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

Owen Hudson1, Sumyya Waliullah1, Justin Hand2, Romina Gazis-Seregina3, Fulya Baysal-Gurel4, Md Emran Ali1
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Hudson, O., Waliullah, S., Hand, J., Gazis-Seregina, R., Baysal-Gurel, F., Ali, M. E. Detection of Phytophthora capsici in Irrigation Water using Loop-Mediated Isothermal Amplification. J. Vis. Exp. (160), e61478, doi:10.3791/61478 (2020).

Abstract

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 µm) method that captures the pathogen’s spores (zoospores) and is later used to amplify the pathogen’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.

Introduction

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/µL); 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/µL 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.

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Protocol

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 sample24,25. 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.

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Representative Results

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 (0.1–0.5 µM each); LF, LB (0.5–1.0 µM each) and FIP, BIP (0.8–2.4 µM each)], durations (30–70 min), and temperatures (55–70 °C). The final LAMP primer mix used in this study was: 0.2 µM of each F3 and B3 primer, 0.8 µM of each Loop-F and Loop-B primer, 1.6 µM of each FIP and BIP primer. Optimization of reaction temperature was performed in the portable amplification instrument (e.g., Genie III) by determining what temperature performed the fastest reaction with no additional negative amplification. The optimal temperature was confirmed to be 64 °C (data not shown). The optimal time for running the assay at 64 °C was 45 min, as the lowest concentrations that were positive for detection (1.2 x 102 spores/mL) still amplified by 40 min, while higher concentrations amplified at 20 min (Figure 2D). The amplified LAMP products were further observed on 1% agarose gel stained with a nucleic acid stain to confirm amplification. All reactions were repeated at least three times.

Isolates of P. capsici were taken from Tennessee, Florida, and Georgia and were submitted to the same protocol described in the methods. All samples of P. capsici isolates were amplified successfully in all runs of the assay (Figure 3).

Detection and sensitivity testing of Phytophthora capsici in irrigation water using portable LAMP assay
We standardized this filter paper-based LAMP method under laboratory conditions using a serial dilution of P. capsici spore suspensions (Figure 1). Serial dilutions were made from a P. capsici spore suspension starting at 4.8 x 104 zoospores/mL and run with the LAMP assay in triplicate. Spore concentrations are shown rather than DNA concentration due to the method involved for DNA extraction. CTAB DNA extraction of the highest spore concentration was 4.5 ng/µL measured by Nanodrop, and the magnetic bead DNA extraction protocol yielded 3.8 ng/µL26. The newly designed LAMP primer set could detect a concentration as low as 1.2 x 102 spores/mL (Figure 2B, 2C, & 2D) with all methods of extraction. The sensitivity shown in the graph of amplification on the portable amplification instrument was identical to that shown in the UV image with no additional sensitivity. The same serial dilution was run in a LAMP reaction using the colorimetric dye to determine the level of sensitivity to the naked eye for field detection. The lowest observable concentration was 4.8 x 103 spores/mL (Figure 2C).

To evaluate the ability of this assay using real-world samples, water samples were collected from seven ponds used for commercial vegetable production in Tift County, Georgia (Table 1, Figure 5, and Supplementary Figure 1). Out of the 7 ponds, 3 showed positive LAMP results (P1, P4, and P6) (Figure 6A, 6B, & 6C). These results suggest that the portable filter paper-based LAMP method could be very useful for detection of the pathogen even with a low zoospore concentration. This demonstrates the applicability of LAMP as a more sensitive detection assay than PCR for screening irrigation water contamination by P. capsici.

Comparative analysis of different methods: Traditional baiting, conventional PCR, and the portable LAMP-based assay
In order to compare the detection sensitivity of LAMP with conventional PCR, the DNA extracted from the serial dilution of spore suspensions were run in a PCR reaction. Results showed that conventional PCR was 40x less sensitive than LAMP, only able to detect a zoospore concentration as low as 4.8 x 103 spores/mL (Figure 2A). Additionally, the DNA samples obtained from filtered irrigation pond water were tested using conventional PCR, and only one of the three positive samples (P4) was successfully amplified as expected band size plus significant contaminates resulting in some smearing and unspecific bands (Figure 6D). Table 3 displays the differences between detection methods using such variables as time, cost, sensitivity, and preparation required. LAMP was the least expensive method among these three methods and it was also the fastest, ranging from 30-60 min for amplification (DNA extraction excluded). Conventional PCR ranged from 120-180 min for amplification (DNA extraction excluded).

Finally, to determine the specificity of the primers, samples of closely related oomycete pathogens (Phytophthora sansomeana, Phytophthora sojae, Phytophthora cinnamomi, Phytophthora palmivora, Pythium ultimum var. ultimum, Phytopythium vexans, Phytopythium helicoides, and Pythium aphanidermatum) were obtained, DNA was extracted using the same protocol for consistency and evaluated by the new LAMP assay with a positive and negative control (Figure 4) to determine specificity of the primers. All non-target samples were negative using the optimized 64 °C for 45 min. This was observed on the real-time amplification graph and imaged on a 1% agarose gel under UV light.

Figure 1
Figure 1: Diagram showing the different steps involved in Phytophthora capsici from a serial dilution of a concentrated spore suspension under laboratory conditions. Please click here to view a larger version of this figure.

Figure 2
Figure 2: Laboratory optimization of the limit for detection of Phytophthora capsici(A) Conventional PCR assay was carried out using specific P. capsici primers on serial dilution factors and visualized on 1% agarose gel. 1, Ladder; 2-7, 4.8 x 104, 4.8 x 103, 4.8 x 102, 2.4 x 102, 1.2 x 102 spores/mL, respectively and 7, negative water control. (B) LAMP assay serial dilution factors visualized on 1% agarose gel. 1-6, a decreasing spore concentration: 4.8 x 104, 4.8 x 103, 4.8 x 102, 2.4 x 102, 1.2 x 102 spores/mL, and 7, negative water control. (C) LAMP results visualized using the colorimetric dye. 1-6, a decreasing spore concentration: 4.8 x 104, 4.8 x 103, 4.8 x 102, 2.4 x 102, 1.2 x 102 spores/mL, and 7, negative water control. (D) LAMP results visualized on the amplification graph. Red = 4.8 x 104, Dark blue = 4.8 x 103, Orange = 4.8 x 102, Light blue = 2.4 x 102, Green = 1.2 x 102 spores/mL, Pink = Negative control (other Phytophthora species), Yellow = ddH2O. (E) A standard curve showing quantification of the values shown in the real-time results. Ln (Spore count) is shown on the X-axis, and minutes to amplification on the Y-axis. (F) LAMP assay with published primer (Dong et al. 2015) on serial dilution factors visualized on 1% agarose gel. 1-6, a decreasing spore concentration: 4.8 x 104, 4.8 x 103, 4.8 x 102, 2.4 x 102, 1.2 x 102 spores/mL, and 7, negative water control. Please click here to view a larger version of this figure.

Figure 3
Figure 3: Amplification of P. capsici DNA from various locations. (A) LAMP results visualized on 1% agarose gel. 1, PC_TN1; 2, PC_TN2; 3, PC_FL1; 4,PC_FL2; 5, PC_GA1; 6, PC_GA2; N, Negative control. Samples 1 and 2 were isolated from TN; samples 3 and 4 isolated from FL; samples 5 and 6 were isolated from GA. (B) Results visualized using the colorimetric dye. 1, PC_TN1; 2, PCTN2; 3, PC_FL1; 4,PC_FL2; 5, PC_GA1; 6, PC_GA2. (C) Results visualized on the amplification graph. Red, PC_TN1; Orange, PCTN2; Yellow, PC_FL1; Green, PC_FL2; Dark blue, PC_GA1; Light blue, PC_GA2; Pink, negative control. Please click here to view a larger version of this figure.

Figure 4
Figure 4: Specificity determination of LAMP assay using DNA from non-target species P. capsici. (A) LAMP assay reaction with related non-target species on agarose gel and visualized on 1% agarose gel. L, Ladder; 1, Phytophthora sansomeana; 2, Phytophthora sojae; 3, Phytophthora cinnamomi; 4, Phytophthora palmivora; 5, Pythium ultimum var. ultimum; 6, Phytopythium vexans; 7, Negative control; 8, Phytophthora capsica. (B) LAMP results visualized using the colorimetric dye. 1, Phytophthora sansomeana; 2, Phytophthora sojae; 3, Phytophthora cinnamomi; 4, Phytophthora palmivora; 5, Pythium ultimum var. ultimum; 6, Phytopythium vexans; 7, Negative control; 8, Phytophthora capsici (C) LAMP results visualized on the amplification graph. Red = Phytophthora capcisi, all other non-target species samples were not amplified. Please click here to view a larger version of this figure.

Figure 5
Figure 5: Pictures showing the sampling and processing of recycled water for the detection of Phytophthora capsici in the field. Please click here to view a larger version of this figure.

Figure 6
Figure 6: Results from on-site detection of Phytophthora capsici in irrigation water sources. (A) Agarose gel showing results from the LAMP amplification of tested water from seven farm in South Georgia. Sample names from left to right: P1, P2, P3, P4, P5, P6, P7, Negative control, N. (B) LAMP results visualized using warmstart colorimetric dye of field samples: P1, P2, P3, P4, P5, P6, P7, Negative control, N. (C) Results from LAMP amplification of field samples using graph. Red: P1, Green: P2, Purple: P3, Yellow: P4, Blue: P5, Orange: P6, Pink: P7, Negative control, N. (D) Agarose gel showing conventional PCR results of the amplification using specific primers PC-1/PC-2 (note than only one site tested positive in comparison to three in LAMP). Please click here to view a larger version of this figure.

Pond name County, State Target crops for irrigation PCR Detection LAMP Detection History of Disease (Y/N)
P1 Tift, GA Vegetables ­ + N
P2 Tift, GA Vegetables - - N
P3 Tift, GA Vegetables - - N
P4 Tift, GA Vegetables + + N
P5 Tift, GA Vegetables - - N
P6 Tift, GA Vegetables - + N
P7 Tift, GA Vegetables - - N

Table 1: Detection of irrigation water from Southern GA.

Primer type Primer name Sequence 5’-3’ Source
LAMP PCA3-F3 TGTGTGTGTGTTCGATCACA This study
PCA3-B3 TTTTTGCGTGCGTCCAGA This study
PCA3-FIP GACACCAAGCACTCGTACTOGTTTTTACAATTGTGCAGAGGGAGGA This study
PCA3-BIP AGAACGAGTATTCGGCGGCGTTTTGAAAAAGGACCACCCCCG This study
PCA3-LF TGTCGAATGGATTTGCGATCTT This study
PCA3-LB ATACGCAGGTCATTTGACTGAC This study
PCR PC-1 GTCTTGTACCCTATCATGGCG Zhang et al., 2006
PC-2 CGCCACAGCAGGAAAAGCATT Zhang et al., 2006

Table 2: Primers used in this study.

Parameters Traditional Conventional PCR LAMP
Sensitivity NA 4.8 X 102 spores/ml 1.2 X 102 spores/ml
Time 2 weeks or longer 2-3 hours (not including DNA extraction) 30 mins - 1 hour (not including DNA extraction)
Preparation • Media creation
• Plating and isolation and
• Designing a trap
• Spore collection using Filter paper
• DNA extraction and
• PCR assay
• Spore collection using Filter paper
• DNA extraction and
• LAMP assay
Materials • Autoclave
• Media and plates
• Incubation room
• Flow hood
• Eggplant
• Milk Crate
• Thermal cycler
• Agar gel
• Gel Doc

• Heat block or
• Genie III
Cost $5.00 per trap $0.60 per reaction $0.75 per reaction

Table 3: Comparison of methods for detection of P. capsici.

Supplementary Figure 1: Location of Tift County, GA and samples of irrigation water taken from various locations from within the state. Positive samples were shown as red dots. Please click here to download this figure.

Supplementary Figure 2: Design of LAMP primer set. Arrows show direction of how primers are read. Please click here to download this figure.

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Discussion

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 for the pathogen by using susceptible host tissue (e.g., fruit, leaves) suspended in the pond and wait for an infection to take place, then remove the fruit/leaves and confirm the diagnosis with microscopy or molecular methods13,14. These methods are limiting due to the amount of time required to run the detection test (2 weeks or longer), and the labor and equipment required. Additionally, extensive experience and knowledge in visual diagnosis, pathogen morphology, and taxonomy are required for accurate results. Molecular techniques such as PCR, qPCR, and DNA hybridization require significantly less time (3-4 h) than the traditional methods of detection; however, they require expensive equipment and a laboratory setting. Additionally, these techniques do not allow for the processing of large volumes of water. Serological assays too, fall short in their detection ability due to non-target positive reactions, and no species-specific assays for Phytophthora species have been developed. Loop-mediated isothermal amplification (LAMP) has recently been used as an on-site diagnosis technique for rapid and sensitive detection of multiple pathogens as the assay only requires a single temperature rather than a thermal cycler28,29. LAMP can be run in the field using a colorimetric dye for visual confirmation or using a real-time amplification machine for results in less than one hour30.

The goal of this experiment was to develop a rapid and sensitive method to detect the presence of Phytophthora capsici in water sources either on-site or in a laboratory. To increase the speed of detection and to combat the limitations of the previously mentioned methods for detecting P. capsici in irrigation water, we designed a method using filter paper to capture the spores and extract their DNA from a larger volume of water. After spores were captured using the filter paper technique and DNA was extracted, the presence of the pathogen was confirmed based on a newly designed LAMP primer set specific to P. capsici. Detection sensitivity and specificity was compared using LAMP and PCR. In all 3 replications and with all of the zoospore concentrations, LAMP was a quicker and more sensitive detection method (Table 3). This method is not limited by having a small sample volume as traditional methods, as this method can be test up to 1 L of water at a single time, increasing the chances of pathogen detection. It was noted in testing that pouring irrigation water slowly through the Buchner funnel at a speed of no more than 40 mL per second increased the spore capture ability of the filter paper.

To validate the detection protocol, water samples from the field where P. capsici was suspected to be present were also taken (Supplementary Figure 1) to test the designed method with a practical scenario. Out of the 7 farms tested, 3 were positive for the presence of P. capsici using the LAMP assay (Figure 6A-6C) while only one farm was positive when using the conventional PCR assay (Figure 6D), showing LAMP as a more sensitive assay for this method of testing irrigation water. Although, this filter based LAMP assay could detect DNA from a concentration as low as 1.2 x 102 zoospores/mL which was significantly less than the original sensitivity of this assay (0.01 ng genomic DNA equivalent to ~5 spores) with unfiltered zoospore suspension. The previous LAMP assay by Dong et. al. (2015)20 was run on the same serial dilution and the level of sensitivity was the same (Figure 2F). The level of detection for a PCR assay with unfiltered spore solution have also shown a higher level of detection (equivalent ~10 spores) which was very similar to the previous PCR based findings10. The spore detection limit of the traditional baiting method of detection was not checked as a single spore could cause infection depending on its individual ability. The decreased sensitivity found in both LAMP and PCR assays is likely due to some spores flowing through or around the filter paper, or once attached to the filter paper, unable to be extracted with 100% efficiency. Nevertheless, this new LAMP and filter system can process and analyze a much higher volume of water than previous methods and confers a higher level of specificity and speed for in-field detection.

Of the extraction methods used, magnetic bead-based extraction was the most rapid and did not require the use of external machines such as a bead beater or centrifuge making it useful for in-field extractions and compliments the portable feature of the LAMP assay. The CTAB based method yielded the highest concentration of DNA but took the longest amount of time, while the commercial plant DNA extraction kit (e.g., DNeasy) was second in both time required and DNA concentration acquired.

With both CTAB and the commercial plant DNA extraction kit, during the homogenization of filter paper it was noted that homogenization was more successful and yielded a higher concentration if the CTAB solution (or extraction buffer) was added to the tube with the pieces of filter paper before bead beating or hand homogenization commenced. Bead beating was done 3 times for one minute each, but vortexing and agitating the tube in between each round was necessary for complete homogenization in all extraction methods. Importantly, the filter paper is subject to getting stuck on the sides or bottom of the tube, so it is crucial to make sure the filter paper is off the walls so that it gets homogenized.

The total time required for amplification is 90-120 min and can be easily done in the field for on-site diagnosis. This filter method is also designed to filter significant amounts of water to increase the possible chances for the detection of the pathogen. This method is also applicable to many pathogens that can accumulate in a water source, particularly genera such as: Pythium, Phytophthora, Fusarium, and bacteria; the only change required will be the development of an equally specific LAMP primer set for the targeted pathogen30.

A significant output of this work is the development of a highly sensitive and rapid filter paper-based LAMP assay for the detection of P. capsici in irrigation water sources. We expect that this study will lead to an increase in awareness of contamination of recycled irrigation water, eventually improving management of Phytophthora associated diseases, and consequently reduce production costs and increase crop yield. Such information is highly needed to improve vegetable production sustainability and enhance profitability of vegetable productions.

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Disclosures

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

Acknowledgments

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.

Materials

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

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References

  1. Hong, C., Moorman, G. J. Plant pathogens in irrigation water: challenges and opportunities. Critical Reviews in Plant Sciences. 24, (3), 189-208 (2005).
  2. Malkawi, H. I., Mohammad, M. J. Physiology, Genetics, Morphology, & Microorganisms, E. o. Survival and accumulation of microorganisms in soils irrigated with secondary treated wastewater. Journal of Basic Microbiology. 43, (1), 47-55 (2003).
  3. Bush, E. A., Hong, C., Stromberg, E. L. Fluctuations of Phytophthora and Pythium spp. in components of a recycling irrigation system. Plant Disease. 87, (12), 1500-1506 (2003).
  4. Ghimire, S. R., et al. Distribution and diversity of Phytophthora species in nursery irrigation reservoir adopting water recycling system during winter months. Journal of Phytopathology. 159, (11-12), 713-719 (2011).
  5. Hausbeck, M. K., Lamour, K. H. Phytophthora capsici on vegetable crops: research progress and management challenges. Plant Disease. 88, (12), 1292-1303 (2004).
  6. Gevens, A., Donahoo, R., Lamour, K., Hausbeck, M. Characterization of Phytophthora capsici from Michigan surface irrigation water. Phytopathology. 97, (4), 421-428 (2007).
  7. Thomson, S., Allen, R. Occurrence of Phytophthora species and other potential plant pathogens in recycled irrigation water. Plant Disease Reporter. 58, (10), 945-949 (1974).
  8. Lamour, K. H., Stam, R., Jupe, J., Huitema, E. The oomycete broad-host-range pathogen Phytophthora capsici. Journal of Molecular Plant Pathology. 13, (4), 329-337 (2012).
  9. Sanogo, S., Ji, P. Water management in relation to control of Phytophthora capsici in vegetable crops. Agricultural Water Management. 129, 113-119 (2013).
  10. Zhang, Z., Li, Y., Fan, H., Wang, Y., Zheng, X. Molecular detection of Phytophthora capsici in infected plant tissues, soil and water. Plant Pathology. 55, (6), 770-775 (2006).
  11. Trout, C., Ristaino, J., Madritch, M., Wangsomboondee, T. Rapid detection of Phytophthora infestans in late blight-infected potato and tomato using PCR. Plant Disease. 81, (9), 1042-1048 (1997).
  12. Sankaran, S., Mishra, A., Ehsani, R., Davis, C. A review of advanced techniques for detecting plant diseases. Commputers and Electronics in Agriculture. 72, (1), 1-13 (2010).
  13. Wang, Z., et al. Development of an improved isolation approach and simple sequence repeat markers to characterize Phytophthora capsici populations in irrigation ponds in southern Georgia. Applied and Environmental Microbiology. 75, (17), 5467-5473 (2009).
  14. Ali-Shtayeh, M., MacDonald, J. Occurrence of Phytophthora species in irrigation water in the Nablus area (West Bank of Jordan). Phytopathologia Mediterranea. 143-150 (1991).
  15. Pringsh, P. Comparison of serological and culture plate methods for detecting species of Phytophthora, Pythium, and Rhizoctonia in ornamental plants. Plant Disease. 74, (9), 655 (1990).
  16. Stewart-Wade, S. M. Plant pathogens in recycled irrigation water in commercial plant nurseries and greenhouses: their detection and management. Irrigation Science. 29, (4), 267-297 (2011).
  17. Aragaki, M., Uchida, J. Y. Morphological distinctions between Phytophthora capsici and P. tropicalis sp. nov. Mycologia. 93, (1), 137-145 (2001).
  18. Tomlinson, J., Boonham, N. Potential of LAMP for detection of plant pathogens. CAB Reviews Perspectives in Agriculture Veterinary Science Nutrition and Natural Resources. 3, (066), 1-7 (2008).
  19. Li, P., et al. A PCR-based assay for distinguishing between A1 and A2 mating types of Phytophthora capsici. Journal of the American Society for Horticultural Science. 142, (4), 260-264 (2017).
  20. Dong, Z., et al. Loop-mediated isothermal amplification assay for sensitive and rapid detection of Phytophthora capsici. Canadian Journal of Plant Pathology. 37, (4), 485-494 (2015).
  21. Khan, M., et al. Comparative evaluation of the LAMP assay and PCR-based assays for the rapid detection of Alternaria solani. Frontiers in Microbiology. 9, 2089 (2018).
  22. Sowmya, N., Thakur, M., Manonmani, H. K. Rapid and simple DNA extraction method for the detection of enterotoxigenic Staphylococcus aureus directly from food samples: comparison of PCR and LAMP methods. Journal of Applied Microbiology. 113, (1), 106-113 (2012).
  23. Waliullah, S., et al. Comparative analysis of different molecular and serological methods for detection of Xylella fastidiosa in blueberry. PLOS ONE. 14, (9), 0221903 (2019).
  24. Böhm, J., et al. Real-time quantitative PCR: DNA determination in isolated spores of the mycorrhizal fungus Glomus mosseae and monitoring of Phytophthora infestans and Phytophthora citricola in their respective host plants. Journal of Phytopathology. 147, 409-416 (1999).
  25. Klimczak, L., Prell, H. J. C. Isolation and characterization of mitochondrial DNA of the oomycetous fungus Phytophthora infestans. Current Genetics. 8, (4), 323-326 (1984).
  26. Ghimire, S. R., et al. Detection of Phytophthora species in a run-off water retention basin at a commercial nursery in plant hardiness zones 7 b of Virginia in winter. Phytopathology. 96, (6), (2006).
  27. Feng, W., Hieno, A., Kusunoki, M., Suga, H., Kageyama, K. J. P. LAMP detection of four plant-pathogenic oomycetes and its application in lettuce fields. Plant Disease. 103, (2), 298-307 (2019).
  28. Aglietti, C., et al. Real-time loop-mediated isothermal amplification: an early-warning tool for quarantine plant pathogen detection. AMB Express. 9, (1), 50 (2019).
  29. Almasi, M. A. Development of a colorimetric loop-mediated isothermal amplification assay for the visual detection of Fusarium oxysporum f. sp. melonis. Horticultural Plant Journal. 5, (3), 129-136 (2019).
  30. Gill, D. J. Pathogenic Pythium from irrigation ponds. Plant Disease Reporter. 54, (12), 1077-1079 (1970).

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