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

1. On-site detection of Phytophthora capsici from irrigation water using portable loop-mediated isothermal amplification

1. Setting up the pump and filter
   1. 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.
   2. 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 µm.
      NOTE: The filter paper must fit to the edges of the Buchner funnel so that minimal water will flow around the filter paper.
2. Water sampling and filtering
   1. Take water samples from the targeted source. Water may have small amounts of debris but not significant sediment or soil.
   2. Pour up to 1,000 mL (1 Liter) 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.
   3. 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 µL 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.
   4. 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 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 buffer.
3. Magnetic bead-based extraction of DNA from filter paper
   1. To the 1.5 mL tube (which now contains approximately 200-300 µL of extraction buffer), add 20 µL of proteinase K and 10 µL of 10 ng/µL RNase.
   2. Incubate at room temperature for 15 min, vortex or shake the tube every 3 min.
   3. Add 500 µL of magnetic beads with the binding buffer to the sample and mix well by shaking. Then incubate for 5 min at room temperature.
   4. 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.
   5. Remove the tube from the magnetic separator. Add 500 µL 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.
   6. Repeat step 1.3.5 with 500 µL of Wash Buffer 2.
   7. Repeat step 1.3.5 with 500 µL of 80% ethanol.
   8. Airdry the magnetic bead pellet for 15 min at room temperature (18-27 °C) with the lid open. If temperatures do not permit, incubate in a gloved hand with the cap open for 15 min.
   9. Remove the tube from the magnetic separator. Add 50 µL of elution buffer and re-suspend the beads by pipetting up and down for 1 min.
   10. 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 °C freezer.
4. Application of newly developed LAMP assay
   1. Prepare LAMP primer mix using 0.2 µM of each F3 and B3 primer, 0.8 µM of each Loop-F and Loop-B primer, and 1.6 µM of each FIP and BIP primer (Table 2).
   2. Add the following LAMP solution to a single PCR tube, or to each individual tube in the 8 tube strip: 2.5 µL of primer mix (step 1.4.1), 12.5 µL of LAVA LAMP master mix, 1 µL of extracted DNA, and 9 µL of ddH₂O. The total volume is 25 µL.
      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.
   3. 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 µL of extracted DNA for 1 µL of the positive control or ddH₂O.
   4. Set the samples into a heat block (or portable amplification instrument) set to 64 °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 sample^24,25. Results were compared in Table 2. If concentration of DNA can be quantified, use between 1-10 ng genomic DNA.
5. Visualization of results
   1. If performed in a laboratory setting, view the amplification products by loading 5 µL of each sample into a 1% agarose gel, running them in a gel electrophoresis machine, and imaging them in a UV imaging machine.
   2. If a colorimetric dye (e.g., Warmstart) was used, view the color change to determine results as positive or negative.
   3. If a portable amplification instrument (e.g., Genie III) was used, view the amplification graph on the screen to determine the results.
6. 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.
   1. Add 1 µL of each DNA extraction to individual tubes containing the following components: 12.5 µL of Green PCR master mix, 9.5 µL of ddH₂O, and 1 µL of forward and reverse primers (Table 2).
   2. Spin down each sample using a microcentrifuge and place tubes into a thermal cycler.
   3. Use the following thermal cycler settings in accordance to the previous publications: 94 °C for 5 min, 30 cycles of denaturation at 94 °C for 30 s, annealing at 54 °C for 30 s, extension at 72 °C for 1 min, and final extension at 72 °C for 10 min.
   4. 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.
7. 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.
   1. First, obtain a healthy eggplant fruit (a susceptible host for P. capsici) and surface sterilize by washing the fruit surface with 70% isopropyl alcohol.
   2. 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.
   3. 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 °C for 5 days.
   4. View plates under a compound microscope for traditional morphological identification at 4 days after isolation.

2. Determining the detection limit of zoospore concentration

1. Making zoospore suspension
   1. 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 °C. Multiple plates can be used to obtain a larger amount of zoospore suspension.
   2. Incubate the plates under continuous light at room temperature for 3 days to stimulate sporulation.
   3. Flood the plates by adding 15 mL of ddH₂O to each plate and place them in a 4 °C fridge for 25 min. Then return to room temperature for 30 min.
   4. Agitate plates to dislodge zoospores and pipette the solution into a single 50 mL tube from all the plates.
   5. To obtain an accurate estimate of zoospore concentration, add 10 µL of the spore suspension onto a hemocytometer and observe under a microscope to count zoospores and estimate the average concentration.
2. Serial dilution
   1. 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.
   2. 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.
3. Detection of zoospore concentration limit
   1. 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.
4. 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.
   1. If colorimetric dye (e.g., Warmstart) is used, use the following the solution: 2.5 µL of primer mix, 12.5 µL of colorimetric dye, 0.5 µL of Green fluorescent dye, 1 µL of extracted DNA, and 8.5 µL of ddH₂O. The total volume is 25 µL.
   2. When using the portable amplification instrument with LAVALAMP mastermix, have an initial step of 95 °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.
   3. 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.
   4. 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.
   5. 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.
5. Detection limit determination and comparison with conventional PCR method
   1. Use DNA extracted in steps in section 1.3 to compare the detection level of conventional PCR with that of the LAMP assay.
   2. Add 1 µL of DNA to a PCR tube that contained 1 µL of both forward and reverse PCR primers (Table 2), 12.5 µL of Green PCR Mastermix, and 9.5 µL of ddH₂O for a total of 25 µL.
   3. Amplify samples in a thermal cycler using the following conditions: 94 °C for 5 min, 30 cycles of denaturation at 94 °C for 30 s, annealing at 54 °C for 30 s, extension at 72 °C for 1 min, and final extension at 72 °C for 10 min.
   4. 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.