethanol but not completely dried out.</li>
		<li>Remove the microtube from the magnetic tube rack, add 50-200 &#181;L of Elution Buffer (Bottle No.7) and re-suspend the beads by repeated pipetting until the beads are uniformly dispersed and let it stand for 30 s. 
		Note: In the above steps, the purified DNA is released from the magnetic beads into the elution buffer.</li>
		<li>Place the microtube on the magnetic tube rack. Wait at least 2 min or until all the beads in the microtube attach to the magnetic-side wall.</li>
		<li>Transfer the supernatant that now contains the purified DNA/RNA to a 0.5 mL microtube for use in the downstream steps.</li>
	</ol>
	</li>
	<li>Portable real-time PCR 
	Note: A portable thermocycler and the PCR assay kit were used according to the manufacturer&#39;s instructions (see the Table of Materials).
	<ol>
		<li>Open and run the thermocycler-associated software, select Target detection test and input all the description information into the Name &#38; Details, Notes, Samples, and Tests data entry fields. 
		Note: Wells #1 and #2 are designated by the software for the negative control and positive control, respectively.</li>
		<li>Prepare PCR reagents prior to use. Transfer 500 &#181;L of the master mix re-suspension buffer into the tube containing lyophilized master mix and mix well by inversion. Transfer the entire master mix into the brown microtube labeled primers/probe (Table 2).
		<ol>
			<li>Cap and shake the microtube to mix. Thorough mixing is required to ensure that all components are re-suspended completely. Let this mixture sit for 5 min before use. 
			Note: Store the reaction mix at -20 &#176;C after use.</li>
		</ol>
		</li>
		<li>Prepare a negative control by transferring 10 &#181;L of the prepared reaction mix from the previous step into a 0.2 mL PCR tube and then add 10 &#181;L of sterile nuclease-free deionized water.</li>
		<li>Prepare a positive control by transferring 10 &#181;L of the prepared reaction mix from the step 1.2.2 into a 0.2 mL PCR tube and then add 10 &#181;L of positive control template.</li>
		<li>For each sample, transfer 10 &#181;L of the prepared reaction mix from the previous step into a 0.2 mL PCR tube, and then add 10 &#181;L of sample DNA prepared from step 1.1.16.</li>
		<li>Load the wells of the thermocycler with the contents from their respective PCR tubes as described in step 2.1.1.</li>
		<li>Once all the necessary information has been entered and confirmed, select Start Run and choose either the ethernet-connected instrument or a USB drive. 
		Note: If the USB drive option is selected, the run file must be saved on the drive to be used with the thermocycler (e.g., F:\genesig). The run will begin immediately after the drive is inserted into the thermocycler.</li>
	</ol>
	</li>
	<li>Data analysis of portable real-time PCR.
	<ol>
		<li>Once the run has finished, open the run file (.usb) from the USB drive using the thermocycler-associated software or directly view the run results in the software by clicking Results.</li>
		<li>Before analyzing results, save the completed run to avoid losing data.</li>
		<li>In the Results tab, view the status of the run, categorized by samples. 
		Note: Data that can be obtained in this tab are the status of results and copy number detected in the sample.</li>
		<li>Click on the Details tab to view the amplification curves. When the target is successfully detected, Cq (quantification cycle) values of both the target and internal control are displayed. 
		Note: These values are calculated in the final report and are used to determine whether a sample is positive for the target and if there are problems with the reaction or the DNA samples.</li>
	</ol>
	</li>
</ol>
2. Other protocols
<ol>
	<li>Alternative lab-based DNA extraction methods
	<ol>
		<li>CTAB-phenol-chloroform based methods
		<ol>
			<li>Perform CTAB-phenol-chloroform based methods, following the Doyle method18 and the Dellaporta method19 as described previously.</li>
		</ol>
		</li>
		<li>DNA mini-preparation method 
		Note: The Edwards method20 was performed as follows.
		<ol>
			<li>Add 500 mg of soil, followed by five 1.4 mm ceramic beads and 750 &#181;L of Edwards buffer (200 mM Tris, pH 8.0, 200 mM NaCl, 25 mM EDTA, 0.5% SDS) to a microtube and mix well.</li>
			<li>Incubate the microtube at 65 &#176;C for 5 min.</li>
			<li>Homogenize the sample with a bead beater homogenizer for 60 s (or by using a mortar and pestle).</li>
			<li>Centrifuge the sample at 14,000 x g for 5 min.</li>
			<li>Transfer 500 &#181;L of supernatant to a fresh microtube and then mix with 500 &#181;L of chilled isopropanol. Mix by inverting the tube 10 times.</li>
			<li>Centrifuge the sample at 14,000 x g for 15 min to pelletize the DNA.</li>
			<li>Decant the supernatant and let the DNA pellet air dry at room temperature until the remaining ethanol has evaporated.</li>
			<li>Wash the DNA pellet with 750 &#181;L of chilled 70% ethanol.</li>
			<li>Air dry the pellet before re-suspending in 50-100 &#181;L of TE buffer (10 mM Tris-HCl, pH 8.0 and 1 mM EDTA).</li>
		</ol>
		</li>
		<li>Other Alternative methods
		<ol>
			<li>Perform Silica-base DNA extraction using kit #1&#160;(MP BIO&#160;Fast DNA Spin) and kit #2 (Zymo BIOMICS DNA Miniprep Kit) according to the manufacturers&#39;&#160;instructions.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Conventional lab-based real-time PCR. 
	Note: A conventional thermocycler was used with mastermix for probe-based PCR, various primers and oligonucleotide probes (Table 2).
	<ol>
		<li>Using non-transparent bottomed PCR tubes or a PCR plate, prepare 20 &#181;L reactions for all DNA samples to be analyzed, as well as a negative control (nuclease-free deionized water) and a positive template control prepared in-house.</li>
		<li>For each PCR tube or well, prepare a mixture including 10 &#181;L of the mastermix, 7 &#181;L of nuclease-free deionized water, 2 &#181;L of 2 &#181;M primers/probe, and 1 &#181;L of DNA sample (from step 1.1.16, or 2.1) or control template, per sample.</li>
		<li>Close the PCR tubes or plate and begin the reaction by selecting the appropriate PCR program.</li>
	</ol>
	</li>
	<li>Data analysis of conventional lab-based real-time PCR
	<ol>
		<li>Use thermocycler-associated software to analyze the results from the conventional thermocycler. To begin data analysis, transfer the run file from the thermocycler to a USB drive, insert a USB drive and select Export.</li>
		<li>Open the data file (.pcrd) from the exported run in the thermocycler-associated software.</li>
		<li>Highlight the well of a sample by locating its corresponding well on the instrument. Amplification curves and standard curves (if applicable) are automatically generated. If the sample information is not entered, click Plate Setup or a similar function to begin data input before analyzing data.</li>
		<li>View the data on the quantification tab; this can be exported for data analysis with a third-party software such as CSV, XML, or HTML file readers.</li>
		<li>Obtain Cq data based on the determined thresholds and compare it with the positive and negative controls. 
		Note: If the DNA standards of the target were used in the assay, compare the sample Cq data to that of the standards to determine the Cq cut-off</li>
	</ol>
	</li>
</ol>

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W., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+new+large-scale+soil+DNA+extraction+procedure+and+real-time+PCR+assay+for+the+detection+of+Sclerotium+cepivorum+in+soil.">A new large-scale soil DNA extraction procedure and real-time PCR assay for the detection of Sclerotium cepivorum in soil.</a> Eur J Plant Pathol. 134 (3), 467-473 (2012).</li><li>Miles, T. D., Martin, F. N., Coffey, M. D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Development+of+Rapid+Isothermal+Amplification+Assays+for+Detection+of+Phytophthora+spp+in+Plant+Tissue.">Development of Rapid Isothermal Amplification Assays for Detection of Phytophthora spp in Plant Tissue.</a> Phytopathology. 105 (2), 265-278 (2014).</li><li>Notomi, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Loop-mediated+isothermal+amplification+of+DNA.">Loop-mediated isothermal amplification of DNA.</a> Nucleic Acids Res. 28 (12), e63-e63 (2000).</li><li>Narayanasamy, P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> Microbial Plant Pathogens: Detection and Management in Seeds and Propagules. , (2016).</li><li>Braun-Kiewnick, A., Altenbach, D., Oberh&#228;nsli, T., Bitterlin, W., Duffy, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+rapid+lateral-flow+immunoassay+for+phytosanitary+detection+of+Erwinia+amylovora+and+on-site+fire+blight+diagnosis.">A rapid lateral-flow immunoassay for phytosanitary detection of Erwinia amylovora and on-site fire blight diagnosis.</a> Journal of Microbiological Methods. 87 (1), 1-9 (2011).</li></ol>

The authors declare no competing financial interests.

As shown in Table 1, recent technological advances in the molecular identification of pathogenic agents have increased the efficacy, accuracy, and speed of diagnosis, which have contributed to the detection of pre-symptomatic infections27. Regarding on-site diagnosis, LAMP and lateral-flow methods are frequently used because they are portable and provide immediate results at a lower cost. However, in the case of serological methods, species-specific detection can be hard to achieve. This occasionally causes misdetection of off-target microbes such as common soil inhabitants. For example, there can be cross reactivity between the serological tests of Phytophthora spp. and Pythium spp. in the case of potato pathogens28, indicating that there are sometimes difficulties detecting the targeted plant pathogens.
In the present study, we have developed an optimized protocol for on-site molecular detection of soil-borne potato pathogens using the portable real-time PCR system by comparing its capabilities with that of a conventional lab-based real-time PCR system. We found that the on-site method specifically detects the potato pathogens in the soil sample, although sensitivity is ~10 times lower than that of an equivalent lab-based assay. It is also worth considering that in this case both the laboratory and field test did not use a biologically relevant sample size. Large sample sizes are required for use in routinely screening field soils as previously described29,30,&#160;where sample sizes of between 250 g to 1 kg are processed, although these methods require skilled operators and sophisticated equipment to extract DNA. Typically, a large-scale soil DNA extract is taken from a single aggregate soil sample representative of numerous subsamples over 1 to 4 hectares6,29,30. However, the protocol developed here is quick, easy-to-use for users with no prior experience in molecular diagnostics and can be used outside of a lab. As the method is rapid and relatively cheap compared to large-scale soil extraction, it could be used to screen many small-scale samples taken from a similar sampling area to large-scale aggregate samples. This could overcome some of the deficiencies of a small sample size and determine additional information on the spatial distribution of the pathogen in the field. In addition, the portability and speed of the method means that it can also be used in demonstration activities to growers for educational and engagement purposes.
Another consideration is that many real-time PCR assays are already published for a wide range of plant pathogens5. This system can make use of these existing assays without the need to design new LAMP primers to enable in field testing. A frequent criticism of LAMP assays is that they can be difficult to design31. Portable PCR, therefore, allows the relatively easy implementation of a wide range of readily available pathogen tests for on-site testing.
Traditional methods can be often costly, laborious, inaccurate, and time-consuming. The simplicity of the on-site method we developed allows growers and industry workers to perform pathogen detection by themselves and perhaps generate a result much quicker than sending to a diagnostic laboratory that could be some distance away. The promptness and sensitivity of the portable PCR method can help growers avoid potential secondary infections, which can further increase of the pathogen population and inadvertent spread (via equipment or humans). In conclusion, the on-site method developed in the present study enables accurate and relatively sensitive detection of important soil-borne pathogens in the field. Our hope is that the on-site method developed in this study will contribute in a current diagnostic pipeline (Figure 6), not only by providing quick and accurate answers to epidemiological questions about plant diseases in the field but also by providing increased understanding of the biology and epidemiology of plant pathogens.

Accurate and rapid identification of causative pathogens significantly impacts decisions regarding plant disease management. Soil-borne diseases are particularly difficult to diagnose because the soil environment is extremely large relative to plant mass, and complex, making it a challenge to understand all the aspects of soil-borne diseases. Moreover, soil-borne diseases can be symptomless during early infection stages, dependent on environmental stressors, and some have long latent periods that result in delayed diagnoses1. Many soil-borne pathogens have developed survival structures, such as specialized spores or melanized hyphae, which can survive in the soil for many years even in the absence of their hosts. Utilized approaches for soil-borne disease management include: avoiding known infested fields, using pathogen-free certified seeds and seedlings, keeping equipment sanitary, and restricting the movement of soil and water when possible. Knowledge of the pathogen presence through molecular detection strategies can also play a useful role by informing timely decisions regarding early-stage treatments or pre-plant assessments of the fields. On-site testing provides additional advantages of providing a rapid result without sending the sample to a diagnostic laboratory that maybe some distance away and also can engage the grower if such a diagnostic is performed &#39;field-side&#39; in their presence.
For on-site diagnosis based on molecular detection, sensitivity, specificity, robustness (repeatability and reproducibility), and efficiency (i.e., simplicity and cost performance) are crucial factors for consideration. Lateral flow devices (LFDs) such as the Immunostrip and PocketDiagnostic, are popular methods for on-site pathogen detection because of their simplicity as a one-step assay. However, LFDs may not be the right diagnostic tool in all situations because they lack the sensitivity and specificity, and occasionally provides ambiguous results if the target pathogen is in low concentrations and can cross-react with similar species or genera2. Loop-mediated isothermal amplification (LAMP) is also applicable for on-site pathogen detection and is particularly inexpensive due to low-cost reagents, reaction conditions that remain constant, and simple colorimetric visual analysis. However, both LFDs and LAMP are typically used qualitatively, although both approaches can be used quantitatively with more expensive equipment3. The polymerase chain reaction (PCR) offers high specificity, high sensitivity, and a quantitative capability in comparison to the aforementioned methods of detection. However, the conventional lab-based PCR technology requires expensive laboratory equipment and skilled personnel, which is a major disadvantage in adopting this technology as a detection method for on-site purposes.
In this protocol, an on-site diagnostic method using a portable real-time PCR instrument is demonstrated. Real-time PCR technology offers advantages over other methods in terms of quantitative accuracy, sensitivity, and versatility, and has been widely used for the detection of a broad range of plant pathogens4,5, including various potato pathogens in soil6. Because of the recent trends of the fast-growing, competitive market, equipment required for PCR technology has continued to develop to be more compact and less expensive7. The protocol is composed of the following steps: magnetic bead-based nucleic acid extraction, portable real-time PCR (fluorogenic probe-based assay), and quantitative data analysis that can be all done remotely using a laptop computer (Figure 1).
Using the portable PCR protocol developed here, soil samples were analyzed to detect the soil-borne pathogen, Spongospora subterranea. Spongospora was chosen as it is an important potato pathogen as the causal agent of powdery scab8. In recent decades, the presence of this disease is considered to have spread to many regions where potatoes are grown9,10.&#160;Powdery scab, through the presence of pimple like lesions on the tubers can cause considerable qualitative yield losses to potato growers. In addition, S.&#160;subterranea can vector Potato Mop Top Virus (PMTV), which can cause internal lesion symptoms in tubers (known as spraing)11,12. Therefore, it is important to know if S. subterranea is present in fields prior to planting6. We also demonstrate the usefulness of this protocol for the detection of Rhizoctonia solani Anastomosis Group 3 (AG3) and PMTV. Although several anastomosis groups of Rhizoctonia solani cause diseases in potatoes, AG3 is arguably the most important worldwide13, causing stem canker and black scurf resulting in marketable yield losses of up to 30%14. PMTV causes necrotic lesions within the tubers, which are commonly called spraing. This virus has recently been reported for the first time in several states in the Pacific Northwest15,16,17,&#160;and is of increasing concern to growers in this important potato growing region. In addition to determining the effectiveness of portable PCR for these important diseases, optimum DNA extraction methodology and soil sample size were also investigated in this study.
The results suggest that the portable PCR method is versatile and applicable for the detection of different pathogens. The on-site detection method we developed can allow the frontline workers in agriculture (e.g., growers) to make earlier decisions regarding disease management, such as variety selections or rotations, and can quantify a plant pathogen in the sample during a field survey, prior to planting, to avoid potential disease outbreaks.

<table border="0" cellpadding="0" cellspacing="0" width="1073">
	<tbody>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">White shell PCR plate</td>
			<td style="width:183px;">Bio-Rad</td>
			<td style="width:195px;">HSP9601</td>
			<td style="width:247px;"></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">CFX Manager</td>
			<td style="width:183px;">Bio-Rad</td>
			<td style="width:195px;">1845000</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">SsoAdvanced Universal Probes Supermix&#160;</td>
			<td style="width:183px;">Bio-Rad</td>
			<td style="width:195px;">1725280</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">CFX96 Touch qPCR instrument</td>
			<td style="width:183px;">Bio-Rad&#160;</td>
			<td style="width:195px;">1855195</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Ethylalcohol, pure 200 proof</td>
			<td style="width:183px;">Decon Laboratories</td>
			<td style="width:195px;">04-355-222</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Masterclear cap strips &#38; real&#8209;time PCR tube strips</td>
			<td style="width:183px;">Eppendorf</td>
			<td>9510222109</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Ethylenediaminetetraacetic Acid (EDTA)</td>
			<td style="width:183px;">Fisher BioReagents</td>
			<td style="width:195px;">BP118-500</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Phenol, Saturated&#160;</td>
			<td style="width:183px;">Fisher BioReagents</td>
			<td style="width:195px;">BP1750I-400</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Tris(hydroxymethyl)aminomethane (Tris)</td>
			<td style="width:183px;">Fisher BioReagents</td>
			<td style="width:195px;">BP152-5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">q16 real-time PCR machine</td>
			<td style="width:183px;">genesig</td>
			<td style="width:195px;">MBS486001</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Ultrapure Sodium Dodecyl Sulfate (SDS)</td>
			<td style="width:183px;">Invitrogen</td>
			<td style="width:195px;">15525-017</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">2-Propanol (Isopropanol)</td>
			<td style="width:183px;">JT Baker</td>
			<td style="width:195px;">67-63-0</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Chloroform</td>
			<td style="width:183px;">JT Baker</td>
			<td style="width:195px;">9180-03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Hydrochloric Acid, 36.5-38.0% (HCl)</td>
			<td style="width:183px;">JT Baker</td>
			<td style="width:195px;">9535-03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">iso-Amyl Alcohol</td>
			<td style="width:183px;">JT Baker</td>
			<td style="width:195px;">9038-01</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">FastDNA Spin kit&#160;</td>
			<td style="width:183px;">MP Biomedicals</td>
			<td style="width:195px;">6560-200</td>
			<td>Silica-based DNA extraction kit #1</td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Luna Universal One-Step RT-qPCR Kit</td>
			<td style="width:183px;">New England Biolabs</td>
			<td style="width:195px;">E3005S</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">0.1ml Low Profile Individual PCR Tubes</td>
			<td style="width:183px;">Phenix Research</td>
			<td style="width:195px;">MPC-100LP</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Easy DNA/RNA Extraction Kit&#160;</td>
			<td style="width:183px;">Primerdesign</td>
			<td style="width:195px;">genesigEASY-EK</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">genesig Easy 1.5 mL tubes</td>
			<td style="width:183px;">Primerdesign</td>
			<td style="width:195px;">genesigEASY-1.5</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Magnetic tube rack</td>
			<td style="width:183px;">Primerdesign</td>
			<td style="width:195px;">genesigEASY-MR</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Spongospora subterranea f. sp. subterranea genesig Easy Kit</td>
			<td style="width:183px;">Primerdesign</td>
			<td style="width:195px;">Path-S.subterranea-EASY</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Hexadecyltrimethylammonium bromide (CTAB)</td>
			<td style="width:183px;">Sigma-Aldrich</td>
			<td style="width:195px;">H6269-100G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Pellet pestles</td>
			<td style="width:183px;">Thermo Fisher Scientific</td>
			<td style="width:195px;">12-141-364</td>
			<td></td>
		</tr>
		<tr height="21">
			<td height="21" style="height:21px;width:451px;">Sodium chloride (NaCl)</td>
			<td style="width:183px;">Thermo Fisher Scientific</td>
			<td style="width:195px;">7647-14-5</td>
			<td></td>
		</tr>
		<tr height="22">
			<td height="22" style="height:22px;width:451px;">DNA Miniprep kit&#160;</td>
			<td style="width:183px;">ZymoBIOMICS</td>
			<td style="width:195px;">D4300</td>
			<td>Silica-based DNA extraction kit #2</td>
		</tr>
	</tbody>
</table>

on-site molecular detection of soil-borne phytopathogens using a portable real-time pcr system

On-site diagnosis of plant diseases can be a useful tool for growers for timely decisions enabling the earlier implementation of disease management strategies that reduce the impact of the disease. Presently in many diagnostic laboratories, the polymerase chain reaction (PCR), particularly real-time PCR, is considered the most sensitive and accurate method for plant pathogen detection. However, laboratory-based PCRs typically require expensive laboratory equipment and skilled personnel. In this study, soil-borne pathogens of potato are used to demonstrate the potential for on-site molecular detection. This was achieved using a rapid and simple protocol comprising of magnetic bead-based nucleic acid extraction, portable real-time PCR (fluorogenic probe-based assay). The portable real-time PCR approach compared favorably with a laboratory-based system, detecting as few as 100 copies of DNA from Spongospora subterranea. The portable real-time PCR method developed here can serve as an alternative to laboratory-based approaches and a useful on-site tool for pathogen diagnosis.

Comparison of DNA extraction methods
The compatibility of a magnetic bead-based DNA extraction method with real-time PCR was evaluated by detecting the amounts of S. subterranea DNA in a soil sample from fields infested with the pathogen. As shown in Supplemental Figure 1, the magnetic bead-based method was compared with the other methods including a CTAB-phenol-chloroform based method18, quick DNA mini-preparation methods19,20, and other standard silica-based DNA extraction kits. DNA samples extracted using the six different methods were subjected to conventional lab-based real-time PCR. The results suggested that the magnetic bead-based method is comparable with the other methods, although silica-based DNA extraction kit showed the best performance among the methods we tested. All kits contain guanidinium thiocyanate or guanidinium hydrochloride: both are powerful chaotropic agents, which denature most of cellular proteins including RNases and DNases. Therefore, using the methods is suitable for both DNA and RNA extractions.
Comparison between a portable real-time PCR and a conventional lab-based real-time PCR
To compare the sensitivity and specificity of a portable PCR to a conventional lab-based PCR, absolute quantification of the pathogen DNA was performed using different amounts of the S. subterranea ITS gene, which was carried by the pGEM-T vector21. A series of 10-fold dilutions of the ITS gene (106 to 100 copies) were analyzed using the SsTQ primers/probe set22. The results demonstrated that the portable PCR method detected the target pathogen DNA (~100 copies), although the sensitivity was 10 times lower than that of the conventional lab-based PCR method, which detected at least 10 copies (Figure 2).
For further validation, artificially infested soils were tested. S. subterranea sporosori were obtained from powdery scab root galls from potato roots. The soils were infested with sporosori suspensions at a final concentration of 105 sporosori/g dry weight of soil. Using the magnetic bead-based method, DNA was extracted from the infested soil samples, and 10-fold serial dilutions were prepared to obtain concentrations equivalent to 105, 104, 103, 102, 101, and 100 sporosori/g dry weight of soil. The DNA samples were used for PCR using the SPO primer/probe set23. The results showed that the portable PCR method has comparable analytical capability to a conventional lab-based PCR method but, again, the sensitivity was reduced by a factor of ~10 (Figure 3).
Finally, we tested a soil sample from a field that was naturally contaminated with S. subterranea. The magnetic bead-based DNA extraction was performed on different amounts of soils (10, 20, 50 and 100 mg of soil per 500 &#181;L of extraction buffer solution). The results suggested that the optimal weight of soil as a starting material for the DNA extraction was 50-100 mg (Figure 4). Soil amounts outside the range caused a failure of the downstream PCR steps. This effect might be because when excess amounts of soil are used as starting material, contaminations (e.g., phenolic compounds) can interfere with the PCR24. In the case of lower volumes of soil, the amount of extracted DNA may be lower than the detection limit of PCR (e.g., the yield of total DNA extracted from 10-20 mg soil was varied). Sensitivity was quite comparable between the portable PCR and conventional PCR methods. Similar results were obtained in DNA samples by different extraction methods (Supplemental Figure 2)
Detection of other pathogens by the on-site detection system using a portable real-time PCR
We tested the portable PCR method to detect other important soil-borne potato pathogens, R. solani AG3 and PMTV. In this study, we performed real-time PCR using the RsTq primers/RQP1 probe set25 for R. solani AG3 detection with DNA from pure culture. We also performed real-time PCR using the PMTV-D primer/probe set26 for PMTV detection with RNA from a spraing symptomatic tuber sample was used. As shown in Figure 5, the portable PCR method successfully detected both pathogens. The results were comparable between the portable and conventional instruments, suggesting that the portable PCR method is versatile and applicable to other pathogen detections if the primer sequences designed for real-time PCR are available.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/56891/56891fig1.jpg" /> 
Figure 1. Procedure of a portable real-time PCR system for on-site pathogen detection. The protocol is composed of steps in the following order: lysate preparation by brief homogenization (A), magnetic bead-based nucleic acid extraction (B), portable real-time PCR (C), and quantitative data analysis using a laptop computer (D). Note that all steps can be completed on site.
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/56891/56891fig2v2.jpg" /> 
Figure 2. Comparison of sensitivity between a portable PCR and a conventional lab-based PCR. Quantification of the pathogen DNA was performed using different amounts of the S. subterranea ITS gene (106 to 100 copies) with the SsTQ primers/probe set. Linear regression between log value of S. subterranea plasmid DNA and reciprocal Log value of Cq on the conventional thermocycler (A) and the portable thermocycler (B). <a href="/files/ftp_upload/56891/56891fig2v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/56891/56891fig3v2.jpg" /> 
Figure 3. Comparison of detection performance in artificially infested soils with S. subterranea. The soils were artificially infested (105 to 100 sporosori/g dry weight of soil) with S. subterranea sporosori suspensions. Using the magnetic bead-based method, DNA was extracted from the infested soil samples. PCRs were performed using the soil samples with the SPO primer/probe set. Linear regression between log value of the starting quantity in sporosori per gram of soil and the reciprocal log value of Cq on the conventional thermocycler (A) and the portable thermocycler (B). <a href="/files/ftp_upload/56891/56891fig3v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/56891/56891fig4v2.jpg" /> 
Figure 4. Comparison of starting amount of soil samples for DNA extraction. The magnetic bead-based method was used for DNA extraction from 10, 20, 50, and 100 mg of soil samples. Real-time PCRs were performed using the portable thermocycler. Standard curves represent the relationship between the amount of total DNA extracted from the soil samples (x-axis) and the amounts of PCR product (y-axis) amplified by the Sss primer/probe set. <a href="/files/ftp_upload/56891/56891fig4v2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/56891/56891fig5.jpg" /> 
Figure 5. Detection of other potato pathogens, R. solani and PMTV. Real-time PCRs were performed using the portable thermocycler and the conventional thermocycler. R. solani AG3 was detected in total DNA extracted from pure culture using RsTq primers and the RQP1 probe (A) PMTV was detected in total RNA extracted from a PMTV-infected tuber sample using the PMTV-D primer/probe set (B). <a href="/files/ftp_upload/56891/56891fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/56891/56891fig6.jpg" /> 
Figure 6. A diagnostic pipeline for phytopathogens. Flowchart shows a general workflow for phytopathogen diagnosis. Note that the traditional step, e.g., visual identification, can be omitted if on-site molecular detection is utilized, which makes the entire process of diagnosis simple and fast. <a href="/files/ftp_upload/56891/56891fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td></td>
			<td>Portable real-time PCR</td>
			<td>Real-time PCR</td>
			<td>LAMP</td>
			<td>ELISA</td>
			<td>Lateral-flow</td>
		</tr>
		<tr>
			<td>Cost per target reaction</td>
			<td>$0.60-$8.47</td>
			<td>$0.60</td>
			<td>$0.75</td>
			<td>$0.60</td>
			<td>$4.74</td>
		</tr>
		<tr>
			<td>Sensitivity</td>
			<td>100 copies</td>
			<td>10 copies</td>
			<td>10 copies</td>
			<td>1-10 sporosori33 
			1-10 ng/mL (protein)33</td>
			<td>1-10 sporosori34 
			5x105CFU/mL35</td>
		</tr>
		<tr>
			<td>Time Expense</td>
			<td>90 minutes</td>
			<td>80-240 minutes</td>
			<td>50-90 minutes32</td>
			<td>3-24 hours</td>
			<td>10-15 minutes</td>
		</tr>
		<tr>
			<td>Preparation Required</td>
			<td>&#9679;Nucleic acid extraction 
			&#9679; Primer design</td>
			<td>&#9679;Nucleic acid extraction 
			&#9679; Primer/probe design</td>
			<td>&#9679; Nucelic acid extraction 
			&#9679; Primer design</td>
			<td>&#9679; Protein extraction 
			&#9679; Antibodies</td>
			<td>N/A</td>
		</tr>
		<tr>
			<td>Other materials required</td>
			<td>&#9679; Portable thermocycler</td>
			<td>&#9679; Conventional thermocycler</td>
			<td>&#9679; Colormetric stain 
			&#9679; Incubator</td>
			<td>&#9679; Plate reader 
			&#9679; Washing equipment</td>
			<td>N/A</td>
		</tr>
	</tbody>
</table>
Table 1. Comparative chart of molecular and serological detection methods for phytopathogens
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Primer</td>
			<td colspan="2">Sequence (5&#8242;&#8211;3&#8242;)a</td>
			<td>Targetb</td>
		</tr>
		<tr>
			<td>SsTQ-F13</td>
			<td colspan="2">CCGGCAGACCCAAAACC</td>
			<td>ITS1-ITS2 in S. subterranea</td>
		</tr>
		<tr>
			<td>SsTQ-R13</td>
			<td colspan="2">CGGGCGTCACCCTTCA</td>
			<td>ITS1-ITS2 in S. subterranea</td>
		</tr>
		<tr>
			<td>SsTQ-P13</td>
			<td colspan="2">[FAM]CAGACAATCGCACCCAGGTTCTCATG[TAM]</td>
			<td>ITS1-ITS2 in S. subterranea</td>
		</tr>
		<tr>
			<td>Genesig S.subterranea primer/probe</td>
			<td colspan="2">N/A</td>
			<td>Actin in S. subterranea</td>
		</tr>
		<tr>
			<td>SPO1014</td>
			<td colspan="2">GGTCGGTCCATGGCTTGA</td>
			<td>ITS in S. subterranea</td>
		</tr>
		<tr>
			<td>SPO1114</td>
			<td colspan="2">GGCACGCCAATGGTTAGAGA</td>
			<td>ITS in S. subterranea</td>
		</tr>
		<tr>
			<td>SPOPRO114</td>
			<td colspan="2">[FAM]CCGGTGCGCGTCTCTGGCTT[TAM]</td>
			<td>ITS in S. subterranea</td>
		</tr>
		<tr>
			<td>RsTqF119</td>
			<td colspan="2">AAGAGTTTGGTTGTAGCTGGTCTATTT</td>
			<td>ITS1-ITS2 in R. solani</td>
		</tr>
		<tr>
			<td>RsTqR119</td>
			<td colspan="2">AATTCCCCAACTGTCTCACAAGTT</td>
			<td>ITS1-ITS2 in R. solani</td>
		</tr>
		<tr>
			<td>RQP119</td>
			<td colspan="2">[FAM]TTTAGGCATGTGCACACCTCCCTCTTTC[TAM]</td>
			<td>ITS1-ITS2 in R. solani</td>
		</tr>
		<tr>
			<td>Genesig PMTV primer/probe</td>
			<td colspan="2">N/A</td>
			<td>CP-RT in PMTV</td>
		</tr>
		<tr>
			<td>PMTV-D-F20</td>
			<td colspan="2">AGAATTGRCATCGAAACAGCA</td>
			<td>CP in PMTV</td>
		</tr>
		<tr>
			<td>PMTV-D-R20</td>
			<td colspan="2">GTCGCGCTCCAATTTCGTT</td>
			<td>CP in PMTV</td>
		</tr>
		<tr>
			<td>PMTV-D-P20</td>
			<td colspan="2">[FAM]CCACAAACAGACAGGTATGGTCCGGAA[TAM]</td>
			<td>CP in PMTV</td>
		</tr>
		<tr>
			<td colspan="4">a Oligo DNA primers were modified with FAM (6-carboxyfluorescein) or TAM (5-carboxytetramethylrhodamine)</td>
		</tr>
		<tr>
			<td colspan="4">b ITS: Internal transcribed spacers, CP: coat protein; CP-RT: coat protein readthrough</td>
		</tr>
	</tbody>
</table>
Table 2. Primers used in this study
Supplementary Figure 1. Comparison of the DNA extraction methods for the detection of the powdery scab pathogen. Six different DNA extraction methods (A-F) were compared for the detection of the powdery scab pathogen, S. subterranea in soil samples. (B, D, F). DNA was extracted using silica-based kit #1 (see the Table of Materials for all kit names), silica-based kit #2, and magnetic bead-based kit, repsectively. PCR was performed using the conventional lab-based PCR thermocycler. Standard curves represent the relationship between the amount of total DNA extracted from the soil samples and the amounts of PCR product amplified by the SsTQ primers/probe set. <a href="/files/ftp_upload/56891/Supplementary_Figure_1.pdf" target="_blank">Please click here to download this figure.</a>
Supplementary Figure 2. Comparison of the limit of detection between a portable PCR and a conventional lab-based PCR. Total DNA was isolated from a soil sample using three different extraction methods: (A, B) Doyle method, (C, D) the silica-based kit #2, and (E, F) the magnetic bead-based kit. Graphs shown on the left are data using the portable thermocycler with the Sss primers/probe set, while the graphs on the right represent data generated using the conventional lab-based thermocycler with the SsTQ primers/probe set. <a href="/files/ftp_upload/56891/Supplementary_Figure_2.pdf" target="_blank">Please click here to download this figure.</a>

Watch this Scientific Journal Video about On-Site Molecular Detection of Soil-Borne Phytopathogens Using a Portable Real-Time PCR System at JoVE.com

On-Site Molecular Detection of Soil-Borne Phytopathogens Using a Portable Real-Time PCR System

Rapid and accurate detection of plant pathogens on-site, especially soil-borne pathogens, is crucial to prevent further inoculum production and proliferation of plant diseases in the field. The method developed here using a portable real-time PCR detection system enables on-site diagnosis under field conditions.

On-site diagnosis of plant diseases can be a useful tool for growers for timely decisions enabling the earlier implementation of disease management ...

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16.4K Views.  Washington State University. Rapid and accurate detection of plant pathogens on-site, especially soil-borne pathogens, is crucial to prevent further inoculum production and proliferation of plant diseases in the field. The method developed here using a portable real-time PCR detection system enables on-site diagnosis under field conditions.

Video: On-Site Molecular Detection of Soil-Borne Phytopathogens Using a Portable Real-Time PCR System

Simultaneous DNA-RNA Extraction from Coastal Sediments and Quantification of 16S rRNA Genes and Transcripts by Real-time PCR

Real Time Polymerase Chain Reaction also known as quantitative PCR (q-PCR) is a widely used tool in microbial ecology to quantify gene abundances of taxonomic and functional groups in environmental samples. Used in combination with a reverse transcriptase reaction (RT-q-PCR), it can also be employed to quantify gene transcripts. q-PCR makes use of highly sensitive fluorescent detection chemistries that allow quantification of PCR amplicons during the exponential phase of the reaction. Therefore, the biases associated with &#39;end-point&#39; PCR detected in the plateau phase of the PCR reaction are avoided. A protocol to quantify bacterial 16S rRNA genes and transcripts from coastal sediments via real-time PCR is provided. First, a method for the co-extraction of DNA and RNA from coastal sediments, including the additional steps required for the preparation of DNA-free RNA, is outlined. Second, a step-by-step guide for the quantification of 16S rRNA genes and transcripts from the extracted nucleic acids via q-PCR and RT-q-PCR is outlined. This includes details for the construction of DNA and RNA standard curves. Key considerations for the use of RT-q-PCR assays in microbial ecology are included.

A protocol to quantify bacterial 16S rRNA genes and transcripts from coastal sediments via real-time PCR is provided. The methodology includes the co-extraction of DNA and RNA; preparation of DNA-free RNA; and 16S rRNA gene and transcript quantification via RT-q-PCR, including standard curve construction.

Real Time Polymerase Chain Reaction also known as quantitative PCR (q-PCR) is a widely used tool in microbial ecology to quantify gene abundances of ...

Measuring and Mapping Patterns of Soil Erosion and Deposition Related to Soil Carbonate Concentrations Under Agricultural Management

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonate (CaCO3) profiles. The objective is to describe a simple conceptual model and detailed protocol for repeatable field and laboratory measurements of these quantities. Here, accurate elevation is measured using a ground-based differential global positioning system (GPS); other data acquisition methods could be applied to the same basic method. Soil samples are collected from prescribed depth intervals and analyzed in the lab using an efficient and precise modified pressure-calcimeter method for quantitative analysis of inorganic carbon concentration. Standard statistical methods are applied to point data, and representative results show significant correlations between changes in soil surface layer CaCO3 and changes in elevation consistent with the conceptual model; CaCO3 generally decreased in depositional areas and increased in erosional areas. Maps are derived from point measurements of elevation and soil CaCO3 to aid analyses. A map of erosional and depositional patterns at the study site, a rain-fed winter wheat field cropped in alternating wheat-fallow strips, shows the interacting effects of water and wind erosion affected by management and topography. Alternative sampling methods and depth intervals are discussed and recommended for future work relating soil erosion and deposition to soil CaCO3.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes in elevation are related to changes in near-surface soil carbonates. Repeatable methods for field and laboratory measurements of these quantities and data analysis methods are described here.

Spatial patterns of soil erosion and deposition can be inferred from differences in ground elevation mapped at appropriate time increments. Such changes ...

Specific and Accurate Detection of the Citrus Greening Pathogen Candidatus liberibacter spp. Using Conventional PCR on Citrus Leaf Tissue Samples

Citrus greening, also known as huanglongbing, is a destructive citrus disease ravaging citrus farms globally. This disease causes asymmetrical yellow leaf mottling, vein yellowing, defoliation, root decay, and ultimately, the death of the citrus plant. When infected, the citrus plants have stunted growth and produce flowers out of season. These flowers rarely yield fruit, and those that do yield small, bitter, irregularly shaped citrus fruit that are not desirable. This disease is spread by the Asian citrus psyllid, Diaphorina citri, and by the grafting of infected citrus tissue. The pathogen has a long and variable incubation period within the citrus plant&#8212;sometimes years, before symptoms appear. Attempts to culture this pathogen in vitro have been unsuccessful, possibly due to the low and uneven concentration of the pathogen within infected citrus tissue, or because it is difficult to replicate the environmental conditions conducive to growth of the pathogen. It is very difficult to identify the disease before it has spread, due to its long incubation period and researchers&#39; inability to culture the pathogen. As a result, the disease only becomes apparent after suddenly destroying a citrus farmer&#39;s entire yield. Presented here is a method for the accurate and specific detection of the citrus greening pathogen, Candidatus liberibacter spp. using a genomic DNA extraction kit and PCR. This method is simple, efficient, cost effective, and adaptable for quantitative analysis. This method can be adapted for use on any citrus tissue; however, it is potentially limited by the amount of pathogen present in the tissue. Nevertheless, this method will allow citrus farmers to identify infected citrus plants earlier, and curb the spread of this destructive disease before it can further spread.

Specific and Accurate Detection of the Citrus Greening Pathogen Candidatus liberibacter  spp. Using Conventional PCR on Citrus Leaf Tissue Samples

Citrus Greening is a particularly destructive disease affecting citrus crops globally. Presented here is a simple method using PCR and genomic DNA extraction of citrus leaf tissue for the accurate and precise identification of the citrus greening pathogen, Candidatus liberibacter spp.

Citrus greening, also known as huanglongbing, is a destructive citrus disease ravaging citrus farms globally. This disease causes asymmetrical yellow leaf ...

Self-standing Electrochemical Set-up to Enrich Anode-respiring Bacteria On-site

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be critical for microbial energy production and persistence in many subsurface environments, especially those lacking soluble terminal electron acceptors. While EET-capable microbes have been successfully isolated from various environments, the diversity of bacteria capable of EET is still poorly understood, especially in difficult-to-sample, low energy or extreme environments, such as many subsurface ecosystems. Here, we describe an on-site electrochemical system to enrich EET-capable bacteria using an anode as a respiratory terminal electron acceptor. This anode is connected&#160;to a cathode capable of&#160;catalyzing abiotic oxygen reduction. Comparing this approach with electrocultivation methods that use a potentiostat for poising the electrode potential, the two-electrode system does not require an external power source. We present an example of our on-site enrichment utilized in an alkaline pond at the Cedars, a terrestrial serpentinization site in Northern California. Prior attempts to cultivate mineral reducing bacteria were unsuccessful, which is likely due to the low-biomass nature of this site and/or the low relative abundance of metal reducing microbes. Prior to implementing our two-electrode enrichment, we measured the vertical profile of dissolved oxygen concentration. This allowed us to place the carbon felt anode and platinum-electroplated carbon felt cathode at depths that would support anaerobic and aerobic processes, respectively. Following on-site incubation, we further enriched the anodic electrode in the laboratory and confirmed&#160;a distinct microbial community compared to the surface-attached or biofilm communities normally observed at the Cedars. This enrichment subsequently led to the isolation of the first electrogenic microbe from the Cedars. This method of on-site microbial enrichment has the potential to greatly enhance the isolation of EET-capable bacteria from low biomass or difficult to sample habitats.

On-site microbial enrichment or in situ cultivation techniques can facilitate the isolation of difficult-to-culture microbial taxa, especially from low-biomass or geochemically extreme environments. Here, we describe an electrochemical set-up without using an external power source to enrich microbial strains that are capable of extracellular electron transport (EET).

Anaerobic respiration coupled with electron transport to insoluble minerals (referred to as extracellular electron transport [EET]) is thought to be ...

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

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

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

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

Detection of Viruses from Bioaerosols Using Anion Exchange Resin

This protocol demonstrates a customized bioaerosol sampling method for viruses. In this system, anion exchange resin is coupled with liquid impingement-based air sampling devices for efficacious concentration of negatively-charged viruses from bioaerosols. Thus, the resin serves as an additional concentration step in the bioaerosol sampling workflow. Nucleic acid extraction of the viral particles is then performed directly from the anion exchange resin, with the resulting sample suitable for molecular analyses. Further, this protocol describes a custom-built bioaerosol chamber capable of generating virus-laden bioaerosols under a variety of environmental conditions and allowing for continuous monitoring of environmental variables such as temperature, humidity, wind speed, and aerosol mass concentration. The main advantage of using this protocol is increased sensitivity of viral detection, as assessed via direct comparison to an unmodified conventional liquid impinger. Other advantages include the potential to concentrate diverse negatively-charged viruses, the low cost of anion exchange resin (~$0.14 per sample), and ease of use. Disadvantages include the inability of this protocol to assess infectivity of resin-adsorbed viral particles, and potentially the need for the optimization of the liquid sampling buffer used within the impinger.

An anion exchange resin-based method, adapted to liquid impingement-based bioaerosol sampling of viruses is demonstrated. When coupled with downstream molecular detection, the method allows for facile and sensitive detection of viruses from bioaerosols.

This protocol demonstrates a customized bioaerosol sampling method for viruses. In this system, anion exchange resin is coupled with liquid ...

A Precise and Autonomous System for the Detection of Insect Emergence Patterns

Existing systems to measure insect emergence patterns have limitations; they are only partially automated and are limited in the maximum number of emerging insects they can detect. In order to obtain precise measurement of insect emergence, it is necessary for systems to be semi-automated and able to measure large numbers of emerging insects. We addressed these issues by designing and building a system that is automated and can measure emergence of up to 1200 insects. We modified the existing &#34;falling-ball&#34; system using Arduino microcontrollers to automate data collection and expand the sample size through multiple data channels. Multiple data channels enable the user to not only increase their sample size, but also allows for multiple treatments to be run simultaneously in a single experiment. Furthermore, we created an R script to automatically visualize the data as a bubble plot, while also calculating the median day and time of emergence. The current system was designed using 3D printing so the user can modify the system to be adjusted for different species of insects. The goal of this protocol is to investigate important questions in chronobiology and stress physiology, using this precise and automated system to measure insect emergence patterns.

Measurement of insect emergence patterns requires precision. Existing systems are only semi-automated and sample size is limited. We addressed these issues by designing a system using microcontrollers to precisely measure the time of emergence of large numbers of emerging insects.

Existing systems to measure insect emergence patterns have limitations; they are only partially automated and are limited in the maximum number of ...

Detection of 3-Nitrotyrosine in Atmospheric Environments via a High-performance Liquid Chromatography-electrochemical Detector System

3-nitrotyrosine (3-NT) is generated from the tyrosine residue in atmospheric bio-aerosol proteins via a reaction with ozone (O3) and nitrogen dioxide (NO2). Stable 3-NT is a specific marker for oxidative damage and is reported to have a promotive effect to elicit allergies. In the present study, we report the development of a highly sensitive assay to quantify 3-NT in air sampler filters to collect &#60; 2.5 &#181;m of particulate matter (PM2.5) from urban environmental air, including bio-aerosol. In this method, a 6 mm-diameter round hole was cut from the filters of air samplers and mixed with a nonspecific protease cocktail in order to hydrolyze proteins. Protein samples digested to the amino acid level were tested for 3-NT using a high-performance liquid chromatography-electrochemical detector (HPLC-ECD). The maximum 3-NT content was detected in a prefilter for PM of sizes from 4.5 to 7.3 &#956;m, with a detection limit of 1.13 pg/m3.

Detection of 3-Nitrotyrosine in Atmospheric Environments via a High-performance Liquid Chromatography-electrochemical Detector System

We present a method to detect 3-nitrotyrosine chemical modifications of atmospheric proteins with 6 mm-diameter rounds cut from air sampler prefilters using a high-performance liquid chromatography-electrochemical detector (HPLC-ECD).

3-nitrotyrosine (3-NT) is generated from the tyrosine residue in atmospheric bio-aerosol proteins via a reaction with ozone (O3) and nitrogen dioxide ...

Detection of Tilapia Lake Virus Using Conventional RT-PCR and SYBR Green RT-qPCR

The aim of this method is to facilitate the fast, sensitive and specific detection of Tilapia Lake Virus (TiLV) in tilapia tissues. This protocol can be used as part of surveillance programs, biosecurity measures and in TiLV basic research laboratories. The gold standard of virus diagnostics typically involves virus isolation followed by complementary techniques such as reverse-transcription polymerase chain reaction (RT-PCR) for further verification. This can be cumbersome, time-consuming and typically requires tissue samples heavily infected with virus. The use of RT-quantitative (q)PCR in the detection of viruses is advantageous because of its quantitative nature, high sensitivity, specificity, scalability and its rapid time to result. Here, the entire method of PCR based approaches for TiLV detection is described, from tilapia organ sectioning, total ribonucleic acid (RNA) extraction using a guanidium thiocyanate-phenol-chloroform solution, RNA quantification, followed by a two-step PCR protocol entailing, complementary deoxyribonucleic acid (cDNA) synthesis and detection of TiLV by either conventional PCR or quantitative identification via qPCR using SYBR green I dye. Conventional PCR requires post-PCR steps and will simply inform about the presence of the virus. The latter approach will allow for absolute quantification of TiLV down to as little as 2 copies and thus is exceptionally useful for TiLV diagnosis in sub-clinical cases. A detailed description of the two PCR approaches, representative results from two laboratories and a thorough discussion of the critical parameters of both have been included to ensure that researchers and diagnosticians find their most suitable and applicable method of TiLV detection.

This protocol diagnoses Tilapia Lake Virus (TiLV) in tilapia tissues using RT-PCR methodologies. The entire method is described from tissue dissection to total RNA extraction, followed by cDNA synthesis and detection of TiLV by either conventional PCR or quantitative PCR using dsDNA binding a fluorescent binding dye.

The aim of this method is to facilitate the fast, sensitive and specific detection of Tilapia Lake Virus (TiLV) in tilapia tissues. This protocol can be ...

A New Portable In Vitro Exposure Cassette for Aerosol Sampling

This protocol introduces a new in vitro exposure system, capable of being worn, including its characterization and performance. Air-liquid interface (ALI) in vitro exposure systems are often large and bulky, making transport to the field and operation at the source of emission or within the breathing zone difficult. Through miniaturization of these systems, the lab can be brought to the field, expediting processing time and providing a more appropriate exposure method that does not alter the aerosol prior to contacting the cells. The Portable In vitro Exposure Cassette (PIVEC) adapts a 37 mm filter cassette to allow for in vitro toxicity testing outside of a traditional laboratory setting. The PIVEC was characterized using three sizes of copper nanoparticles to determine deposition efficiency based on gravimetric and particle number concentration analysis. Initial cytotoxicity experiments were performed with exposed lung cells to determine the ability of the system to deposit particles while maintaining cell viability. The PIVEC provides a similar or increased deposition efficiency when comparing to available perpendicular flow in vitro exposure devices. Despite the lower sample throughput, the small size gives some advantages to the current in vitro ALI exposure systems. These include the ability to be worn for personal monitoring, mobility from the laboratory to the source of emission, and the option to set-up multiple systems for spatial resolution while maintaining a lower user cost. The PIVEC is a system capable of collecting aerosols in the field and within the breathing zone onto an air-interfaced, in vitro model.

A New Portable In Vitro Exposure Cassette for Aerosol Sampling

Here, we present a protocol to perform portable cellular aerosol exposures and measure cellular response. The method uses cells, grown at the air-liquid interface, mimicking in vivo physiology. Cellular response to copper nanoparticle aerosols was observed as oxidative stress through reactive oxygen species generation and cytotoxicity as lactate dehydrogenase release.

This protocol introduces a new in vitro exposure system, capable of being worn, including its characterization and performance. Air-liquid interface (ALI) ...