Detection of Airborne Sporangia of Pseudoperonospora cubensis and P. humuli in Michigan Using Burkard Spore Traps Coupled to Quantitative PCR

Pseudoperonospora cubensis (Berk. & M. A. Curtis) Rostovzev, the causal agent of cucurbit downy mildew (CDM), infects approximately 20 cucurbit genera including the economically important crops of cucumber (Cucumis sativus), cantaloupe (Cucumis melo var. cantalupensis), squash (Cucurbita), watermelon (Citrullus lanatus), and pumpkin (Cucurbita pepo; Cohen et al. 2015; Lebeda and Cohen 2011). CDM symptoms include angular, chlorotic lesions that coalesce and become necrotic, resulting in leaf blight and death; pathogen sporulation occurs on the abaxial side of the leaf (Salcedo et al. 2020). In cucumber, foliar blighting resulting from CDM can result in yield reduction (Hausbeck et al. 2019; Perla et al. 2019; Reuveni et al. 1980).

In the United States, Michigan is the largest producer of pickling cucumber and the second-largest producer of cucumber for the fresh market, with approximately 4.1 million hundredweight (cwt) of cucumbers sold in 2018 (U.S. Department of Agriculture 2020). For nearly 40 years, resistant cucumber cultivars had been used successfully to mitigate CDM (Brzozowski et al. 2016). In 2004, a highly virulent strain of P. cubensis emerged in the United States overcoming this host resistance (Thomas et al. 2017), and since then, fungicides have been relied on for control (Blum et al. 2011; Holmes et al. 2015). However, fungicide-resistant P. cubensis isolates have presented crop protection challenges. Single-site fungicides including mefenoxam and azoxystrobin were ineffective when the pathogen emerged in 2004 (Ernest et al. 2005; Gevens and Hausbeck 2005; Thornton et al. 2006). Since that time, P. cubensis resistance to dimethomorph (Zhu et al. 2007) and mandipropamid (Blum et al. 2011; Hausbeck and Cortright 2010) has been reported in the United States (Holmes et al. 2015; Keinath 2015). Similarly, reduced efficacy of fluopicolide against CDM has been observed in field trials in Michigan (Hausbeck and Linderman 2014), Georgia (Langston and Sanders 2013), and North Carolina (Adams and Quesada-Ocampo 2014). While propamocarb was effective against CDM for several years, since 2013 its efficacy appeared to be compromised in field trials in North Carolina (Keinath 2015; Thomas et al. 2018), Pennsylvania (Gugino and Grove 2016), and Michigan (Hausbeck et al. 2017; Hausbeck and Linderman 2014).

P. cubensis is an obligate pathogen and its survival depends on the availability of susceptible hosts (Cohen et al. 2015). The pathogen does not survive in regions that experience frost; instead, its sporangia are dispersed to northern latitudes from overwintering sites (Ojiambo and Holmes 2011; Quesada-Ocampo et al. 2012). Airborne sporangia concentrations influence CDM onset (Granke et al. 2014) and under conducive weather conditions, P. cubensis sporangia can spread rapidly within and between fields (Ojiambo et al. 2015). Airborne concentrations of P. cubensis sporangia in Michigan’s cucumber fields have been monitored using Burkard spore traps (Burkard Manufacturing) with light microscopy to identify and enumerate the pathogen sporangia based on morphology (Granke et al. 2014; Granke and Hausbeck 2011). Pseudoperonospora humuli, the causal agent of hop downy mildew (HDM), is nearly identical to P. cubensis morphologically (Runge and Thines 2011), but rarely infects cucurbits in the United States (Mitchell et al. 2011). Approximately 400 ha of hops are planted in Michigan (Michigan Department of Agriculture & Rural Development 2018) and P. humuli is prevalent (Lizotte and Miles 2020). Thus, relying on morphological identification, alone, to monitor airborne sporangial concentrations of P. cubensis could result in inaccurate estimations of the pathogen’s presence and concentration.

PCR-based methods have been used successfully to detect and quantify airborne plant pathogens such as Peronospora effusa (Klosterman et al. 2014), Peronospora schachtii (Klosterman et al. 2014), Claviceps purpurea (Dung et al. 2018), and P. humuli, infecting spinach (Pinacia oleracea), beet (Beta vulgaris), grass-seed (Lolium perenne), and hop (Humulus lupulus; Gent et al. 2009), respectively. A quantitative PCR (qPCR) assay was developed that differentiates between P. cubensis and P. humuli sporangia (Summers et al. 2015). This assay, with or without microscopic visualization of spore trap tapes, could accelerate the speed and accuracy of P. cubensis detection and inform the initiation of fungicide sprays. The objective of our study was to improve the detection of airborne concentrations of P. cubensis sporangia by adapting a qPCR-based assay (Summers et al. 2015) that distinguishes between P. cubensis and P. humuli using Burkard spore trap samples collected in the field.

Materials and Methods

In vitro evaluations to assess the sensitivity of the qPCR-based assay were performed using isolates of P. cubensis (CDM23) and P. humuli (HDM19) obtained in 2017 using methods similar to those described by Thomas et al. (2017). Briefly, diseased tissue was placed in a moist chamber overnight to induce sporulation. Sporangia from a single cucumber leaf lesion or an infected hop basal shoot (spike) were suspended in 1 ml of distilled water and the resulting inoculum (1,000 to 10,000 sporangia/ml) applied to the abaxial side of detached leaves of ‘Vlaspik’ cucumber or ‘Centennial’ hop, respectively, contained in Petri dishes (100 × 15 mm). Inoculated leaves were then incubated in a growth chamber at 18°C under a 12/12‐h light/dark cycle. Seven to 10 days post-inoculation, sporangia were gently rinsed from infected leaves using a Preval spray power unit filled with distilled water. A new set of leaves was inoculated with the resulting sporangia.

Collection of sporangia and extraction of genomic DNA.

P. cubensis (CDM23) and P. humuli sporangia (HDM19) were gently rinsed from host tissue into centrifuge tubes (50 ml) using a Preval spray power unit filled with distilled water. The sporangial suspension was concentrated by centrifugation (5424R centrifuge, Eppendorf) at 14,000 rpm for 5 min and homogenized in MP Biomedicals Lysing Matrix H impact-resistant 2-ml tubes using a Qiagen TissueLyser II (Qiagen, Valencia, CA) for 4 min at 30 Hz. DNA was extracted using a Macherey-Nagel NucleoSpin Plant II isolation kit (Macherey-Nagel, Bethlehem, PA) following manufacturer’s instructions, and the DNA concentration was determined using a Life Technologies Qubit double-stranded DNA High Sensitivity Assay Kit (Life Technologies, Carlsbad, CA).

Competitive qPCR internal control.

A competitive positive internal control (IC) was designed in this study and incorporated into every qPCR reaction to monitor for the presence of PCR inhibitors in each sample. The IC consisted of a single-stranded linear synthetic DNA that utilizes the same primers of the target mitochondrial cox2 gene, and an additional fluorogenic probe (ICprobeJ2: /5CYS/A+GCATTATT+GTTTAT+CATATATACA/3IABkFQ/) for amplification and detection. The sequence of the IC (Table 1) showed no significant nucleotide identity to any known naturally occurring PCR-amplifiable nucleotide sequences reported in the NCBI database.

Table 1. Primers and locked nucleic acid (LNA) probes for the quantitative-PCR assay differentiating Pseudoperonospora cubensis and P. humuli using the 105 single nucleotide polymorphisms in the mitochondrial Cox2 gene^v

	Code name	Concentrationw	Sequence 5′->3′
Primer	RT33Fx	600 nM	AACTCCCGTTATGGAAGGTATT
Primer	RT182Rx	600 nM	CCATGTACAACAGTAGCTGGA
Probe	CUBprobeSNP105x	250 nM	HEX/A+C+AAA+C+G+AATA+CT/BHQy
Probe	HUMprobeSNP105x	500 nM	FAM/AA+C+AAA+C+A+AATA+CTG/BHQy
Probe	ICprobeJ2	250 nM	CYS/A+GCATTATT+GTTTAT+CATATATACA/BHQy
Internal control	IC	7.5 × 10−10 nM (0.75 aMz)	AACTCCCGTTATGGAAGGTATTATCATTAATCAT
			GATTTGTATATATGATAAACAATAATGCTATAAC
			ATAGAGTCTCTTTCATGAATAATCCAGCTACTGT
			TGTACATGG

^v CUB, P. cubensis; HUM, P. humuli; IC, internal control; HEX, hexachlorofluorescein; FAM, fluorescein amidite; CYS, cysteine. nM, nanomolar.

^w Concentrations used in a 20-μl quantitative-PCR reaction.

^x Primers and LNA probes were adapted from Summers et al. (2015).

^y LNAs in the probes are followed by a plus (+) sign.

^z Attomole (aM) = 10⁻¹⁸ moles per liter.

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qPCR protocol with purified DNA.

All qPCR experiments were conducted using a modified version of the protocol described by Summers et al. (2015) in accordance with the Minimum Information for Publication of Quantitative Real-Time Experiments guidelines (Bustin et al. 2009). The Summers et al. (2015) assay was modified by changing the commercial master mix, Bio-Rad IQ Supermix (Bio-rad, Hercules, CA), to the IDT Prime-Time Gene Expression Master Mix (IDT, Skokie, IL). This new master mix reduced the variation between technical replicates and increased the amplification efficiency of qPCR reactions. Additionally, we also added an internal-positive control to identify any alterations in amplification efficiency in field air samples. qPCR reactions with a final volume of 20 µl were manually assembled in Bio-Rad MLL9651 96-well white plates containing 10 µl of the Prime-Time Gene Expression Master Mix, 2 µl of template DNA, 600 nM of each primer (RT33F and RT182R), 500 nM of the locked nucleic acid (LNA) probe HUMprobeSNP105, 250 nM of the LNA probe CUBprobeSNP105, 250 nM of the LNA probe ICprobeJ2, and 7.5 × 10⁻¹⁰ nM (0.75 aM) of our IC (Table 1). The IC was set to this concentration to obtain a C_q value of 29 without affecting the sensitivity or specificity of the other probes. Negative control reactions lacking the DNA template were included in every plate run. The qPCR protocol was run on a Bio-Rad CBio-FX 96 Touch qPCR system and included an initial denaturation step at 95°C for 3 min followed by 38 cycles of 95°C for 10 s and 65°C for 45 s. Two technical replicates of each sample were run and the average C_q value and standard deviation were calculated using Bio-Rad CFX Manager software (version 3.1; Bio-Rad #1845000).

Sensitivity and specificity of qPCR with LNA probes.

The Burkard spore traps use a vacuum pump to draw air (approximately 10 liters/min) into a collection chamber containing a reel, covered with Melinex tape (DuPont; Burkard Manufacturing); the reel was mounted on a clockwork mechanism (Hirst 1952). The Melinex tape (Burkard Manufacturing Co. Ltd., U.K.) was coated with an adhesive of petroleum jelly and paraffin (9:1 weight/weight) dissolved in sufficient toluene to provide adequate coverage of tape at the desired thickness (Granke et al. 2014).

The sensitivity and specificity of the qPCR assay was evaluated using three different experiments; the first generated a standard curve using pure DNA of P. humuli and P. cubensis, the second used samples that contained a mixture of P. humuli and P. cubensis DNA, and the third used samples of P. cubensis sporangia that also contained the Melinex tape and the adhesive used on the Burkard spore trap tapes. The first procedure included 10-fold dilutions of genomic DNA from two independent DNA extractions from each pathogen isolate (‘CDM23’ and ‘HDM19’) that were used to generate standard curves ranging from 10 to 100,000 fg. Three technical replicates of each sample dilution were tested using qPCR, and the average C_q values with the standard deviation were calculated using the Bio-Rad CFX Manager software. Mean C_q values were plotted against the log₁₀ of template DNA concentrations and used to generate standard curves. The second procedure included the evaluation of mixed-DNA samples to assess the specificity of the qPCR assay. An in vitro assessment was used to determine whether the assay could detect P. cubensis and P. humuli in mixed samples containing DNA from both pathogens. Ten-fold dilutions of genomic DNA from each species were made from 100 to 100,000 fg and used as templates, both separately and mixed in varying concentrations (Table 2), for the qPCR assay described above. Three technical replicates of each concentration and mixture were run and the average C_q values and standard deviations were calculated using Bio-Rad CFX Manager software. The third procedure included five independent extractions from solutions containing 10, 100, or 1,000 sporangia of P. cubensis prepared using a hemocytometer counting cell chamber. Sporangia were homogenized in impact-resistant 2-ml tubes using a Qiagen TissueLyser (4 min at 30 Hz) and DNA was extracted using a Macherey-Nagel NucleoSpin Plant II isolation kit. Subsequently, 2 µl of the extraction products were evaluated using qPCR. Finally, 9- × 48-mm sections (representing a 24 h-sampling period) of Melinex tape with adhesive were spiked with 10, 20, 50, 100, or 300 P. cubensis sporangia. DNA was extracted and evaluated using qPCR as previously described. Three technical replicates of an average of four independent extractions of each sample dilution were tested and the average C_q values with standard deviation were calculated using the Bio-Rad CFX Manager software.

Table 2. Threshold cycle (C_q) values of quantitative PCR assays using locked nucleic acid probes and varying concentrations of genomic DNA^z

DNA			Probe
	Pseudoperonospora cubensis	Pseudoperonospora humuli	CUBprobeSNP105		HUMprobeSNP105
			Cq-HEX	SD	Cq-FAM	SD
Unmixed	10,000 fg	–	28.39 a	0.10	NA	NA
	1,000 fg	–	32.12 b	0.21	NA	NA
	100 fg	–	35.49 c	0.74	NA	NA
	–	10,000 fg	29.88 nc	0.34	28.29 d	0.08
	–	1,000 fg	32.96 nc	0.17	31.70 e	0.30
	–	100 fg	NA	NA	36.61 f	0.14
Mixed	10,000 fg	10,000 fg	28.67 a	0.23	28.52 d	0.60
	1,000 fg	1,000 fg	31.44 b	0.37	32.42 e	0.08
	100 fg	100 fg	35.32 c	0.98	35.11 f	0.59
	10,000 fg	1,000 fg	28.37 a	0.15	32.07 e	0.33
	10,000 fg	100 fg	28.39 a	0.26	NA	NA
	1,000 fg	10,000 fg	29.78	0.37	27.96 d	0.34
	100 fg	10,000 fg	30.58	0.24	24.99 d	5.91

^z The hexachlorofluorescein (HEX)-labeled probe (CUBprobeSNP105) was designed to detect only DNA from P. cubensis, and the fluorescein amidite (FAM)-labeled probe (HUMprobeSNP105) was designed to detect only DNA from P. humuli. C_q values with the same letter are not significantly different (Student’s t-test; P = 0.05). fg, femtogram; –, not DNA added; NC, not used for comparison in the Student’s t-test; NA, not defined; SD, standard deviation.

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Collection of field samples for screening using qPCR.

Airborne sporangial concentrations were monitored during the cucumber growing season (May to September) in 2018 and 2019 using Burkard spore traps. Each year, a spore trap was placed 20 m from a commercial cucumber field located in Muskegon County in northwest Michigan and a cucumber research plot at the Michigan State University Plant Pathology Farm located in Ingham County in south-central Michigan. The Michigan State University cucumber research plot (0.25 ha) was direct seeded during the last week of July and was located 200 m from an abandoned hop research yard (0.25 ha) where systemically infected basal shoots (spikes) were observed beginning in late April 2019. An additional Burkard spore trap was placed in a commercial hop yard in Berrien County in 2019.

The reel in each Burkard trap was covered with a Melinex tape coated with an adhesive as described previously. The tape was removed weekly and cut longitudinally along the center line into two subsections of 9- × 336-mm each (Rogers et al. 2009). The first section was then cut into 48-mm lengths (equivalent to a monitoring period of 24 h), scored at hourly intervals (2 mm), and stained to facilitate counting according to the protocol described by Granke et al. (2014). The second section was also cut into 48-mm lengths, placed into impact-resistant HP Biomedicals Lysing Matrix H 2-ml tubes, and subjected to DNA extraction as previously described using a Macherey-Nagel NucleoSpin Plant II isolation kit (Macherey-Nagel, Bethlehem, PA). Subsequently, 2 µl of the extraction product was evaluated using qPCR. Fields were scouted weekly for signs and/or symptoms of P. cubensis. Leaf samples with lesions resembling CDM symptoms and signs of the pathogen were returned to the laboratory and examined using light microscopy to verify the presence of sporangia.

Results

Sensitivity and specificity of qPCR.

Using 10-fold dilutions of P. cubensis and P. humuli DNA, the qPCR assay exhibited a significant linear response with an efficiency of 93.6% (R² = 0.99) and 90.7% (R² = 1), respectively (Fig. 1A and C). Both species-specific LNA probes detected each pathogen within total DNA template amounts ranging between 100 and 100,000 fg per reaction (Fig. 1A and C). The average C_q values for samples containing 100 fg of P. humuli and P. cubensis DNA were <35.5. Most samples with concentrations <100 fg were either not detected or were detected without reasonable certainty (<95% of the times tested); thus, 100 fg of template DNA was considered as the lower limit of detection of the qPCR assay for both species. Although the LNA probes were specifically designed to detect either P. humuli or P. cubensis based on the recognition of a single nucleotide polymorphism (SNP) at the 105-base of the cox2 gene (Summers et al. 2015), the hexachlorofluorescein-labeled LNA probe CUBprobeSNP105 for P. cubensis detection showed non-specific amplification of P. humuli DNA. However, the amplification curves of P. humuli DNA with this probe did not show the same shape as those generated using the P. cubensis DNA (Fig. 1B). P. cubensis samples were classified as “positive” based on the shape of the amplification curve using the probe CUBprobeSNP105 and no amplification of the fluorescein amidite-labeled LNA probe HUMprobeSNP105. This probe (HUMprobeSNP105), designed to recognize P. humuli, was highly specific and no background signal was observed when P. cubensis DNA was analyzed (Fig. 1D).

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Additionally, when mixed samples containing DNA from both species were assessed using the qPCR assay, no significant changes were observed in the C_q values of the samples containing P. cubensis or P. humuli DNA in a 1:1 ratio (Table 2). However, when 10,000 fg of P. cubensis DNA was mixed with 100 fg of P. humuli DNA, no detection of P. humuli DNA was observed (Table 2). Detection of P. cubensis occurred in all mixtures, but when 10,000 fg of P. humuli DNA was mixed with 1,000 or 100 fg of P. cubensis DNA, the detection of P. cubensis occurred at significant lower C_q values than in reactions including only P. cubensis DNA (Table 2). Mixed samples containing DNA from both species generated amplification curves with both probes (CUBprobeSNP105 and HUMprobeSNP105). This clearly differentiated them from samples containing only P. cubensis DNA for which there was only amplification with the CUBprobeSNP105 probe (Supplementary Fig. S1A and C). Although the samples containing only P. humuli DNA also generated amplification curves with both probes (Supplementary Fig. S1B), the amplification curves generated with the CUBprobeSNP105 probe for mixed samples showed faster growth in the exponential phase of the curves. This was the case for mixed samples containing DNA from both species in a 1:1 ratio and samples containing P. cubensis and P. humuli DNA in 10:1 ratio (Supplementary Fig. S1C and D). Only the amplification curves of the samples containing P. cubensis and P. humuli DNA in the ratios 1:10 and 1:100 were not clearly differentiated from the amplification curves containing only P. humuli DNA (Supplementary Fig. S1B and E).

Both LNA probes detected DNA from extractions containing 10, 100, or 1,000 sporangia of P. cubensis and P. humuli. Upon regression analysis, a linear relationship between C_q values and DNA extracted from purified sporangia was observed for both species (P. cubensis, R² = 0.99, P_value = 0.03; and P. humuli, R² = 0.98, P_value = 0.084; Fig. 2A). The average C_q value for detecting 10 P. cubensis sporangia was <35.5 (Fig. 2A) and samples with <10 sporangia could not be detected with reasonable certainty (>95% of the times tested). C_q values ≤35.5 were classified as specific to P. humuli and P. cubensis and were used as a threshold to evaluate field samples.

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The sensitivity of the assay to detect P. cubensis was minimally affected by the adhesive applied to the Melinex tape (Fig. 2B). DNA was detected from extractions of tapes spiked with 20, 50,100, or 300 sporangia and a reduced number of samples (4/10) containing 10 sporangia had average C_q values <35.5. All the extractions showed C_q values that were significantly different from the background signal observed in the negative controls. The relationship between sporangial numbers and C_q values was significant (P = 0.003) and the assay exhibited a linear response with a R² value of 0.99 (Fig. 2B). The average C_q values of the different sporangial dilutions were within the 95% confidence interval. However, a high standard error of the mean was observed among biological replicates of extractions with the same number of sporangia (Fig. 2B), indicating that the extraction affects the precision of quantifying sporangia using the qPCR-based assay.

Assessment of field samples using light microscopy and qPCR.

A total of 560 samples collected from May to August in 2018 and 2019 were assessed using qPCR. P. cubensis or P. humuli DNA was detected using qPCR on field samples with <10 sporangia (Fig. 2C). Approximately 90% of all samples that tested positive for either pathogen using qPCR (204 out of 227) had one or more sporangia on the corresponding half of the tape analyzed using light microscopy. The average C_q value of the IC remained relatively constant and had an average of 28.8 ± 1.7 (standard deviation) among all the field samples evaluated. Regression analysis indicated that the number of sporangia on the second half of the tape of the samples (quantified using light microscopy) explained 54% (R² = 0.54) and 10% (R² = 0.10) of the variation in the C_q values of P. cubensis and P. humuli, respectively (Fig. 2C).

Using light microscopy, Pseudoperonospora spp. sporangia were first detected in 2018 during May (Muskegon County commercial field, 15 May) and June (Ingham County research field, June 13; Fig. 3). From May to July at the research field, fewer than five sporangia/day were observed via light microscopy with P. cubensis DNA, which was confirmed using qPCR on 13 June and 10, 23 July; P. humuli was confirmed on 19 June and 7 July (Fig. 3A). At the commercial field, <10 sporangia/day were detected using light microscopy from May to July, except for 21 June (Fig. 3B). Using qPCR, we confirmed the presence of P. humuli DNA on 29 May to 4 June and 21 to 26 June (Fig. 3B), while P. cubensis DNA was detected on 5 and 13 June (Fig. 3B). Airborne sporangial concentrations increased during the first week of August and reached the maximum during the third week of the month in both locations monitored in 2018 (Fig. 3A and B). CDM symptoms were observed at the research field on 15 August; a peak of 16 sporangia was observed via light microscopy 10 days prior (5 August) and P. cubensis DNA was confirmed by qPCR on 1 to 6 August and 10 August (Fig. 3A). After CDM symptoms were observed in the research field, daily sporangial counts via light microscopy exceeded 10 sporangia/day with P. cubensis DNA detected nearly every day using qPCR (Fig. 3A and B). CDM symptoms were observed at the commercial field on 7 August; a peak of 22 sporangia was observed via light microscopy 4 days prior (3 August) and P. cubensis DNA was confirmed by qPCR on 3 and 5 August, and 28, 30, and 31 July (Fig. 3B). The day after the detection of CDM symptoms, >80 sporangia/day were captured by the spore traps and P. cubensis DNA was detected every day using qPCR (Fig. 3B).

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In 2019, Pseudoperonospora spp. sporangia were first detected in May across all locations using light microscopy (Fig. 4). During May and June, concentrations of airborne sporangia exceeded 10 sporangia/day in the commercial hop yard (Fig. 4A) and the research field (Fig. 4B). Using qPCR, P. humuli DNA was detected several times in the commercial hop yard from May through August while P. cubensis DNA was only detected on 12, 14, and 18 August (Fig. 4A). Based on data from light microscopy, the research field that was in proximity to a non-treated hop yard had >10 sporangia/day during the last week of May, the first and fourth week of June, and the first week of July (Fig. 4B). At this location, P. humuli DNA was detected from May to July using qPCR (Fig. 4B). During August, <10 sporangia/day were observed at the research field. CDM symptoms were confirmed at this site on 21 August and P. cubensis DNA was verified by qPCR on 11, 19, and 20 August. After CDM symptoms, P. cubensis DNA was detected from 22 to 31 August (Fig. 4B). In the commercial cucumber field, <10 sporangia/day were observed using light microscopy from May through July except for 18, 21, and 23 June (Fig. 4C). P. humuli DNA was confirmed with qPCR on 22 and 23 June (Fig. 4C). At this location, the sporangial counts increased from the third to the last week of August. CDM symptoms were confirmed on 16 August, and P. cubensis DNA was detected using qPCR in air samples every day from 12 to 31 August (Fig. 4C).

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Discussion

The ability to detect and differentiate between P. cubensis and P. humuli in field air samples using qPCR represents an important advance for CDM monitoring and management. The qPCR detection of airborne sporangia could be used as a decision-making tool to initiate fungicide sprays (Dhar et al. 2019) or as a complementary variable to forecast the risk (Carisse et al. 2009) of CDM outbreaks in Michigan. Early and specific detection of P. cubensis sporangia could ensure timely crop protection and avoid unnecessary fungicide applications. P. cubensis does not overwinter in Michigan; for disease to occur, the pathogen must be introduced into the state’s growing regions annually (Naegele et al. 2016). Burkard spore traps coupled with light microscopy have been used since 2007 to alert Michigan growers to an influx of P. cubensis sporangia into their growing region. However, HDM is prevalent in the state (Lizotte and Miles 2020) where approximately 400 ha of hops have been planted (Hop Growers of America 2019). Using a qPCR assay, we were able to distinguish between the morphologically similar sporangia of P. cubensis and P. humuli collected from Burkard spore traps. During the 2 years of monitoring using Burkard spore traps coupled with the qPCR assay in cucumber fields, P. cubensis sporangia were detected approximately 5 to 10 days before CDM symptoms were observed in monitored cucumber fields.

We adapted the qPCR assay developed by Summers et al. (2015) to a high degree of sensitivity for use with the Burkard spore trap samples. Using DNA extracted from purified sporangial suspensions of P. cubensis and P. humuli, we were able to detect DNA concentrations ranging from 100 fg to 100,000 fg. This sensitivity was validated with the detection of the two downy mildew pathogens in field samples containing <10 sporangia (C_q < 35.5). We split the tape of Burkard spore traps to facilitate the comparison between light microscopy and qPCR, and observed that the number of sporangia deposited onto one-half of the tape was linearly correlated with the C_q values obtained after the assessment of the other half using qPCR. However, the change in the number of sporangia on field samples quantified using light microscopy explained only 54 and 10% of the variation in the C_q values of P. cubensis and P. humuli, respectively, suggesting a limited capacity of the qPCR assay for the absolute quantification of sporangia in field samples.

The low correlation between C_q values and sporangial numbers of field samples may be explained by the high variation in the yield of DNA extraction among samples (Summers et al. 2015), inaccurate visual quantification, the low specificity of one of our probes, and possibly, the multicopy nature of the target sequence (Dung et al. 2018; Klosterman et al. 2014; Kunjeti et al. 2016). Similarly, the yield variation of DNA extractions among samples may also explain the variation observed in the C_q values of samples with the same number of sporangia in vitro. This variation is introduced in all the samples collected in the field and may reduce the precision for the quantification of sporangia using the extraction protocol and qPCR assay described in this study. However, assessing the first half of the spore trap tape using qPCR could reduce the number of samples that require microscopic analysis for spore quantification, accelerating the turn-around time associated with monitoring airborne P. cubensis sporangia. The reduction of variation in the yield of DNA extractions and the utilization of a qPCR assay based on a single-copy marker may be more appropriate for quantification (Rahman et al. 2020), but may result in a system with reduced sensitivity when compared with the multicopy system that we used.

The non-specific amplification of P. humuli DNA affected the quantification capacity of the assay when both species were present in the same reaction. However, the inclusion of a second probe ensured that the detection of each species was possible even when P. cubensis and P. humuli were present in the same sample. The non-specific amplification of P. humuli did not occur under the conditions described by Summers et al. (2015), and was a consequence of the change in the commercial master mix used for the qPCR reactions (Supplementary Fig. S2). The master mix used in this study reduced the variation among technical replicates (data not shown) and increased the amplification efficiency of the qPCR reactions (exponential phase), but it affected the specificity of the assay. Different qPCR master mixes influence how oligonucleotides (primers and probes) bind to target regions (Morinha et al. 2020); thus, the suitability of new reagents must be carefully evaluated, as they may condition the results of the qPCR. The detection of both species using our qPCR assay was hindered only in samples with a significantly higher amount of P. humuli compared with P. cubensis (i.e., samples containing P. cubensis and P. humuli DNA in the ratios 1:10 and 1:100). In these cases, or in locations where a higher number of P. humuli sporangia relative to the number of P. cubensis sporangia is expected (e.g., hop yards), the use of the Bio-Rad IQ Supermix, (Bio-rad, Hercules, CA) as described by Summers et al. (2015) for qPCR reactions, should allow a more accurate evaluation of the samples.

During 2 years of monitoring in commercial cucumber fields, we did not detect any periods when both pathogens were detected simultaneously; however, overlapping periods may have occurred at the commercial hop yard during late August. The identification of genetic regions with a higher number of polymorphisms has allowed the design of more specific primers and probes for P. cubensis detection (Rahman et al. 2020). Using this new set of primers and probes in combination with the probe HUMprobeSNP105 (for detection of P. humuli) could ensure both specific detection and quantification of P. cubensis and P. humuli sporangia, using qPCR even during periods when both species are present.

Despite the limitations of the qPCR assay described in this study, we were able to detect low atmospheric concentrations of P. cubensis and P. humuli (<10 sporangia/day). Detection of P. cubensis before symptoms developed in the field was linked to a sporangial concentration <10 sporangia/day as estimated using light microscopy. In other crops including lettuce (Lactuca sativa) and onion (Allium cepa), measurements of aerial spore load (sporangia/day) have been used to guide fungicide application to control Bremia lactucae (Dhar et al. 2019) and Botrytis squamosa (Carisse et al. 2009), respectively. In these systems, fungicide applications began once spore loads reached a critical level between 300 and 500 spores/day (10 sporangia/m³). In the cucumber fields monitored in 2018, CDM symptoms were observed after airborne P. cubensis sporangial concentrations were >15 sporangia/day as determined via light microscopy. This suggests that the critical concentration for CDM could be close to this number, depending on the coincident environmental conditions. Using the qPCR assay, P. cubensis sporangia were detected before concentrations reached >15 sporangia/day. More than 15 sporangia/day were also observed 1 month before CDM was detected in the cucumber fields (June 2018 and 2019); however, these sporangia were identified as P. humuli using qPCR. In Michigan, information on the airborne concentration of sporangia is used to provide an early warning for growers that CDM is likely to occur (Hausbeck 2020) and prompt the application of fungicides. Using light microscopy only, P. humuli sporangia could have triggered unnecessary fungicide applications, highlighting the importance of a qPCR assay system that reliably distinguishes between P. cubensis and P. humuli.

Using light microscopy and qPCR, differences in the airborne sporangial concentrations of P. humuli and P. cubensis were detected between the 2 years of monitoring. From June to August 2018 at the commercial cucumber field, we detected higher airborne sporangial concentrations of P. cubensis compared with 2019. A relatively cold and rainy spring delayed the planting of cucumbers for pickling in 2019 (U.S. Department of Agriculture 2020). This reduced host availability may have also resulted in reduced infection and P. cubensis sporangia production. Similarly, from May to July 2018, low concentrations of P. humuli (<10 sporangia/day) were detected in the two monitored fields whereas a higher concentration (>10 sporangia/day) was observed at the three locations monitored in 2019. P. humuli overwinters in dormant hop buds or crowns, growing into expanding basal shoots in spring and early summer (Coley-Smith 1962). Extended periods of wetness, high relative humidity, and temperatures <20°C (Gent and Ocamb 2009; Royle 1973) occurred during the cold and rainy spring of 2019 (National Oceanic and Atmospheric Administration 2019) and may have favored the pathogen’s reproduction and infection.

These results suggest that the qPCR-based assay allowed for precise monitoring of airborne P. cubensis and P. humuli sporangia over two different years; the specific detection of these two species was not possible using light microscopy only. Cucumber growers in Michigan desire to know when sporangia of P. cubensis have arrived in their production region/field so that scouting efforts can be intensified, and costly fungicide programs initiated. The information on P. cubensis detection derived from spore traps coupled with qPCR could be used by growers to make informed decisions regarding fungicide usage leading to increased efficiency. Judicious use of fungicides may slow the development of pathogen resistance and decrease the cost associated with CDM control. The deployment of a broader network of spore traps and the evaluation of air samples using qPCR could also improve the risk assessment of CDM epidemics. Future evaluation of more cost-effective spore traps such as impaction traps for the monitoring of P. cubensis is essential to increase the geographic coverage of the spore trapping network in Michigan. The use of more spore traps at a local level could make the monitoring more geographically precise and trigger the execution of disease management practices only in fields at high risk of infection based on the qPCR detection of P. cubensis and the local environmental conditions.

Acknowledgments

We thank the following undergraduate research assistants: C. Adams, H. Taddonio, L. Espinoza, J. Chan, G. Kenny, and A. Job. We also thank D. Higgins for helpful discussions.