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Early Detection of the Root-Knot Nematode Meloidogyne hapla Through Developing a Robust Quantitative PCR Approach Compliant with the Minimum Information for Publication of Quantitative Real-Time PCR Experiments Guidelines

    Affiliations
    Authors and Affiliations
    • Johannes Tavoillot1 †
    • Thierry Mateille1
    • Nadine Ali2
    • Anne-Marie Chappé3
    • Jean-François Martin1
    1. 1Biology Center for Population Management (CBGP), Université de Montpellier, Center for International Cooperation in Agricultural Research for Development (CIRAD), National Institute for Agricultural Research (INRAE), Institut Agro, Institute of Research for Development (IRD) Montpellier, 34988 Montpellier, France
    2. 2Department of Plant Protection, Tishreen University, Tishreen, Syria
    3. 3Nematology Unit, Anses, French Agency for Food, Environmental and Occupational Health and Safety/Plant Health Laboratory, 35653 Le Rheu, France
    Published Online:9 Nov 2021https://doi.org/10.1094/PDIS-11-20-2408-RE

    Abstract

    Root-knot nematodes (RKNs) are major threats to crops through attacking the roots, which induces an abnormal development of the plant. Meloidogyne hapla is of particular concern, as it is currently expanding its distribution area and displays a wide host range. Effective plant protection against this RKN requires early detection, as even a single individual can cause severe economic losses on susceptible crops. Molecular tools are of particular value for this purpose, and among them, quantitative PCR (qPCR) presents many advantages (i.e., sensitivity, specificity, and rapidity of diagnosis at a reduced cost). Although a few studies have already been proposed for detecting M. hapla through this technique, they lack experimental details and performance testing, suffer from low taxonomic resolution, and/or require expensive hydrolysis probes. Here, we propose a qPCR detection method that uses SYBR Green with developed primers amplifying a fragment of the cytochrome oxidase I mitochondrial region. The method was developed and evaluated following the minimum information for publication of quantitative real-time PCR experiments (MIQE) guidelines to ensure its quality (i.e., sensitivity, specificity, repeatability, reproducibility, and robustness). The results demonstrate that the newly developed method fulfills its goals, as it proved specific to M. hapla and allowed for a reproductible detection level as low as 1.25 equivalent of a juvenile individual. All criteria associated with the MIQE guidelines were also met, so the method is of general use for the reliable early detection of M. hapla.

    The root-knot nematodes (RKNs) Meloidogyne spp. (Tylenchida, Meloidogynidae) are soilborne pests of major crops. These obligate endoparasites decrease plant growth and reduce crop yields. The total global yield loss is estimated at 10% (Raaijmakers et al. 2009), resulting in a worldwide annual loss of U.S. $100 billion (Bird et al. 2009) and local impact affecting agricultural production (40% of farms in southern France are affected; Djian-Caporalino 2012). Among the nearly 100 Meloidogyne species described (Hunt and Handoo 2009), three common tropical species (Meloidogyne javanica, Meloidogyne arenaria, and Meloidogyne incognita) and one temperate species (Meloidogyne hapla; Moens et al. 2009) are responsible for most crop damage. Although classified as a cryophilic species (Lyons et al. 1975), M. hapla Chitwood is cosmopolitan, including temperate, cold northern countries (Hunt and Handoo 2009; Orton 1974) and the Mediterranean basin (Dabaj and Jenser 1987). It is found equally in natural ecosystems (Ali et al. 2016) and vegetable shelters (Djian-Caporalino 2012). Severe infections and damage by M. hapla are more and more frequent, especially in organic soils (Abawi et al. 1997). The spread in vegetable crops was ascribed to the wide use of Mi gene-resistant cultivars that do not control M. hapla (Liu and Williamson 2006). M. hapla has a wide host range, including >550 crops and weeds (Jepson 1987). Negative economic impact by M. hapla occurs mostly on vegetables with a damage threshold density of less than one egg per square centimeter of soil, especially leafy greens (Widmer et al. 1999), which represent a basic component in the vegetable cropping systems in southern Europe. In this context, an early diagnosis is key in controlling the pest. However, symptoms on plants (stunted growth, wilting, leaf yellowing, and root galling) are observed late in the crop cycle and are therefore poor indicators to prevent its occurrence.

    RKNs are difficult to characterize because they are embedded in the soil and plant roots. Moreover, the specific identification of Meloidogyne species has always been a challenge, especially because of cryptic species complexes, sexual dimorphism in some cases, and polyploidy (Blok and Powers 2009; Janati et al. 1982). The most common methods are based on morphological criteria (e.g., perineal pattern of the females, stylet knob shape of the males, stylet length, and distance of the dorsal gland orifice from the stylet base; Hunt and Handoo 2009). These diagnostic features are difficult to recover because (i) Meloidogyne males are often rare (most species are parthenogenetic); (ii) females are embedded in roots, and the distortion of their shape prevents reproducible morphometrics; and (iii) juveniles do not provide enough discriminating characteristics. Species among Meloidogyne are very similar in shape (De Oliveira et al. 2011), and their morphological diagnostic requires much time and systematics expertise.

    Alternative identification methods of RKNs have been established using biochemical markers (Bergé 1975). They are based on the electrophoretic separation of isoenzymes from individual samples of females (e.g., esterases, malate dehydrogenases, glutamate oxaloacetate transaminases, superoxide dismutases) with specific molecular weights (Esbenshade and Triantaphyllou 1985). The electrophoretic migration of the isoenzyme bands is evaluated by a migration ratio (Rm) and compared with that of a control population (usually M. javanica). This method is widely used routinely, although it is not reliable when multiple species are mixed in a sample. Moreover, it can be used only with females because of the high protein concentration required (Esbenshade and Triantaphyllou 1990), which calls for long and tedious individual extraction of the females outside the roots by hand.

    Molecular methods became essential to identify nematodes especially in the case of cryptic species (Gamel et al. 2014). These methods are reliable, fast, accurate, and relatively simple to implement. They are based on the detection of DNA polymorphic markers using the PCR technique, such as PCR restriction fragment length polymorphism (Curran et al. 1986), microsatellite DNA markers (Castagnone-Sereno et al. 1995), and random amplified polymorphic DNA (RAPD) genotyping (Randig et al. 2001). Other molecular markers such as sequence characterized amplified regions (SCARs) are based on the amplification of a DNA region with species-specific primers from RAPD fragments (Zijlstra et al. 2000). SCAR primers are available for M. arenaria, M. javanica, M. incognita (Zijlstra et al. 2000), M. hapla, Meloidogyne chitwoodi, and Meloidogyne fallax (Zijlstra 2000). This last method is easy to use and does not require DNA sequencing, but the SCAR markers developed so far do not amplify populations of M. hapla recovered from natural ecosystems (Ali et al. 2016).

    Finally, two nuclear markers, internal transcribed spacer (ITS) and D2D3 (gene portion 28S), provide sufficient genetic variation to distinguish nematode genera and species (Zijlstra et al. 1995) and allow characterization of Meloidogyne species (Ali et al. 2015; Castillo et al. 2003; De Ley et al. 1999) but require PCR amplification and Sanger sequencing.

    Efforts in nematode DNA barcoding highlighted the usefulness of the mitochondrial gene (Derycke et al. 2010; Kiewnick et al. 2015). This gene has advantageous properties because it exhibits multiple copies per cell as well as higher polymorphism levels than nuclear markers in general. It also demonstrates interspecific variability and strong maternal heritability that is especially useful for parthenogenetic species (Blouin 2002). Those properties enhance the detection of plant-parasitic nematodes and allow for the characterization of species among Meloidogyne (Powers 2004).

    Although Sanger DNA sequencing is a powerful way to characterize diversity for individuals, the detection of specific M. hapla haplotypes can also be efficiently conducted by quantitative PCR (qPCR). This methodology is based on fluorescent labels and allows for the specific detection of minute amounts of DNA matrices during the amplification. Many studies have shown how qPCR may be a fast, cheap, and sensitive technology for proper detection of plant-parasitic nematodes as long as qPCR primer pairs specific to each species of interest are available (Kawanobe et al. 2015). This approach was adapted to detect and identify Meloidogyne species (Berry et al. 2008; Braun-Kiewnick et al. 2016). However, most of the tests developed for M. hapla are either based on the noncoding ITS region (Dong et al. 2013; Watanabe et al. 2013) using mostly expensive hydrolysis probes (Sapkota et al. 2016) or based on 16D10 nuclear effector genes (Gorny et al. 2019), lacking experimental details and method performance tests, precluding its reproducible use by the scientific community (Bustin et al. 2009). In this context, and considering the expansion of M. hapla in vegetable-producing areas, this study aimed to provide a standardized and reliable tool to distinguish M. hapla from other nematode species. The method developed here is based on qPCR assays using the SYBR Green fluorophore and newly developed cytochrome oxidase I (COI)-specific primers. It was extensively tested on M. arenaria, M. incognita, M. javanica, and less closely related nematodes used as outgroups, and was assessed for standard performance criteria for sensitivity, specificity, repeatability, reproducibility, and robustness, complying with the minimum information for publication of quantitative real-time PCR experiments (MIQE) guidelines.

    Materials and Methods

    Specimen collection and DNA extraction.

    The development of a qPCR identification method for M. hapla required assessment of multiple RKN samples to assess the genericity of the diagnostic for M. hapla populations and also to insure the specificity of the detection with regard to other Meloidogyne species. The sampling design intended to address a diversity of geographic origin and of taxonomic divergence (Table 1).

    Table 1. Description of the samples used to validate the performance criteria of the quantitative PCR assay for Meloidogyne hapla identification

    A total of 15 nematode samples were tested, including 2 samples that were taxonomically distant species (i.e., Pratylenchus spp. and Paratylenchus spp.) as outgroups, 3 Meloidogyne samples belonging to a cryptic species complex (i.e., M. javanica, M. arenaria, and M. incognita), and 10 samples from M. hapla populations with distinct geographic origins.

    Each sample was inoculated separately onto a sensitive tomato plant (Solanum lycopersicum var. Roma VF) and bred in a growth chamber under controlled atmosphere (25 ± 2°C).

    Meloidogyne females and egg masses were extracted (manual dissection) from the infested roots under a stereomicroscope, saved individually in 5 µl of Trudgill buffer supplemented with 20% sucrose as described in Supplementary Material, then crushed, and stored at –20°C. After hatching of the corresponding egg masses (Madulu and Trudgill 1994), the hatched second-stage juveniles (J2) were conditioned (20 individuals per tube) in distilled water and stored at –20°C. Samples from Pratylenchus and Paratylenchus spp. were directly extracted after crushing and filtration of the infested roots (Coyne et al. 2010) and stored as for Meloidogyne species. For each sample, DNA extractions were performed from individuals using the DNeasy blood and tissue extraction kit (Qiagen, Courtaboeuf, France) according to the manufacturer’s instructions with slight modifications (incubation at 56°C for 4 h and final elution in 50 µl of distilled water heated to 70°C) to produce a reference DNA bank.

    Nematode identification.

    We identified the different Meloidogyne species using biochemical markers, based on electrophoretic separation of proteins, allowing the revelation of isoenzyme (i.e., esterase, the most common system used in the genus) (Bergé 1975). After grinding and centrifugation of conditioned females and electrophoresis on vertical polyacrylamide gel electrophoresis, a revelation of enzyme products by staining with 1-naphtylacetate (Brewer and Singh 1970) was performed (Supplementary Material). Migration reports (i.e., Rm) bands of esterase profiles were calculated for each female and compared with those of control populations of M. javanica and M. hapla live stock kept in the laboratory. Finally, the results were compared with those described in the literature (Esbenshade and Triantaphyllou 1985). Then, we confirmed this diagnostic result with a molecular characterization using the DNA extracted from juveniles. The complete protocol is provided in Supplementary Material. To summarize, we have used two different molecular markers: the SCARs to determine M. javanica, M. arenaria, and M. incognita species (Zijlstra et al. 2000), and, depending on the diagnosis obtained, we performed a second amplification of a portion of the 28S rDNA (D2D3) allowing the identification of M. hapla (Castillo et al. 2003). Pratylenchus spp. and Paratylenchus spp. populations were characterized after amplification and sequencing of the mDNA COI gene using universal primers COIF and COIR (Blouin 2002; Lazarova et al. 2006).

    Development of species-specific primers.

    The locus selected for identifying M. hapla is a portion of gene coding for the first subunit of cytochrome oxidase 1 (COI), an enzyme involved in the mitochondrial respiratory chain. Its rate of multicopy number per cell and high degree of polymorphism, interspecific variability, and maternal heritability (interesting for Meloidogyne with parthenogenetic reproduction) make this gene an excellent candidate as a molecular marker suitable for M. hapla diagnosis (Harris et al. 1990).

    A collection of COI sequences was created by combining all sequences available from the GenBank database corresponding to the four major species of RKNs (Blaxter et al. 1998) and COI sequences obtained from the RKN samples available in the laboratory with COIF and COIR primers (Supplementary Material). ecoPrimers-OBItools software (Riaz et al. 2011) was then used to design primers specific to M. hapla. The parameters selected for primer design included amplicon size (between 280 and 320 bp), primer size (minimum 20, maximum 25, optimum 22 nucleotides), melting temperature (Tm) (minimum 55°C, maximum 62°C, optimum 60°C), percentage of G and C nucleotides (GC%) (between 40 and 60%), quorum strict correspondence (98%), and quorum sensitivity (95%). After manual optimization of the different parameters and the 3′ specificity for forward and reverse primers, we retained a pair of primers specific to M. hapla (COIspecF/COIpsecR) to develop the qPCR method for M. hapla diagnosis as described in Supplementary Material. We confirmed on agarose gel that the qPCR product is of the size expected (339 pb).

    qPCR optimization and performance assessment.

    Newly developed methods used as standard assays in diagnostic analyses require reliability, cost effectiveness, and ease of application. This is why we chose qPCR with SYBR Green for the development of a robust approach for detecting M. hapla. This technique is based on the same principle as conventional PCR but is distinguished by the real-time detection of the amplicons during the amplification with a fluorescent dye. To monitor the incorporation of the dye during the PCR, we used a thermal cycler (Roche LC480) and specific intercalating dye of double-stranded DNA (LC480 SYBRGreen I Master, Roche Applied Science, Meylan, France) with high solution stability and very low residual fluorescence. Using this approach, the fluorescent signal is directly proportional to the amount of DNA amplified and indicates both the presence and quantity (when a quantification standard scale is used) of DNA marker from the organism targeted.

    The qPCR cycling conditions for COIspecF and COIspecR primers (specific to M. hapla) and COIF and COIR (universal primers) were 95°C initial denaturation for 10 min, 60°C annealing for 20 s, and 72°C extension for 30 s (45 cycles), followed by a final extension at 72°C for 10 min. All qPCR reactions were conducted in 20-µl final volume containing 3 µl of DNA extract, 1× PCR LC480 SYBRGreen I Master (Roche Applied Science), each primer at 0.25 µM, and 6 µl of distilled water. Amplifications were performed in a Roche LightCycler 480 Gradient Thermocycler. Positive controls (DNA extract obtained from a monospecific breeding and confirmed M. hapla population) as well as negative controls (distilled water) were included in all qPCR experiments, and all samples were tested as triplicates.

    To assess the performance of the method, we followed the recommendations given in the MIQE guidelines (Bustin et al. 2009) as well as in the methodological guide for characterizing the performance criteria of the French Agency for Food, Environmental and Occupational Health & Safety (Laurentie et al. 2015) to build the experimental design (Fig. 1).

    Fig. 1.

    Fig. 1. Experimental design used to validate diagnostic performance criteria. The dotted frames delimit the experiments conducted for each of the five performance criteria.

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    “Analytical sensitivity” refers to the minimum number of target DNA molecules in a sample that can be measured accurately with an assay. To estimate the sensitivity, we established the limit of detection (LOD), which is the smallest concentration that can be detected with reasonable certainty (95% probability is commonly used). A range of sequential dilutions (dilutions in eight steps, sequential factor 1/2) from a DNA extract of 20 J2s of M. hapla was established, and each dilution level was amplified with the specific primers COIspecF and COIspecR. Species diversity among Meloidogyne involved specimens’ size differences for the juveniles, which directly affected the initial concentration of extracted DNA. We could not measure it and rather expressed the relative concentration of each dilution level, and thus the LOD, as the number of theoretical number of individuals corresponding to each dilution factor (20, 10, 5 … juveniles). After defining the LOD, for all subsequent tests, the DNA extracts used were normalized to an optimal concentration that could be used to determine the other performance criteria (1/2 dilution level).

    “Analytical specificity” refers to detecting the appropriate target sequence rather than other species also present in a sample. To measure specificity, we ran two separate amplifications, one with M. hapla-specific primers (COIspecF and COIspecR) and the other with the universal COI primers on standardized DNA extraction from each nematode sample.

    Finally, we evaluated the analytical performance of the qPCR diagnostic method. First, we tested for repeatability, which refers to the precision of the assay for a given sample repeatedly analyzed under the same measuring conditions (same operator, same equipment, same laboratory). Second, we evaluated the reproducibility (i.e., the precision of the test under variable measuring conditions: same operator, different equipment, different laboratories).

    To evaluate these criteria, we repeated in three separate reactions (two within the same laboratory 2 days apart and a third assay in a different laboratory) the amplification with the specific primer pairs of a serial dilution of DNA extractions (20 J2s) from one M. hapla sample as well as from the 15 other nematode samples in three technical replicates.

    Last, we measured robustness, which refers to the reliability of the method in routine conditions. To measure its ability to withstand small changes in experimental conditions, three distinct amplifications were carried out with three different annealing temperatures (59, 60, and 61°C) on an M. hapla population and the same serial dilutions as previously detailed with M. hapla-specific primers.

    All three criteria were expressed as average crossing point (Cp; the minimum number of amplification cycles for which amplified DNA is detectable; i.e., for which there is both exponential amplification and a fluorescence level significantly higher than the background noise), and associated standard deviation (SD; dispersion indicator to measure the variability of the measured values).

    Data processing and statistical analysis.

    All criteria were evaluated after controlling the calibration curves and validation of two performance parameters associated with qPCR: amplification efficiency (E) in percent and linearity (expressed as the correlation coefficient R2). Efficiency rates were obtained from the slopes of the standard curves as follows:

    E=([10(−1/slope)]−1)×100.

    Linearity was measured as the coefficient of determination of the standard calibration curves obtained by linear regression analysis.

    Results

    Biochemical and molecular identification of nematode collection.

    An electrophoretic analysis of esterase isoenzyme patterns was conducted on conditioned Meloidogyne females to provide references for species identification.

    The “J3” phenotype of M. javanica (three bands’ Rm, 46, 54.5, and 58.9%) was used as a reference (control population) to calculate Rm of each population and confirm their identification (Esbenshade and Triantaphyllou 1985): M. javanica ‘Egypt’ (bands 1, 2, and 3 with Rm 46, 54.6, and 58.9%), M. arenaria ‘France’ (bands 4 and 5 with Rm 53.75 and 56.25%), M. incognita ‘France’ (band 1 with Rm 46%), and M. hapla (band 6 with Rm 49.66% for all 10 tested populations; Fig. 2A). To confirm this identification, we performed a complementary molecular characterization on the DNA extracted from juveniles by amplification of specific SCARs for M. javanica (670 bp), M. arenaria (420 bp), and M. incognita (1,200 bp; Fig. 2B) and by amplification, sequencing, and taxonomic assignation (through BLASTn) of a fragment of 28S D2D3 for M. hapla (Table 2). Pratylenchus spp. and Paratylenchus spp. populations were identified by amplification, sequencing, and taxonomic assignation with a fragment of mDNA COI gene (Table 2).

    Fig. 2.

    Fig. 2. Esterase phenotypes of Meloidogyne populations and sequence characterized amplified region (SCAR) phenotypes of Meloidogyne populations. A, Esterase phenotypes of Meloidogyne populations (R, Meloidogyne javanica reference as a standard for each migration; F1, M. javanica ‘Egypt’; F2, Meloidogyne arenaria ‘France’; F3, Meloidogyne incognita ‘France’; F4, populations of Meloidogyne hapla). The upper part of the figure schematizes the obtained migration profiles for each species. The lower part of the figure combines four migration images from the for species of interest. B, SCAR phenotypes of Meloidogyne populations (M, 1-Kb size marker DNA; F1, M. javanica ‘Egypt’; F2, M. arenaria ‘France’; F3, M. incognita ‘France’). The figure combines two migration images from three species of interest.

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    Table 2. Biochemical and molecular diagnostics of plant-parasitic nematodes populations in the laboratory collection population

    Molecular diagnosis of M. hapla by qPCR.

    Analytical and diagnostic sensitivities.

    The method developed should enable the identification of M. hapla specimens and requires performing measures of detection threshold for the target species with certainty. The standard range of dilutions made from a DNA extract of M. hapla allowed such calibration. A regression was obtained for the first five dilution levels with average Cp values of 27.56, 28.5, 29.48, 30.5, and 31.58; an efficiency E of 99.31%; and a correlation coefficient R2 of 0.9992. This advocated setting the LOD at the dilution 1/16, corresponding to a relative theoretical unit of 1.25 individuals, 170 times higher than fluorescence background signal recorded for the negative control. This qPCR assay has the ability to provide a positive result when the targeted species is present in less than this threshold, but, in these circumstances, the absence of the species cannot be guaranteed.

    Analytical specificity.

    Two qPCR amplifications were performed on the 15 nematode samples representing all species tested and on a dilution range with the four highest concentration levels, one with universal primers to verify the presence of DNA in each sample and a second with primers specific to M. hapla to test for their specificity and their ability to detect only the target species. For both essays, standard curves obtained were optimized both for their efficiencies (102.65 and 100.69%, respectively) and correlation coefficient (0.9927 and 0.9997, respectively).

    In the universal amplification, all samples tested were positive (average Cp between 33.27 and 37.67, less than the average Cp of the negative control, 39.00), which allowed validation of the quality of the DNA extraction and therefore the presence of DNA in each sample. For the specific amplification, all 10 M. hapla populations tested positive (average Cp between 30.72 and 34.71; Fig. 3), and no other species displayed any positive amplification. The qPCR assay is 100% specific to those species tested within this study, as all tested target populations were detected and no amplification was observed when other RKNs were tested, confirming the ability of the assay to specifically detect the target species M. hapla.

    Fig. 3.

    Fig. 3. Specificity of Meloidogyne hapla species-specific primers tested on all root-knot nematode (RKN) populations used in this study. Representation of standard curve linearity validated on four levels of sequential dilutions in triplicate (amplification efficiency E, 100.69%; correlation coefficient R2, 0.9997) and representation of average crossing point (Cp) values of all RKN populations amplified with specific primers (●, average Cp values of M. hapla populations; □, average Cp values of other RKN populations).

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    Repeatability and reproducibility.

    As for previous performance tests, serial dilutions of DNA isolated from one M. hapla population (20 juveniles) were prepared to evaluate the repeatability and reproducibility of the qPCR assays. Each dilution as well as the 15 nematode populations were amplified for three replicates and repeated in three separate reactions (two in the same laboratory and another one in a different laboratory) with specific primers.

    Intraassay variability was first tested on two experiments of serial dilutions on normalized DNA with specific primers COIspecF/COIspecR to determine the repeatability. The intraassay SD of the different assays for all serial dilutions ranged from 0.04 to 1.10 and 0.13 to 0.93, respectively. The efficiency E and the correlation coefficient R2 were determined from the standard curves and evaluated respectively at 94.82 and 110.4% with correlation coefficients R2 of 0.9564 and 0.9955. Both assays had a wide dynamic range of reliable amplification linearity of at least four orders of magnitude (from Cp 32.16 to 35.1 for assay 1; from Cp 32.39 to 35.21 for assay 2; Fig. 4A), and all average Cp values obtained for M. hapla populations were <39 when ≥39 for all other RKN populations when they appeared to be amplified.

    Fig. 4.

    Fig. 4. A, Repeatability or intraassay precisions of specific primer pair-based quantitative PCR (qPCR) assays. B, Reproducibility or interassay interlaboratory performance of specific primer-based qPCR assays.

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    The interassay variation was tested with two different laboratories to determine the reproducibility of the two assays. The interassay SDs obtained were from 0.15 to 0.53, and the two amplifications showed reaction efficiencies of 110.4 and 101.4%, with R2 of 0.9955 and 0.9536. Both assays had a wide dynamic range of reliable amplification linearity of at least three orders of magnitude (from Cp 32.39 to 35.1 for assay 1; from Cp 33.11 to 35.08 for assay 2; Fig. 4B), and all average Cp values obtained for M. hapla populations were <39 when ≥39 for all other RKN populations.

    The qPCR method is therefore repeatable and reproducible, as it allowed the specific detection of the target species multiple times within the same laboratory and in a different laboratory.

    Robustness.

    To complete the evaluation of the qPCR performance, the robustness was tested by three distinct amplifications of the same serial dilution each time with three replicates for one M. hapla population, with specific primers and varying by three annealing temperatures that mimic thermocycler small discrepancies (59, 60, and 61°C). All three assays had a wide dynamic range of reliable amplification linearity (from Cp 32.29 to 35.34 for amplification at 59°C, from Cp 32.16 to 35.1 for amplification at 60°C, and from Cp 33.64 to 36.59 for amplification at 61°C). Amplification efficiencies were 94, 95, and 93% with associated R2 values of 0.9927, 0.9564, and 0.9555, and all average Cp values measured for M. hapla populations were <39. Because the target species is detected despite a ±1°C annealing temperature variation, the reliability of the method can be validated under routine usage conditions (Fig. 5).

    Fig. 5.

    Fig. 5. Sequential dilution curves of three intralaboratory amplifications in triplicate at three different annealing temperatures.

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    Discussion

    Observable crop damage caused by nematodes is usually nonspecific and easily confused with symptoms of abiotic or biotic origin. The presence of galls on plant roots remains a good indicator of the presence of RKNs Meloidogyne spp., but it does not allow characterization of the species involved. Such symptoms are observable in the field only quite late in the population dynamics, and do not allow for mitigation of the agronomic damage. The damage threshold for RKN may be a single nematode for susceptible crops, or when climatic and soil conditions are favorable (Peiris et al. 2018). Being able to detect RKN early is central to the effectiveness of crop protection methods (Coyne et al. 2018).

    In this context, molecular diagnostic tools are best suited to resolve this issue. Quantitative real-time PCR in particular, used in research and diagnosis, already proved its effectiveness for the characterization of plant-parasitic nematodes (Braun-Kiewnick and Kiewnick 2018) like M. hapla (Gorny et al. 2019; Sapkota et al. 2016). It meets this imperative for faster and more reliable characterization of harmful nematodes, at a lower cost and high reproducibility. The method developed here differs from that described by Braun-Kiewnick and Kiewnick in 2016 because it uses a specific double-stranded DNA intercalant (SYBR Green), avoiding expensive TaqMan hydrolysis probes that require design expertise and are subject to a significant risk of false-negative findings when polymorphism occurs. Using qPCR enables to monitor the PCR reaction during its exponential phase in which the increase in the quantity of amplicons is highly correlated to the initial quantity of the target matrix. This allowed the establishment of an LOD equivalent to a relative theoretical unit of 1.25 J2 M. hapla, close to the threshold of damage as defined by Peiris et al. (2018).

    A set of target genomic regions based on groups of 18S/28S or ITS ribosomal genes is available and is widely used for phylogenetic analyses between closely related species (Dong et al. 2013; Sapkota et al. 2016) or genes coding for effector proteins (Gorny et al. 2019; Yu et al. 2010). Unfortunately, the previously introduced DNA markers lack resolution to distinguish between closely related species (Janssen et al. 2016). Also, they lack specificity with regard to cross-reactions with nontarget species (Sapkota et al. 2016). This led to the development of a new set of specific primers in an appropriate barcode region located on the gene coding for mitochondrial COI that is fully suitable for this purpose (Gissi et al. 2008; Powers et al. 2018). When developing the method, we paid particular attention to experimental details, and tests of the stability of the performance of the methods developed the basis for the development of a reliable and reproducible diagnostic technique (Bustin et al. 2009, 2010) for routine use. We therefore chose to measure different diagnostic and analytical performance criteria as established by the official guide for characterizing performance criteria (Laurentie et al. 2015). Like Gorny et al. (2019), we controlled sensitivity and established an LOD equivalent to 1.25 juveniles. Like Kiewnick et al. (2015), we measured the specificity of the primers developed and obtained 100% detection of M. hapla populations, without cross-reaction with the other Meloidogyne species tested. We completed these measurements by evaluating and confirming three additional criteria: repeatability, reproducibility, and robustness. This whole set of specifications concurs with the standardized methods of the MIQE guidelines (Bustin et al. 2009) and the minimum standards defined by Bustin et al. (2010). To this end, we set up our experimental design to control sources of variation and validate quality results at every single step of the process. This method has a strong potential as an effective decision support tool, improving fast and effective diagnosis of M. hapla.

    Acknowledgments

    The authors thank Philippe Castagnone (director of Institut Sophia Agrobiotech, National Institute for Agricultural Research (INRAE), Sophia Antipolis, France) and the entire Plant–Nematode Interactions team for providing us with RKN (Meloidogyne hapla) species and Laurent Folcher (head of the National Reference Laboratory covering phytoparasitic nematodes on all matrices, French Agency for Food, Environmental and Occupational Health and Safety/Plant Health Laboratory (ANSES), Le Rheu, France) and the entire nematology unit for training and support in the development of this standardized diagnostic method.

    The author(s) declare no conflict of interest.

    Literature Cited

    The author(s) declare no conflict of interest.