This work received the financial support of Georgia Commodity Commission for Vegetables project ID# FP00016659. The authors thank Dr. Pingsheng Ji, University of Georgia and Dr. Anne Dorrance, Ohio State University for providing pure cultures of Phytophthora spp. We also thank Li Wang and Deloris Veney for their technical assistance throughout the study.

1. On-site detection of Phytophthora capsici from irrigation water using portable loop-mediated isothermal amplification
<ol>
	<li>Setting up the pump and filter
	<ol>
		<li>Attach a filtering flask to a tube that is connected to a hand pump so that when the pump is activated, air will be pulled in through the mouth of the filtering flask.</li>
		<li>Fit the Buchner funnel into the rubber stopper to the mouth of the filtering flask and fit the appropriately sized piece of filter paper into the Buchner funnel so that air is pulled through the filter paper. The filter paper should have a retention size of 15 &#181;m. 
		NOTE: The filter paper must fit to the edges of the Buchner funnel so that minimal water will flow around the filter paper.</li>
	</ol>
	</li>
	<li>Water sampling and filtering
	<ol>
		<li>Take water samples from the targeted source. Water may have small amounts of debris but not significant sediment or soil.</li>
		<li>Pour up to 1,000 mL (1 Liter)&#160;of test water over the filter paper placed inside the Buchner funnel slowly enough to prevent overflow, while the hand pump (or vacuum) is being used to create a suction to pull the water through. 
		NOTE: There is no minimum amount of water that can be tested using this method, and although it at least 50 mL is suggested, 1000 mL is the maximum for this method.</li>
		<li>Using forceps, remove the filter paper from the Buchner funnel and cut it into small pieces with sterile scissors. Add as many pieces (8-12) as can be submerged in the amount of extraction buffer (400 &#181;L for magnetic bead-based extraction) required by the protocol into a 1.5 mL tube. Save remaining pieces of filter paper for processing after the first set has been extracted.</li>
		<li>Vortex or otherwise agitate the pieces of filter paper and extraction buffer for 10 s every minute for 5 min. Then, using forceps, remove the filter paper losing as little of the extraction&#160;buffer as possible. Repeat this step with the remaining pieces of filter paper until all pieces have been vortexed/agitated and soaked in the extraction&#160;buffer.</li>
	</ol>
	</li>
	<li>Magnetic bead-based extraction of DNA from filter paper
	<ol>
		<li>To the 1.5 mL tube (which now contains approximately 200-300 &#181;L of extraction&#160;buffer), add 20 &#181;L of proteinase K and 10 &#181;L of 10 ng/&#181;L RNase.</li>
		<li>Incubate at room temperature for 15 min, vortex or shake the tube every 3 min.</li>
		<li>Add 500 &#181;L of magnetic beads with the binding buffer to the sample and mix well by shaking. Then incubate for 5 min at room temperature.</li>
		<li>Place the tube in the magnetic separator rack for 2 min until all beads have been pulled to the magnet. Remove and discard the supernatant.</li>
		<li>Remove the tube from the magnetic separator. Add 500 &#181;L of Wash Buffer 1 and re-suspend beads by shaking the tube vigorously. Wait for 30 s and then place the tube back in the magnetic separator. Wait for 2 min until all beads have been pulled towards the magnet before removing and discarding the supernatant. 
		NOTE: When waiting for the magnetic beads to magnetize to the separator, we recommend inverting the tube, which can dislodge magnetic beads stuck to the cap and the sides of the tube and result in a greater number of beads being attached.</li>
		<li>Repeat step 1.3.5 with 500 &#181;L of Wash Buffer 2.</li>
		<li>Repeat step 1.3.5 with 500 &#181;L of 80% ethanol.</li>
		<li>Airdry the magnetic bead pellet for 15 min at room temperature (18-27 &#176;C) with the lid open. If temperatures do not permit, incubate in a gloved hand with the cap open for 15 min.</li>
		<li>Remove the tube from the magnetic separator. Add 50 &#181;L of elution buffer and re-suspend the beads by pipetting up and down for 1 min.</li>
		<li>Place tube back in a magnetic separator. Wait 2 min before transferring the supernatant without disturbing the beads to a separate tube for DNA storage. 
		NOTE: Here the experiment can be paused before moving on. Extracted DNA should be stored on ice or in a -20 &#176;C freezer.</li>
	</ol>
	</li>
	<li>Application of newly developed LAMP assay
	<ol>
		<li>Prepare LAMP primer mix using 0.2 &#181;M of each F3 and B3 primer, 0.8 &#181;M of each Loop-F and Loop-B primer, and 1.6 &#181;M of each FIP and BIP primer (Table 2).</li>
		<li>Add the following LAMP solution to a single PCR tube, or to each individual tube in the 8 tube strip: 2.5 &#181;L of primer mix (step 1.4.1), 12.5 &#181;L of LAVA LAMP master mix, 1 &#181;L of extracted DNA, and 9 &#181;L of ddH2O. The total volume is 25 &#181;L. 
		NOTE: If using the portable amplification instrument (e.g., Genie III) with colorimetric dye (e.g., Warmstart), refer to section 2.4.2- 2.4.3.</li>
		<li>Designate two tubes as positive and negative controls. For the positive control, use either a control provided by the LAVA LAMP kit, or a known positive DNA sample. For the negative control, use ddH2O. For both, substitute the 1 &#181;L of extracted DNA for 1 &#181;L of the positive control or ddH2O.</li>
		<li>Set the samples into a heat block (or portable amplification instrument) set to 64 &#176;C for 45 min. 
		NOTE: Other extraction methods can be used here instead of magnetic bead based extraction. CTAB and a commercial DNA extraction kit were both successfully tested using the standard protocols and substituting the filter paper pieces for the plant sample24,25. Results were compared in Table 2. If concentration of DNA can be quantified, use between 1-10 ng genomic DNA.</li>
	</ol>
	</li>
	<li>Visualization of results
	<ol>
		<li>If performed in a laboratory setting, view the amplification products by loading 5 &#181;L of each sample into a 1% agarose gel, running them in a gel electrophoresis machine, and imaging them in a UV imaging machine.</li>
		<li>If a colorimetric dye (e.g., Warmstart) was used, view the color change to determine results as positive or negative.</li>
		<li>If a portable amplification instrument (e.g., Genie III) was used, view the amplification graph on the screen to determine the results.</li>
	</ol>
	</li>
	<li>Application of previously developed PCR assay 
	NOTE: If using a conventional PCR assay, steps 1-3 remain the same, and the following steps should be applied in place of steps 1.4 and 1.5.
	<ol>
		<li>Add 1 &#181;L of each DNA extraction to individual tubes containing the following components: 12.5 &#181;L of Green PCR master mix, 9.5 &#181;L of ddH2O, and 1 &#181;L of forward and reverse primers (Table 2).</li>
		<li>Spin down each sample using a microcentrifuge and place tubes into a thermal cycler.</li>
		<li>Use the following thermal cycler settings in accordance to the previous publications: 94 &#176;C for 5 min, 30 cycles of denaturation at 94 &#176;C for 30 s, annealing at 54 &#176;C for 30 s, extension at 72 &#176;C for 1 min, and final extension at 72 &#176;C for 10 min.</li>
		<li>Run the product in a 1% agarose gel. Observe the presence of bands under UV light where positive reactions will have a band size of ~508 bp.</li>
	</ol>
	</li>
	<li>Traditional method of detection 
	NOTE: There are multiple methods of selective plating for pathogen detection, and the following is a general protocol for P. capsici.
	<ol>
		<li>First, obtain a healthy eggplant fruit (a susceptible host for P. capsici) and surface sterilize by washing the fruit surface with 70% isopropyl alcohol.</li>
		<li>Place the eggplant fruit into milk crates with a flotation device (polyethylene foam or other) and deploy into irrigation ponds. Secure each bait trap to a single point and leave the traps in the water reservoir (recycled irrigation water) for at least 7 days or until fruit rot symptoms are observed. Collect the fruit and transport it to the laboratory.</li>
		<li>Rinse and dry the fruit in a sterile hood before removing small pieces of infected tissues and place them on a plate of PARPH medium amended with 25 mg/L pentachloronitrobenzene, 0.0005% pimaricin, 250 mg/L ampicillin, 10 mg/L rifampicin and 50 mg/L hymexazol. Incubate plates at 25 &#176;C for 5 days.</li>
		<li>View plates under a compound microscope for traditional morphological identification at 4 days after isolation.</li>
	</ol>
	</li>
</ol>
2. Determining the detection limit of zoospore concentration
<ol>
	<li>Making zoospore suspension
	<ol>
		<li>Incubate P. capsici on V8 agar (100 mL of V8 juice, 900 mL of ddH2O, 1 g of CaCO3) plates for 1 week at 26 &#176;C. Multiple plates can be used to obtain a larger amount of zoospore suspension.</li>
		<li>Incubate the plates under continuous light at room temperature for 3 days to stimulate sporulation.</li>
		<li>Flood the plates by adding 15 mL of ddH2O to each plate and place them in a 4 &#176;C fridge for 25 min. Then return to room temperature for 30 min.</li>
		<li>Agitate plates to dislodge zoospores and pipette the solution into a single 50 mL tube from all the plates.</li>
		<li>To obtain an accurate estimate of zoospore concentration, add 10 &#181;L of the spore suspension onto a hemocytometer and observe under a microscope to count zoospores and estimate the average concentration.</li>
	</ol>
	</li>
	<li>Serial dilution
	<ol>
		<li>Add 1 mL of the spore suspension and 9 mL of ddH2O to a separate tube. Repeat this step for as many 10-fold dilutions as desired.</li>
		<li>Spore solutions are then submitted for DNA extraction based on the previous protocol using the filtration method. 
		NOTE: If a larger volume of suspension is desired, double the volume: 2 mL of spore suspension and 18 mL of ddH2O.</li>
	</ol>
	</li>
	<li>Detection of zoospore concentration limit
	<ol>
		<li>Evaluate spore detection limit by running each serial dilution individually through the assay until clearly positive results are no longer observed. Once the final dilution is obtained, dilute by a factor of 2 (4.8 to 2.4 in this example) and run the assay again to get a more accurate detection limit.</li>
	</ol>
	</li>
	<li>Development and optimization of the LAMP method 
	NOTE: LAMP primers were designed based on a 1121-base pair (bp) fragment of P. capsici (Li et al.19) as shown in Supplementary Figure 2.
	<ol>
		<li>If colorimetric dye (e.g., Warmstart) is used, use the following the solution: 2.5 &#181;L of primer mix, 12.5 &#181;L of colorimetric dye, 0.5 &#181;L of Green fluorescent dye, 1 &#181;L of extracted DNA, and 8.5 &#181;L of ddH2O. The total volume is 25 &#181;L.</li>
		<li>When using the portable amplification instrument with LAVALAMP mastermix, have an initial step of 95 &#176;C for 3 min as recommended by the manufacturer, but this is not required. A final annealing step is not required to observe the color change or amplification graph. Do not run a warmstart step if the commercial colorimetric dye is to be used.</li>
		<li>View LAMP assay results in one of the following methods: run samples on a 1% agarose gel or view using a UV imaging machine with the naked eye or view at the Genie III real-time amplification screen.</li>
		<li>Optimize the temperature of the LAMP assay by using the portable amplification instrument and analyzed using the real-time amplification graph for speed and level of sensitivity. Run samples with unique temperatures to determine the fastest amplification with the highest level of sensitivity.</li>
		<li>Determine the detection limit of the LAMP developed assay by making a serial dilution of extracted DNA (as with spore suspension in step 2.2.1) and maintaining reaction conditions as previously described for the LAMP reaction for each dilution.</li>
	</ol>
	</li>
	<li>Detection limit determination and comparison with conventional PCR method
	<ol>
		<li>Use DNA extracted in steps in section 1.3 to compare the detection level of conventional PCR with that of the LAMP assay.</li>
		<li>Add 1 &#181;L of DNA to a PCR tube that contained 1 &#181;L of both forward and reverse PCR primers (Table 2), 12.5 &#181;L of Green PCR Mastermix, and 9.5 &#181;L of ddH2O for a total of 25 &#181;L.</li>
		<li>Amplify samples in a thermal cycler using the following conditions: 94 &#176;C for 5 min, 30 cycles of denaturation at 94 &#176;C for 30 s, annealing at 54 &#176;C for 30 s, extension at 72 &#176;C for 1 min, and final extension at 72 &#176;C for 10 min.</li>
		<li>Run samples in an electrophoresis machine on a 1% agarose gel and view on a UV imaging machine. The excepted band size was 508 bp.</li>
	</ol>
	</li>
</ol>

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A. <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+a+colorimetric+loop-mediated+isothermal+amplification+assay+for+the+visual+detection+of+Fusarium+oxysporum+f.+sp.+melonis.">Development of a colorimetric loop-mediated isothermal amplification assay for the visual detection of Fusarium oxysporum f. sp. melonis.</a> Horticultural Plant Journal. 5 (3), 129-136 (2019).</li><li>Gill, D. J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pathogenic+Pythium+from+irrigation+ponds.">Pathogenic Pythium from irrigation ponds.</a> Plant Disease Reporter. 54 (12), 1077-1079 (1970).</li></ol>

The authors have nothing to disclose or any conflicts of interest.

The testing of irrigation water for phytopathogens is a crucial step for growers using irrigation ponds and recycled water27. Irrigation ponds provide a reservoir and breeding ground for a number of phytopathogens as excess irrigation water is directed from the field to the pond carrying with it any pathogens that may have been present16,27. The traditional method for detection of a plant pathogens in a large water source is to set a bait ...

Recycling water in farms and nurseries is becoming increasingly popular due to the increase in water costs and environmental concerns behind water usage. Many irrigation methods have been developed for growers to reduce the spread and occurrence of plant disease. Regardless of the source of the water (irrigation or precipitation), runoff is generated, and many vegetable and nursery growers have a pond to collect and recycle runoff1. This creates a reservoir for possible pathogen accumulation favoring the spread of pathogens when the recycled water is used to irrigate crops2,3,4. Oomycete plant pathogens particularly benefit from this practice as zoospores will accumulate in water and the primary dispersive spore is self-motile but requires surface water5,6,7. Phytophthora capsici is an oomycete pathogen that affects a significant number of solanaceous and cucurbit crops in different ways8. Often, the symptoms are damping-off of seedlings, root and crown rot; however, in crops such as cucumber, squash, melon, pumpkin, watermelon, eggplant and pepper, entire harvests may be lost due to fruit rot9. Although there are known methods of detecting this plant pathogen, most require an infection to have already taken place which is too late for any preventative fungicides to have a significant effect10.
The traditional method to test irrigation water for the detection and diagnosis of targeted microorganisms is an antiquated approach when speed and sensitivity are crucial to success and profitable crop production11,12. Plant tissue susceptible to the targeted pathogen (e.g., eggplant for P. capsici) is attached to a modified trap that is suspended in an irrigation pond for extended period before being removed and inspected for infection. Samples from the plant tissue are then plated on semi-selective media (PARPH) and incubated for culture growth, then morphological identification is performed using a compound microscope13. There are other similar detection methods for other plant pathogens using selective media and plating small amounts of contaminated water before sub-culturing14,15. These methods require anywhere from 2 to 6 weeks, several rounds of sub-culturing to isolate the organism, and experience on Phytophthora diagnostics to be able to recognize the key morphological characters of each species. These traditional methods do not work well for detection of irrigation water contaminated by P. capsici due to factors such as interference by other microorganisms that are also present in the water sources. Some fast-growing microorganisms like Pythium spp. and water-borne bacteria can overgrow on the plate making P. capsici undetectable16,17.
The purpose of this study was to develop a sensitive and specific molecular method that can be used in both field and laboratory settings to detect P. capsici zoospores in irrigation water. The protocol includes the development of a novel loop-mediated isothermal amplification (LAMP) primer set able to specifically amplify P. capsici, based on a 1121-base pair (bp) fragment of P. capsici18,19. A previously developed LAMP primer from Dong et al. (2015) was used in comparison to the assay that was developed for this study20.
The LAMP assay is a relatively new form of molecular detection that has been demonstrated to be more rapid, sensitive, and specific than conventional polymerase chain reaction (PCR)21. In general, conventional PCR assays cannot detect under 500 copies (1.25 pg/&#181;L); in contrast, previous studies have shown that the sensitivity of LAMP can be 10 to 1,000 times higher than conventional PCR and can easily detect even 1 fg/&#181;L of genomic DNA22,23. Additionally, the assay can be carried out rapidly (often in 30 min) and on-site (in the field) by using a portable heating block for amplification and a colorimetric dye that changes color for a positive sample (removing the need for electrophoresis). In this study, we compared the sensitivity of PCR and LAMP assays using a filter extraction method. The proposed detection method allows researchers and extension agents to easily detect the presence of P. capsici spores from different water sources in less than two hours. The assay is proven to be more sensitive than conventional PCR and was validated in situ by detecting the presence of the pathogen in the irrigation water used by a grower. This detection method will allow growers to estimate the presence and population density of the pathogen in various water sources that are being used for irrigation, preventing devastating outbreaks and economic losses.

<table>
	<tbody>
		<tr>
			<td>Agarose gel powder</td>
			<td>Thomas Scientific</td>
			<td>C997J85</td>
			<td></td>
		</tr>
		<tr>
			<td>Buchner funnel</td>
			<td>Southern Labware</td>
			<td>JBF003</td>
			<td></td>
		</tr>
		<tr>
			<td>Bullet Blender</td>
			<td>Next Advance</td>
			<td>BBX24</td>
			<td></td>
		</tr>
		<tr>
			<td>Centrifuge 5430</td>
			<td>Eppendorf</td>
			<td>22620509</td>
			<td></td>
		</tr>
		<tr>
			<td>Chloroform</td>
			<td>Fischer Scientific</td>
			<td>C298-500</td>
			<td></td>
		</tr>
		<tr>
			<td>CTAB solution</td>
			<td>Biosciences</td>
			<td>786-565</td>
			<td></td>
		</tr>
		<tr>
			<td>Dneasy Extraction Kit</td>
			<td>Qiagen</td>
			<td>69104</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter Flask</td>
			<td>United</td>
			<td>FHFL1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Filter Paper</td>
			<td>United Scientific Supplies</td>
			<td>FPR009</td>
			<td></td>
		</tr>
		<tr>
			<td>Gel Green 10000X</td>
			<td>Thomas Scientific</td>
			<td>B003B68 (1/EA)</td>
			<td></td>
		</tr>
		<tr>
			<td>Genie III</td>
			<td>OptiGene</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Hand pump</td>
			<td>Thomas Scientific</td>
			<td>1163B06</td>
			<td></td>
		</tr>
		<tr>
			<td>Iso-amyl Alcohol</td>
			<td>Fischer Scientific</td>
			<td>BP1150-500</td>
			<td></td>
		</tr>
		<tr>
			<td>LAVA LAMP master mix</td>
			<td>Lucigen</td>
			<td>30086-1</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic bead DNA extraction</td>
			<td>Genesig</td>
			<td>genesigEASY-EK</td>
			<td></td>
		</tr>
		<tr>
			<td>Magnetic Separator</td>
			<td>Genesig</td>
			<td>genesigEASY-MR</td>
			<td></td>
		</tr>
		<tr>
			<td>polyvinylpyrrolidone</td>
			<td>Sigma Aldrich</td>
			<td>PVP40-500G</td>
			<td></td>
		</tr>
		<tr>
			<td>Primers</td>
			<td>Sigma Aldrich</td>
			<td></td>
			<td></td>
		</tr>
		<tr>
			<td>Prism Mini Centrifuge</td>
			<td>Labnet</td>
			<td>C1801</td>
			<td></td>
		</tr>
		<tr>
			<td>T100 Thermal Cycler</td>
			<td>Bio-Rad</td>
			<td>1861096</td>
			<td></td>
		</tr>
		<tr>
			<td>UV Gel Doc</td>
			<td>Analytik Jena</td>
			<td>849-00502-2</td>
			<td></td>
		</tr>
		<tr>
			<td>Warmstart Colorimetric Dye</td>
			<td>Lucigen</td>
			<td>E1800m</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide Mini ReadySub-Cell GT Cell</td>
			<td>Bio-Rad</td>
			<td>1704489EDU</td>
			<td></td>
		</tr>
		<tr>
			<td>70% isopropanol</td>
			<td>Fischer Scientific</td>
			<td>A451-1</td>
			<td></td>
		</tr>
	</tbody>
</table>

detection of phytophthora capsici in irrigation water using loop-mediated isothermal amplification

Phytophthora capsici is a devastating oomycete pathogen that affects many important solanaceous and cucurbit crops causing significant economic losses in vegetable production annually. Phytophthora capsici is soil-borne and a persistent problem in vegetable fields due to its long-lived survival structures (oospores and chlamydospores) that resist weathering and degradation. The main method of dispersal is through the production of zoospores, which are single-celled, flagellated spores that can swim through thin films of water present on surfaces or in water-filled soil pores and can accumulate in puddles and ponds. Therefore, irrigation ponds can be a source of the pathogen and initial points of disease outbreaks. Detection of P. capsici in irrigation water is difficult using traditional culture-based methods because other microorganisms present in the environment, such as Pythium spp., usually overgrow P. capsici making it undetectable. To determine the presence of P. capsici spores in water sources (irrigation water, runoff, etc.), we developed a hand pump-based filter paper (8-10 &#181;m) method that captures the pathogen&#8217;s spores (zoospores) and is later used to amplify the pathogen&#8217;s DNA through a novel loop-mediated isothermal amplification (LAMP) assay designed for the specific amplification of P. capsici. This method can amplify and detect DNA from a concentration as low as 1.2 x 102 zoospores/mL, which is 40 times more sensitive than conventional PCR. No cross-amplification was obtained when testing closely related species. LAMP was also performed using a colorimetric LAMP master mix dye, displaying results that could be read with the naked eye for on-site rapid detection. This protocol could be adapted to other pathogens that reside, accumulate, or are dispersed via contaminated irrigation systems.

Optimization of LAMP method 
In this study, we detected the presence of Phytophthora capsici in irrigation water using a portable loop-mediated isothermal amplification (LAMP) assay. First, the proposed LAMP assay was optimized by testing different LAMP primer concentrations [F3, B3 (0.1&#8211;0.5 &#181;M each); LF, LB (0.5&#8211;1.0 &#181;M each) and FIP, BIP (0.8&#8211;2.4 &#181;M each)], durations (30&#8211;70 min), and temperatures (55&#8211;70 &#176;C). The final LAMP primer mix used i...

Watch this Scientific Journal Video about Detection of Phytophthora capsici in Irrigation Water using Loop-Mediated Isothermal Amplification at JoVE.com

Detection of Phytophthora capsici in Irrigation Water using Loop-Mediated Isothermal Amplification

We developed a method to detect Phytophthora capsici zoospores in water sources using a filter paper DNA extraction method coupled with a loop-mediated isothermal amplification (LAMP) assay that can be analyzed in the field or in the lab.

Detection of Phytophthora capsici in Irrigation Water using Loop-Mediated Isothermal Amplification

Phytophthora capsici is a devastating oomycete pathogen that affects many important solanaceous and cucurbit crops causing significant economic losses in ...

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6.3K Views.  University of Georgia. We developed a method to detect Phytophthora capsici zoospores in water sources using a filter paper DNA extraction method coupled with a loop-mediated isothermal amplification (LAMP) assay that can be analyzed in the field or in the lab.

Video: Detection of Phytophthora capsici in Irrigation Water using Loop-Mediated Isothermal Amplification

Colorimetric Paper-based Detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from Large Volumes of Agricultural Water

This protocol describes rapid colorimetric detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from large volumes (10 L) of agricultural waters. Here, water is filtered through sterile Modified Moore Swabs (MMS), which consist&#160;of a simple gauze filter enclosed in a plastic cartridge, to concentrate bacteria. Following filtration, non-selective or selective enrichments for the target bacteria are performed in the MMS. For colorimetric detection of the target bacteria, the enrichments are then assayed using paper-based analytical devices (&#181;PADs) embedded with bacteria-indicative substrates. Each substrate reacts with target-indicative bacterial enzymes, generating colored products that can be detected visually (qualitative detection) on the &#181;PAD. Alternatively, digital images of the reacted &#181;PADs can be generated with common scanning or photographic devices and analyzed using ImageJ software, allowing for more objective and standardized interpretation of results. Although the biochemical screening procedures are designed to identify the aforementioned bacterial pathogens, in some cases enzymes produced by background microbiota or the degradation of the colorimetric substrates may produce a false positive. Therefore, confirmation using a more discriminatory diagnostic is needed. Nonetheless, this bacterial concentration and detection platform is inexpensive, sensitive (0.1 CFU/ml detection limit), easy to perform, and rapid (concentration, enrichment, and detection are performed within approximately 24&#160;hr), justifying its use as an initial screening method for the microbiological quality of agricultural water.

Colorimetric Paper-based Detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from Large Volumes of Agricultural Water

A protocol involving integrated concentration, enrichment, and end-point colorimetric detection of foodborne pathogens in large volumes of agricultural water is presented here. Water is filtered through Modified Moore Swabs (MMS), enriched with selective or non-selective media, and detection is performed using paper-based analytical devices (&#181;PAD) imbedded with bacterial-indicative colorimetric substrates.

This protocol describes rapid colorimetric detection of Escherichia coli, Salmonella spp., and Listeria monocytogenes from large volumes (10 L) of ...

Laboratory-determined Phosphorus Flux from Lake Sediments as a Measure of Internal Phosphorus Loading

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) loading from lake sediments can account for a substantial portion of the total P load in eutrophic, and some mesotrophic, lakes. Laboratory determination of P release rates from sediment cores is one approach for determining the role of internal P loading and guiding management decisions. Two principal alternatives to experimental determination of sediment P release exist for estimating internal load: in situ measurements of changes in hypolimnetic P over time and P mass balance. The experimental approach using laboratory-based sediment incubations to quantify internal P load is a direct method, making it a valuable tool for lake management and restoration.
Laboratory incubations of sediment cores can help determine the relative importance of internal vs. external P loads, as well as be used to answer a variety of lake management and research questions. We illustrate the use of sediment core incubations to assess the effectiveness of an aluminum sulfate (alum) treatment for reducing sediment P release. Other research questions that can be investigated using this approach include the effects of sediment resuspension and bioturbation on P release.
The approach also has limitations. Assumptions must be made with respect to: extrapolating results from sediment cores to the entire lake; deciding over what time periods to measure nutrient release; and addressing possible core tube artifacts. A comprehensive dissolved oxygen monitoring strategy to assess temporal and spatial redox status in the lake provides greater confidence in annual P loads estimated from sediment core incubations.

Lake eutrophication is a water quality issue worldwide, making the need to identify and control nutrient sources critical. Laboratory determination of phosphorus release rates from sediment cores is a valuable approach for determining the role of internal phosphorus loading and guiding management decisions.

Eutrophication is a water quality issue in lakes worldwide, and there is a critical need to identify and control nutrient sources. Internal phosphorus (P) ...

EPA Method 1615. Measurement of Enterovirus and Norovirus Occurrence in Water by Culture and RT-qPCR. I. Collection of Virus Samples

EPA Method 1615 was developed with a goal of providing a standard method for measuring enteroviruses and noroviruses in environmental and drinking waters. The standardized sampling component of the method concentrates viruses that may be present in water by passage of a minimum specified volume of water through an electropositive cartridge filter. The minimum specified volumes for surface and finished/ground water are 300 L and 1,500 L, respectively. A major method limitation is the tendency for the filters to clog before meeting the sample volume requirement. Studies using two different, but equivalent, cartridge filter options showed that filter clogging was a problem with 10% of the samples with one of the filter types compared to 6% with the other filter type. Clogging tends to increase with turbidity, but cannot be predicted based on turbidity measurements only. From a cost standpoint one of the filter options is preferable over the other, but the water quality and experience with the water system to be sampled should be taken into consideration in making filter selections.

EPA Method 1615 uses an electropositive filter to concentrate enteroviruses and noroviruses in environmental and drinking waters. This manuscript describes the procedure for collecting samples for Method 1615 analyses.

EPA Method 1615 was developed with a goal of providing a standard method for measuring enteroviruses and noroviruses in environmental and drinking waters. ...

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated (WATER-PAM) Fluorometry

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be subsampled throughout the experiment1&#8211;7. Subsampling of large volumes can be problematic for several reasons: 1) it causes variation in the total volume and the surface area:volume ratio of the culture during the experiment; 2) pseudo-replication (i.e., replicate samples from the same treatment flask8) is often employed rather than true replicates (i.e., sampling from replicate treatments); 3) the duration of the experiment is limited by the total volume; and 4) axenic cultures or the usual bacterial microbiota are difficult to maintain during long-term experiments as contamination commonly occurs during subsampling.
The use of microtiter plates enables 1 ml culture volumes to be used for each replicate, with up to 48 separate treatments within a 12.65 x 8.5 x 2.2 cm plate, thereby decreasing the experimental volume and allowing for extensive replication without subsampling any treatment. Additionally, this technique can be modified to fit a variety of experimental formats including: bacterial-algal co-cultures, algal physiology tests, and toxin screening9&#8211;11. Individual wells with an alga, bacterium and/or co-cultures can be sampled for numerous laboratory procedures including, but not limited to: WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry, microscopy, bacterial colony forming unit (cfu) counts and flow cytometry. The combination of the microtiter plate format and WATER-PAM fluorometry allows for multiple rapid measurements of photochemical yield and other photochemical parameters with low variability between samples, high reproducibility and avoids the many pitfalls of subsampling a carboy or conical flask over the course of an experiment.

A Small Volume Bioassay to Assess Bacterial/Phytoplankton Co-culture Using WATER-Pulse-Amplitude-Modulated WATER-PAM Fluorometry

The goal of this procedure is to demonstrate the reproducibility and adaptability of using a microtiter plate format for microalgal screening. This rapid screen combines WATER-Pulse-Amplitude-Modulated (WATER-PAM) fluorometry to measure photosynthetic yield as an indicator of Photosystem II (PSII) health with small volume bacterial-algal co-cultures.

Conventional methods for experimental manipulation of microalgae have employed large volumes of culture (20 ml to 5 L), so that the culture can be ...

EPA Method 1615. Measurement of Enterovirus and Norovirus Occurrence in Water by Culture and RT-qPCR. Part III. Virus Detection by RT-qPCR

EPA Method 1615 measures enteroviruses and noroviruses present in environmental and drinking waters. This method was developed with the goal of having a standardized method for use in multiple analytical laboratories during monitoring period 3 of the Unregulated Contaminant Monitoring Rule. Herein we present the protocol for extraction of viral ribonucleic acid (RNA) from water sample concentrates and for quantitatively measuring enterovirus and norovirus concentrations using reverse transcription-quantitative PCR (RT-qPCR). Virus concentrations for the molecular assay are calculated in terms of genomic copies of viral RNA per liter based upon a standard curve. The method uses a number of quality controls to increase data quality and to reduce interlaboratory and intralaboratory variation. The method has been evaluated by examining virus recovery from ground and reagent grade waters seeded with poliovirus type 3 and murine norovirus as a surrogate for human noroviruses. Mean poliovirus recoveries were 20% in groundwaters and 44% in reagent grade water. Mean murine norovirus recoveries with the RT-qPCR assay were 30% in groundwaters and 4% in reagent grade water.

Here we present a procedure to quantify enterovirus and norovirus in environmental and drinking waters using reverse transcription-quantitative PCR. Mean virus recovery from groundwater with this standardized procedure from EPA Method 1615 was 20% for poliovirus and 30% for murine norovirus.

EPA Method 1615 measures enteroviruses and noroviruses present in environmental and drinking waters. This method was developed with the goal of having a ...

Screening for Endocrine Activity in Water Using Commercially-available In Vitro Transactivation Bioassays

In vitro transactivation bioassays have shown promise as water quality monitoring tools, however their adoption and widespread application has been hindered partly due to a lack of standardized methods and availability of robust, user-friendly technology. In this study, commercially available, division-arrested cell lines were employed to quantitatively screen for endocrine activity of chemicals present in water samples of interest to environmental quality professionals. A single, standardized protocol that included comprehensive quality assurance/quality control (QA/QC) checks was developed for Estrogen and Glucocorticoid Receptor activity (ER and GR, respectively) using a cell-based Fluorescence Resonance Energy Transfer (FRET) assay. Samples of treated municipal wastewater effluent and surface water from freshwater systems in California (USA), were extracted using solid phase extraction and analyzed for endocrine activity using the standardized protocol. Background and dose-response for endpoint-specific reference chemicals met QA/QC guidelines deemed necessary for reliable measurement. The bioassay screening response for surface water samples was largely not detectable. In contrast, effluent samples from secondary treatment plants had the highest measurable activity, with estimated bioassay equivalent concentrations (BEQs) up to 392 ng dexamethasone/L for GR and 17 ng 17&#946;-estradiol/L for ER. The bioassay response for a tertiary effluent sample was lower than that measured for secondary effluents, indicating a lower residual of endocrine active chemicals after advanced treatment. This protocol showed that in vitro transactivation bioassays that utilize commercially available, division-arrested cell &#34;kits&#34;, can be adapted to screen for endocrine activity in water.

Screening for Endocrine Activity in Water Using Commercially-available In Vitro Transactivation Bioassays

A protocol to screen for endocrine activity in organic extracts of water samples, including treated wastewater effluent and surface (receiving) water, was adapted using commercially available division-arrested ("freeze and thaw") in vitro transactivation bioassays.

In vitro transactivation bioassays have shown promise as water quality monitoring tools, however their adoption and widespread application has been ...

A Simple Planting Technique for Re-establishing Trees Where Frequent Inundation Occurs

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with herbaceous plants. This protocol describes a simple, cost-effective tree planting technique needed for restoring degraded Everglade tree islands. The design is patterned after a natural Everglades process that creates floating peat islands, which allows tree survival and growth in flooded conditions and often leads to the development of tree islands. Commercially available peat bags were used as the medium for the growth and establishment of potted native tree saplings. The pop-up configuration floated initially and provided additional elevation to minimize inundation, with a single native tree species sapling and a single tree fertilizer spike. During a 3 year study involving 105 pop-ups, most plants survived (80%) and many thrived. Determining whether this technique can establish trees on a degraded tree island will require longer studies and extensive field tests.

This protocol describes a simple, cost-effective tree planting technique for restoring degraded Everglades tree islands that experience inundation. The design creates an island (pop-up) which floats initially and adds elevation to promote tree survival and growth under flooded conditions.

Many of the Everglades tree islands have lost elevation over the past century and most of their trees have died such that they are now covered with ...

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol for performing in situ juvenile exposure tests within oligotrophic river catchments over one-month and three-month periods. Two methods (in both modifications) are presented to evaluate the juvenile growth and survival rate. The methods and modifications differ in value for the locality bioindication and each has its benefits as well as limitations. The sandy cage method works with a large set of individuals, but only some of the individuals are measured and the results are evaluated in bulk. In the mesh cage method, the individuals are kept and measured separately, but a low individual number is evaluated. The open water exposure modification is relatively easy to apply; it shows the juvenile growth potential of sites and can also be effective for water toxicity testing. The within-bed exposure modification needs a high workload but is closer to the conditions of a natural juvenile environment and it is better for reporting the real suitability of localities. On the other hand, more replications are needed in this modification due to its high-hyporheic environment variability.

Bioindication Testing of Stream Environment Suitability for Young Freshwater Pearl Mussels Using In Situ Exposure Methods

In situ bioindications enable determination of the suitability of an environment for endangered mussel species. We describe two methods based on the juvenile exposure of freshwater pearl mussels in cages to oligotrophic river habitats. Both methods are implemented in variants for open water and hyporheic water environments.

Knowledge of habitat suitability for freshwater mussels is an important step in the conservation of this endangered species group. We describe a protocol ...

Optimized Procedure for Determining the Adsorption of Phosphonates onto Granular Ferric Hydroxide using a Miniaturized Phosphorus Determination Method

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide (GFH), with little effort and high reliability. The phosphonate, e.g., nitrilotrimethylphosphonic acid (NTMP), is brought into contact with the GFH in a rotator in a solution buffered by an organic acid (e.g., acetic acid) or Good buffer (e.g., 2-(N-morpholino)ethanesulfonic acid) [MES] and N-cyclohexyl-2-hydroxyl-3-aminopropanesulfonic acid [CAPSO]) in a concentration of 10 mM for a specific time in 50 mL centrifuge tubes. Subsequently, after membrane filtration (0.45 &#181;m pore size), the total phosphorus (total P) concentration is measured using a specifically developed determination method (ISOmini). This method is a modification and simplification of the ISO 6878 method: a 4 mL sample is mixed with H2SO4 and K2S2O8 in a screw cap vial, heated to 148-150 &#176;C for 1 h and then mixed with NaOH, ascorbic acid and acidified molybdate with antimony(III) (final volume of 10 mL) to produce a blue complex. The color intensity, which is linearly proportional to the phosphorus concentration, is measured spectrophotometrically (880 nm). It is demonstrated that the buffer concentration used has no significant effect on the adsorption of phosphonate between pH 4 and 12. The buffers, therefore, do not compete with the phosphonate for adsorption sites. Furthermore, the relatively high concentration of the buffer requires a higher dosage concentration of oxidizing agent (K2S2O8) for digestion than that specified in ISO 6878, which, together with the NaOH dosage, is matched to each buffer. Despite the simplification, the ISOmini method does not lose any of its accuracy compared to the standardized method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric hydroxide, with little effort and high reliability. In a buffered solution, the phosphonate is brought into contact with the adsorbent using a rotator and then analyzed via a miniaturized phosphorus determination method.

This paper introduces a procedure to investigate the adsorption of phosphonates onto iron-containing filter materials, particularly granular ferric ...

A Loop-mediated Isothermal Amplification (LAMP) Assay for Rapid Identification of Bemisia tabaci

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many countries around the world. Severe yield losses are caused by direct feeding, and even more importantly, also by the transmission of more than 100 harmful plant pathogenic viruses. As for other invasive pests, increased international trade facilitates the dispersal of B. tabaci to areas beyond its native range. Inspections of plant import products at points of entry such as seaports and airports are, therefore, seen as an important prevention measure. However, this last line of defense against pest invasions is only effective if rapid identification methods for suspicious insect specimens are readily available. Because the morphological differentiation between the regulated B. tabaci and close relatives without quarantine status is difficult for non-taxonomists, a rapid molecular identification assay based on the loop-mediated isothermal amplification (LAMP) technology has been developed. This publication reports the detailed protocol of the novel assay describing rapid DNA extraction, set-up of the LAMP reaction, as well as interpretation of its read-out, which allows identifying B. tabaci specimens within one hour. Compared to existing protocols for the detection of specific B. tabaci biotypes, the developed method targets the whole B. tabaci species complex in one assay. Moreover the assay is designed to be applied on-site by plant health inspectors with minimal laboratory training directly at points of entry. Thorough validation performed under laboratory and on-site conditions demonstrates that the reported LAMP assay is a rapid and reliable identification tool, improving the management of B. tabaci.

A Loop-mediated Isothermal Amplification LAMP Assay for Rapid Identification of Bemisia tabaci

This paper reports the protocol for a rapid identification assay for Bemisia tabaci based on loop-mediated isothermal amplification (LAMP) technology. The protocol requires minimal laboratory training and can, therefore, be implemented on-site at points of entry for plant imports such as seaports and airports.

The whitefly Bemisia tabaci (Gennadius) is an invasive pest of considerable importance, affecting the production of vegetable and ornamental crops in many ...