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Irrigation and Rootstocks to Manage Northern Root-Knot Nematode During Wine Grape Vineyard Establishment

    Authors and Affiliations
    • Katherine E. East1
    • Inga A. Zasada2
    • R. Paul Schreiner2
    • Michelle M. Moyer1
    1. 1Department of Horticulture, Washington State University Irrigated Agriculture Research and Extension Center, Prosser, WA 99350
    2. 2United States Department of Agriculture–Agriculture Research Service (USDA-ARS), Horticultural Crops Research Laboratory, Corvallis, OR 97330

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    Vineyard replanting in Washington state can be negatively impacted by the plant-parasitic nematode Meloidogyne hapla. Chemically focused nematode management programs do not offer long-term suppression; however, this may be achieved through the adoption of cultural approaches such as rootstocks and irrigation. Nematode-resistant rootstocks are used extensively in other regions but many have not been tested against M. hapla. Vineyards in eastern Washington are irrigated; therefore, manipulating available soil water may also impact nematode development. In 2017, two field trials were established in eastern Washington to evaluate the effects of (i) late-summer water limitation on M. hapla population development and (ii) host status of 1103 Paulsen, 3309 Couderc, and Matador rootstocks for M. hapla. The efficacy of these cultural management approaches was evaluated under three initial M. hapla densities (0, 50, and 250 M. hapla second-stage juveniles per 250 g of soil) in both trials. Reducing irrigation to manage M. hapla infestation of grape roots was ineffective and may cause harm to the vines by inducing too much water stress. Conversely, rootstocks effectively reduced population densities of M. hapla. Overall, rootstocks show the most promise as a cultural tool to manage M. hapla during the establishment phase in Washington vineyards.

    Washington state is the second-largest producer of wine grapes in the United States (USDA-NASS 2017). After a first initial planting expansion in the 1980s and 1990s, the industry is facing a period of replanting. Unfortunately, plant-parasitic nematodes can result in vineyard replant challenges (East et al. 2021; Zasada et al. 2019). This is exacerbated by the fact that wine grapevines planted in Washington state are Vitis vinifera (L.) and, unlike many other production areas around the world, are routinely planted on their own roots rather than on a resistant rootstock (Moyer and O’Neal 2014). V. vinifera is susceptible to many species of root-knot nematode (Meloidogyne spp.), including the most widespread plant-parasitic nematode in Washington state vineyards, Meloidogyne hapla (Chitwood), the northern root-knot nematode (Zasada et al. 2012). Established vines can support some level of nematode feeding but young vines planted into soils with high densities of M. hapla face intense feeding pressure and may fail to establish adequately (East et al. 2021).

    Most plant-parasitic nematode management strategies in perennial cropping systems focus on the preplanting period and emphasize the use of chemical products. However, changes and reductions in product registration have significantly reduced effective preplant chemical options available to growers (Zasada et al. 2010). This challenge, combined with grower participation in sustainable certification programs mandating reduced chemical inputs, has spurred interest in understanding the potential efficacy of cultural management tools to mitigate M. hapla damage in new (or replanted) vineyards.

    Infection of vine roots requires the second-stage juveniles (J2) of root-knot nematode to move through water in the soil to locate and infect susceptible plant roots. This research asked the question “Can available soil water be adjusted to manage M. hapla in establishing vineyards?” From our previous work in Washington, soil densities of M. hapla J2 begin to increase in late summer through the winter and peak in early spring (East et al. 2019b). The distribution of M. hapla in Washington vineyards is related to soil moisture and fine root biomass (Howland et al. 2014), with M. hapla J2 concentrated in the vine row near irrigation emitters in the top half meter of the soil profile (East et al. 2019a; Howland et al. 2014). Movement in soil by M. javanica, another species of root-knot nematode, was greatest below field capacity in laboratory experiments (–500 kPa), while root invasion by M. javanica was greatest at field capacity (–33 kPa) (Wallace 1966). Given that M. hapla J2 are concentrated under drip emitters in eastern Washington vineyards, it may be possible to reduce irrigation enough to limit densities of M. hapla J2 in the soil or reduce their ability to infest grapevine roots.

    Nematode-resistant rootstocks are another cultural tool for M. hapla management (Anwar et al. 2000). There are a number of rootstocks that are extremely poor hosts for Meloidogyne, though many have not been evaluated for M. hapla specifically (Ferris et al. 2012). In greenhouse evaluations, the rootstocks 3309 Couderc (3309 C), 101-14 Mgt, 110 R, 420 A, Salt Creek, Freedom, Harmony, Riparia Gloire, Matador, and St. George were all poor hosts for M. hapla (Zasada et al. 2019). Thus, the adoption of rootstocks in nematode-infested vineyards may be a sustainable management option for M. hapla. However, grower adoption of grape rootstocks in Washington has been historically low due to the relative ease of trunk retraining in own-rooted vines following winter cold damage. This hesitation is waning, as the industry faces an outbreak of the root pest phylloxera (Daktulosphaira vitifoliae Fitch) for which the only long-term solution is the use of resistant rootstocks (M. Hansen, personal communication). However, rootstock performance, including host status for M. hapla, is needed to provide appropriate recommendations for Washington grape growers.

    Two cultural tactics (irrigation and rootstocks) were evaluated for M. hapla management in Washington in two concurrent replant vineyard trials that were established and evaluated over 2 years. The goals were to (i) examine how late-season deficit irrigation might be used to reduce M. hapla survival and root invasion and (ii) evaluate the host status of three nematode-resistant rootstocks against M. hapla under field conditions.

    Field Site and Preparation

    Because both the irrigation and rootstock experimental vineyards used a similar experimental design, the initial steps for both trials are described together. The timeline for these experiments is described in Table 1. A 0.4-ha site was chosen at the Washington State University Irrigated Agriculture Research and Extension Center in Prosser, WA. This field had been fallow since 2014, with no prior planting records. Prior to site preparation, in spring 2017, extensive sampling confirmed that no M. hapla J2 were present in the soil. Both experiments were arranged as split-plot designs with four replicates, where each replicate was composed of three rows of vines. Each vine row was assigned one of the three initial nematode density treatments: high (approximately 250 M. hapla J2 per 250 g of soil), medium (approximately 50 M. hapla J2 per 250 g of soil), and no (0 M. hapla J2 per 250 g of soil). The proposed action threshold for M. hapla is 100 M. hapla J2 per 250 g of soil (Zasada et al. 2012). Then, within each row, three (irrigation) or four (rootstock) treatments were imposed on six-vine segments.

    TABLE 1 Experimental timeline for the irrigation and rootstock trials

    Planting rows were rototilled to a depth of 10 cm on 4 May 2017. Interrow space (1.2 m wide by 23 m long) was treated with glyphosate (23.4 ml product per liter) (RoundUp; Monsanto Company, Marysville OH) using a hand-held sprayer on 5 May 2017 to reduce weed pressure. On 10 May 2017, a large quantity of soil known to contain a high population density of M. hapla was collected under drip emitters from a 30-year-old Riesling vineyard in Mattawa, WA (East et al. 2019b). The species was confirmed as M. hapla by molecular identification done by the North Carolina Department of Agriculture & Consumer Services (Raleigh, NC). This infested soil was used to prepare M. hapla inoculum for both experiments. Soil of the same series was also collected from a nearby fallow block where M. hapla was absent, to use as the control and to mix with the infested soil to obtain the desired densities of M. hapla in the plots. Both soils were thoroughly mixed using a shovel. M. hapla J2 were extracted from infested and control soils using a semiautomatic elutriator (Seinhorst 1962) followed by sugar centrifugation-flotation (Jenkins 1964). Nematodes were enumerated using a Leica DM IL inverted microscope (Leica Microsystems, Wetzlar, Germany), so that desired densities of M. hapla could be established in different treatment plots by altering the ratio of the infested and control soil additions made. Three 250-g samples composed from three soil cores were evaluated for each soil type. On average, the infested soil had 1,180 ± 87 M. hapla J2 per 250 g of soil, and the control soil had 0 ± 0 M. hapla J2 per 250 g of soil.

    Initial nematode densities were established in entire rows at planting, with each nematode density (no, medium, and high, as described above) randomly allocated to specific rows in 3-row replicates, for a total of 12 rows per experiment (three densities by four replicates). At planting, each vine received 2.5 liters of the appropriate inoculum soil (Fig. 1A and B).

    FIGURE 1

    FIGURE 1 Vineyard planting and destructive harvesting. A, Soil with Meloidogyne hapla (in bucket), being applied in predug hole before planting. B, Rootstock experimental site preplanting, with trenches dug. C, Irrigation experimental site on day of planting. Each row has three irrigation lines, one for each irrigation treatment. D, Planted irrigation experiment with vines at 1.2-m spacing. E, Fully planted rootstock trial, with nursery spacing. F, Vitis vinifera ‘Chardonnay’ plugs in tray (irrigation study; grafted vines used for rootstock study). G, Destructive harvest in fall 2018, digging full vines. H, Whole vines harvested and awaiting transport to be washed and weighed.

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    Irrigation experiment planting.

    Vines were planted on 26 May 2017. Each row consisted of three plots with 6 vines each (irrigation treatments described below) for a total of 18 vines per row. Vines were spaced 1.2 m apart within each plot, and plots were spaced 1.8 m apart. All irrigation treatments were represented in each row (with varying initial nematode density). Each treatment per row had an individual dripline with an on/off valve allowing independent irrigation for each plot with one drip emitter per vine (Fig. 1C and D).

    Irrigation treatments were as follows: (i) full irrigation (full; irrigated to keep soil water potential above –100 kPa), (ii) slight midseason reduction (partial; starting approximately 1 August, irrigated back to field capacity when soil water potential reached –500 to –600 kPa), and (iii) extreme midseason reduction (low; starting approximately 1 August, irrigated back to field capacity only when soil water potential was less than –1,000 kPa). Timing of irrigation cut-off was designed to coincide with the time that M. hapla J2 population densities in the soil increase rapidly (East et al. 2019b). Dielectric soil water potential sensors (MPS-6; Decagon Devices, Pullman, WA) were used to measure soil water potential and schedule irrigation. Three soil water potential sensors were placed at 20 cm below the soil surface under a drip emitter, one per irrigation treatment in the two outermost rows, for a total of six sensors. Average soil water potential was measured at least twice a week; when average soil water potential reached a particular treatment threshold, those treatment plots were watered, returning them to field capacity (–33 kPa). All plots were irrigated to field capacity prior to irrigation shut off (19 October 2017) to maintain sufficient moisture going into winter.

    Rootstock experiment planting.

    The rootstock experiment was planted on 30 May 2017. Twelve rows, spaced 1.2 m apart, were prepared as described in the irrigation experiment. Within each row, 4 rootstock treatment plots were established with six vines each on 0.3-m spacing, for a total of 48 plots (Fig. 1E). The 0.3-m spacing was used because we were less concerned with water movement between vines. All rootstocks had a V. vinifera ‘Chardonnay’ scion (Fig. 1F), and rootstock treatments consisted of (i) Matador; (ii) 1103 Paulsen (1103 P); (iii) 3309 C; and (iv) an own-rooted Chardonnay (control). All rootstocks were irrigated on the same schedule as the full-irrigation treatment in the irrigation experiment.

    Vine and nematode data collection.

    Vine data were collected in both trials in the same way. In fall 2017 (24 to 25 October), every other vine in each treatment plot in both experiments was destructively harvested to measure M. hapla egg density, fine root density, plant root weight, shoot weight, and trunk weight. Each vine was removed in-field using shovels, and most of the soil was removed (Fig. 1G and H). In the lab, each vine was stripped of leaves and roots were carefully washed free of soil under cold running water. Three root branches emanating from the trunk that included woody roots and fine feeder roots were selected at random and removed, and all nine root branches were combined per plot. These root branches were retained in cold storage for later processing to determine M. hapla egg density and root fresh weight (described below). The vine was then divided into shoots (1-year-old growth), the remaining roots (all tissue below the uppermost root, including the crown), and trunk (remaining vine tissue). Each vine portion was placed in a paper bag, weighed (fresh weight), placed in a drying oven at 70°C for 7 days, and then reweighed (dry weight). This process was repeated in fall 2018 (2 to 3 October) with the remaining in-field vines. Pruning weights were collected during the 2017–18 winter (26 February 2018) between year 1 and year 2 of the experiments. Vines were pruned back to two buds and the canes that were removed were collected and weighed.

    Sampling for nematodes was similar in both experiments. In 2017, soil samples were collected on 20 October, with one soil core (2.5 cm in diameter by 45 cm deep) collected per vine from between the drip emitter and the vine, for a total of six cores combined per plot. This process was repeated on 27 February and 27 September 2018, with two soil cores collected per vine and combined into one sample, because there were only three vines per plot in 2018. Nematodes were extracted and quantified as described above. M. hapla eggs were collected from the root branches excised from each three-vine sample during fall destructive harvest in 2017 and 2018 (described above). Roots were placed into a screw-top container with a 10% sodium hypochlorite solution and shaken for 3 min to extract eggs (Hussey and Barker 1973). The solution was poured through stacked 88- and 25-µm sieves, with roots retained on the 88-µm sieve being aggressively washed. Roots were recovered from the 88-µm sieve and eggs were rinsed from the 25-µm sieve to a 50-ml tube using a total of 10 ml of distilled water. Eggs from a 1-ml aliquot were counted under a stereomicroscope and data were expressed as number of M. hapla eggs per gram of wet root.

    Data analysis.

    Parameters in both rootstock and irrigation experiments were analyzed using the standard least-squares model platform in JMP (version 14.0.0; SAS Institute Inc., Cary, NC). M. hapla J2 data were log transformed (log x + 1) to meet assumptions of variance. Irrigation experimental parameters were analyzed as a split plot, with irrigation, nematode density, and nematode density-irrigation interaction as fixed variables and replicate as a random variable. Rootstock experimental parameters were analyzed as a split plot, with rootstock, nematode density, and nematode density–rootstock interaction as fixed variables and replicate as a random variable. Each variable (J2, eggs, dry weight, and pruning weight) collected on a given date was analyzed separately. When appropriate, means separation was performed using Tukey’s honestly significant difference test. There were no significant (P < 0.05) interactions between irrigation and initial nematode density treatments for all data, except that of egg density in 2017 in the irrigation study. No interactions were significant between rootstock and initial nematode density treatments in the rootstock study; therefore, we focused on main effects.

    Irrigation had no effect on M. hapla population densities.

    Irrigation manipulation in late summer had no effect on M. hapla J2 densities (P = 0.27, 0.63, and 0.74 in fall 2017, spring 2018, and fall 2018, respectively). Similarly, there was no difference in M. hapla eggs per gram of root between irrigation treatments in fall 2017 or fall 2018 (P = 0.84 and 0.62, respectively). The aim of reducing irrigation during late summer was to reduce the water film around soil particles or roots to limit M. hapla J2 movement to roots and root invasion. This approach was not successful. Our results are similar to those of Griffen and Jorgensen (1969), where they examined infection of potato (Solanum tuberosum) by M. hapla under soil moisture levels of 25 to 125% field capacity. Over 70% of tubers were infected at all soil moisture levels, with the greatest infectivity at 100% field capacity. In a study by Towson and Apt (1983), survival of M. javanica was reduced whenever soil was either very wet (–16 to –30 kPa; field capacity equal to –33 kPa) or very dry (–1,500 to –9,200 kPa; permanent wilting point equal to –1,500 kPa). The optimal soil moisture for M. javanica survival was –110 kPa. Reducing soil water content low enough to restrict M. hapla J2 invasion of roots or survival in soil may not be possible in young vines without causing too much water stress, because vines exhibited early leaf senescence in the low irrigation treatment applied here (irrigating at –1,000 kPa soil water potential), though there were no effects on vine biomass.

    As expected, high and medium initial nematode density treatments had greater M. hapla J2 soil densities than the no initial nematode density treatment in fall 2017 and 2018 (P = 0.0002 and P < 0.0001, respectively) (Table 2). The effect was most pronounced in spring 2018 (P < 0.0001). In fall 2017, the high initial nematode density treatment had more eggs than the no initial nematode density treatment, and medium initial nematode density treatment was intermediate (P < 0.0001). By fall 2018, high and medium initial nematode density treatments were no longer different from each other, and only differed from the no initial nematode density treatment (P < 0.0001).

    TABLE 2 Irrigation experiment: Influence of preplant Meloidogyne hapla soil density on second-stage juvenile (J2) soil densities, egg densities, and whole-vine dry weights of own-rooted Vitis vinifera ‘Chardonnay’ over 2 years postplantingx

    Keeping soil water near field capacity resulted in greater vine pruning weights regardless of nematode density.

    There were no differences in whole vine dry weight in fall 2017 due to irrigation (P = 0.16) or initial nematode density (P = 0.21; Table 2). In fall 2018, there was no effect of irrigation (P = 0.79) but there was an effect of initial nematode density (P = 0.039; Table 2). The no initial nematode density treatment had greater dry weights than the medium initial nematode density treatment but neither differed from the high initial nematode density treatment. It is difficult to draw any conclusions from this variation without additional years of data. Irrigation altered dormant season pruning weight in spring 2018, with vines under full irrigation having greater pruning weights than those under partial or low irrigation (P = 0.0009; Table 2). There was no effect of initial nematode density treatment on pruning weights (P = 0.33).

    Both the low and partial irrigation treatments supplied less water than desired for early establishment of vines (Evans et al. 1993; Myburgh et al. 1996; Peacock et al. 1977). In the low irrigation treatment, yellowing of leaves associated with water stress was observed 2 weeks after initiating the lowest irrigation treatment. This study was not designed to determine water needs for a young vineyard; rather, it was designed to see whether reducing irrigation could reduce nematode invasion while maintaining vine growth.

    Rootstocks reduced M. hapla population densities.

    In fall 2017, M. hapla J2 population densities in soil were influenced by rootstock (P = 0.0003; Table 3) but not by the initial nematode density (P = 0.058). The own-rooted control vines had the highest density of M. hapla J2, with 1103 P, 3309 C, and Matador having very low M. hapla J2 densities in soil. In spring 2018, there were significant differences among rootstock (P < 0.0001) and initial nematode density (P < 0.0001) treatments. All rootstocks had fewer M. hapla J2 than the own-rooted control, and the high and medium initial nematode density treatments had more M. hapla J2 than the no initial nematode density treatment. Similarly, in fall 2018, both rootstock and initial nematode density treatments were significant parameters (P = 0.0023 and P < 0.0001, respectively). As of spring 2018, high and medium initial nematode density treatments had the greatest M. hapla J2 densities; however, these appeared to be mostly driven by the own-rooted Chardonnay. M. hapla had higher reproduction on susceptible Chardonnay than the three non-vinifera rootstocks, meaning the non-vinifera rootstocks are likely poor hosts for M. hapla. However, in fall 2018, Matador was not different from the control, which indicates that this rootstock may allow for reproduction of M. hapla, though to a lesser extent than a susceptible cultivar in the first year after planting. Whether this rootstock would continue to be a poor host for M. hapla over the lifetime of a vineyard is unknown.

    TABLE 3 Rootstock experiment: Influence of preplant Meloidogyne hapla soil density on second-stage juvenile (J2) soil densities over the 2 years of the study

    There was a 40- to 50-fold difference between M. hapla eggs per gram of root found on the own-rooted Chardonnay versus any of the non-vinifera rootstocks in fall 2017 and fall 2018. In fall 2017, rootstock, initial nematode density, and their interaction significantly altered M. hapla egg densities (P ≤ 0.0001, 0.0013, and 0.0001, respectively) (Fig. 2A). The own-rooted Chardonnay initial high and medium nematode density treatments had the greatest density of eggs/g of root (mean = 2,299 and 1,905 eggs/g of root, respectively), while all other treatment combinations had lower densities of M. hapla eggs (ranging from 0 to 58 eggs/g of root, on average). The differences between the two densities on the susceptible rootstock were driven by the significance of the interaction, though that effect was lost by the next fall.

    FIGURE 2

    FIGURE 2 Meloidogyne hapla egg densities per gram of root by initial nematode density (no, medium, and high) and rootstock (1103 Paulsen [1103 P], 3309 Couderc [3309 C], Matador, and own-rooted Vitis vinifera ‘Chardonnay’ control) in A, fall 2017. M. hapla egg densities per gram root in fall 2018 are shown separately by B, rootstock and C, initial nematode density. Bars are the means of n = 4 and error bars are standard error. Different letters denote significant differences between treatment means at α = 0.05 using Tukey’s honestly significant difference.

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    In fall 2018, both rootstock and initial nematode density treatments significantly altered egg production (P < 0.0001 and 0.014, respectively). Again, own-rooted Chardonnay had more M. hapla eggs than any rootstock (Fig. 2B), and the high initial nematode density treatment had more M. hapla eggs than the no initial nematode density treatment, with the medium initial treatment intermediate between them (Fig. 2C). The final nematode population differences imposed through the initial nematode density inoculations persisted, but only in the susceptible cultivar. The non-vinifera rootstocks performed equally well under a range of nematode pressures but no rootstock was free of M. hapla. Host status can be determined by calculating the percentage of eggs per gram of root, where less than 10% is considered a resistant host (Ferris et al. 2012). In our study, under this definition, 1103 P (2.3%), 3309 C (0.3%), and Matador (0.2%) are all considered resistant to M. hapla after 2 years and could be recommended as rootstocks for M. hapla management.

    For comparison with what might elicit a management action in production vineyards, M. hapla J2 densities were organized into risk categories around the proposed management threshold of 100 M. hapla J2 per 250 g of soil (East et al. 2021) (Fig. 3). In fall 2018, all of the high and medium initial nematode density treatments, regardless of rootstock, were in category 2 or higher, where management action should be considered. Currently, the management approach for nematodes in eastern Washington vineyards heavily relies on preplant soil fumigation, in which nematode density and treatment options would be considered prior to planting, although recent studies show that preplant fumigation approaches are not always effective (East et al. 2021). Postplanting, there are few nematicide options with inconsistent efficacy; ultimately, growers of own-rooted V. vinifera are really limited to preplant chemical management of nematodes, with few options when populations reestablish at a site.

    FIGURE 3

    FIGURE 3 Categorical diagram of Meloidogyne hapla management risk over time. M. hapla second-stage juvenile (J2) density categories: category 1 (blue) = less than 50 M. hapla J2 per 250 g of soil (below management threshold); category 2 (yellow) = 50 to 150 M. hapla J2 per 250 g of soil (around proposed management threshold); and category 3 (red) = more than 150 M. hapla J2 per 250 g of soil (above management threshold) in fall 2017, spring 2018, and fall 2018. Initial M. hapla density treatments were applied at planting in spring 2017: High (approximately 250 M. hapla J2 per 250 g of soil); medium (Med; approximately 50 M. hapla J2 per 250 g of soil); and No (0 M. hapla J2 per 250 g of soil). Rootstocks were: 1103 Paulsen (1103 P), 3309 Couderc (3309 C), Matador, and own-rooted Vitis vinifera ‘Chardonnay’ (control). Each data point within a treatment combination is the average of n = 4.

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    Historically, 1103 P was found to be susceptible to M. incognita and resistant to some other Meloidogyne spp. (Nicol et al. 1999; Téliz et al. 2007). The 3309 C rootstock was considered susceptible to Meloidogyne spp. (Cousins and Walker 2002; Nicol et al. 1999) but now it has also been reported to be tolerant of Meloidogyne spp., where it can develop and reproduce but have no impact on vine growth (McKenry and Anwar 2006). The 3309 C rootstock has also been reported as a poor host for M. hapla in greenhouse studies (Zasada et al. 2019). Our trials confirm the poor host status of both 1103 P and 3309 C, with both rootstocks supporting some nematode reproduction but only after 2 years of exposure to high nematode densities. The own-rooted Chardonnay control was always at the management action recommended level (category 3) in the high and medium initial nematode density treatments during both years.

    Matador is a relatively new rootstock and was designed to be resistant to ‘Harmony’-resistance breaking pathotypes of Meloidogyne sp. (Cousins 2011) and was also reported as a poor host for M. hapla in greenhouse studies (Zasada et al. 2019). In this study, at the high initial nematode density, Matador supported M. hapla densities that fell into category 2 (i.e., requiring potential management intervention) at the earliest sampling time up until fall 2018, unlike the other two rootstocks, which did not support M. hapla densities that would elicit management action (category 1). After 2 years under high nematode pressure, based on egg density, Matador is defined as a poor host for M. hapla; however, looking at J2 densities and relative vine growth, it appears to actually be tolerant; in other words, although it does allow for M. hapla development, the plant vigor was not impacted in this short study. Looking at both sets of information, we would be confident in recommending 1103 P and 3309 C as rootstocks to manage M. hapla but more cautious about recommending Matador without further study.

    As an aside, we did find M. hapla in the no initial nematode density treatment in fall 2017. In this case, M. hapla J2 were only found in three of the eight total plots designated as “no initial nematodes”, including replicate plots planted to 1103 P and Chardonnay; all other no initial nematode density plots had zero nematodes. Later samples from those same plots had zero or almost zero nematodes (about 20 times fewer), indicating that the fall 2017 samples were likely contaminated with nematodes in the lab rather than in the field. Field contamination would have resulted in increased or similar M. hapla J2 densities over time, which was not the case (Fig. 3).

    Rootstocks had greater vine vigor than own-rooted vines.

    There were no differences in whole vine dry weight in either year due to initial nematode density (P = 0.089 and 0.44) but rootstock clearly had an effect (P < 0.0001 and P = 0.0004 in fall 2017 and 2018, respectively) (Table 3). In fall 2017, own-rooted Chardonnay were smaller than the non-vinifera rootstocks. In fall 2018, own-rooted Chardonnay were still the smallest vines but were not different from 3309 C; Matador was intermediate in size, and 1103 P had the most dry mass. Rootstock also influenced pruning weights (P < 0.0001), and own-rooted Chardonnay had smaller pruning weight than all rootstocks.

    Given that the Chardonnay scion grafted on all three non-vinifera rootstocks had greater pruning weights than the own-rooted Chardonnay, this could indicate that planting on rootstocks may impart more vigor to the vines than planting own-rooted V. vinifera. Increased vigor may be a benefit to using nematode-tolerant rootstocks in Washington, because this may lead to quicker establishment of vines and a potentially earlier first crop. In producing vineyards in Washington state, vigor management is mostly achieved using deficit irrigation strategies, which should work equally well to manage the high genetic vigor in vineyards planted to rootstocks once they are established (Keller et al. 2012).

    The need for longer-term field trials in nematode resistance evaluations.

    The gradation in M. hapla densities (0 to 250 M. hapla J2 per 250 g of soil) established at planting persisted over the duration of both the irrigation and rootstock field experiments but had little effect on vine biomass (whole vine biomass or pruning weight). Only in the fall 2018 irrigation experiment was there decreased vine biomass in the medium initial nematode density treatment as compared with the high and no initial nematode density treatment (Table 2). In a perennial cropping system, it may take more than two growing seasons to see an effect of M. hapla parasitism on vine productivity. Because all of the vines in these experiments were harvested in 2 years, as designed, a third year of data could not be collected. In a concurrent long-term field trial in Washington state, dormant vine pruning weights of own-rooted Chardonnay vines did not differ from vines planted on Meloidogyne spp. resistant rootstocks until 3 years after planting (East et al. 2021), indicating that the impact of M. hapla parasitism on grape productivity may not be expressed in the first few seasons. In experiments with ring nematode Mesocriconema xenoplax, reduction in V. vinifera ‘Pinot Noir’ vine biomass did not occur in the first year but was observed by year two in a potted vine experiment (Schreiner et al. 2012) and by the third year in field microplots (Pinkerton et al. 2004). This does not invalidate this study but does indicate that the results should be considered in the larger context. Future field trials examining the impact of Meloidogyne hapla in perennial cropping systems should be designed to run for many years, in order to observe a measurable impact if the intent is to monitor plant decline due to nematode parasitism.

    Conclusions and Recommendations

    The irrigation treatments had no effect on M. hapla J2 soil or egg densities or on whole-vine biomass over the course of the 2-year trial. However, fully irrigated vines had greater pruning weights than the partial or low irrigation treatments. Overall, this indicates that water limitation during the late summer is not an effective tactic to modify M. hapla population densities in newly planted vineyards and may do more harm than good during vineyard establishment.

    Rootstocks are a good option to manage M. hapla. All three rootstocks considered were poor hosts for M. hapla, though Matador may be less resistant than 3309 C and 1103 P. Although nematodes were capable of reproducing on the tested rootstocks, reproduction was 40 to 50 times less than on own-rooted Chardonnay. Matador and 1103 P rootstocks had greater pruning weights and whole-vine biomass in both years than own-rooted vines, though there was no effect of the different initial nematode density treatments, indicating that these rootstocks are more vigorous than V. vinifera under eastern Washington conditions. Greater vigor due to rootstock could be managed using cultural practices (irrigation) and may even be desired in some cases. All three rootstocks are recommended for replanting vineyards in M. hapla-infested soil, though Matador may need further evaluation for long-term performance.

    Finally, it may not be feasible to understand the impact of M. hapla from short-term studies for vineyards that might be in production for many decades. Longer-term studies are needed to gain a more realistic view of how management tools such as rootstocks fare over time in mitigating nematode parasitism.


    We thank A. Boren, M. McCoy, J. Newhouse, C. Wram, M. Mireles, A. McDaniel, and J. Pinkerton for their technical assistance.

    The author(s) declare no conflict of interest.

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    Funding: Funding for this project has been provided through the Washington State Grape and Wine Research Program; funding sources include Washington State Wine Commission, Auction of Washington Wines, State Liter tax, and Washington State University Agriculture Research Center; with partial support from USDA-ARS Current Research Information System 2072-22000-043-00D and USDA National Institute of Food and Agriculture Hatch project 1016563.

    The author(s) declare no conflict of interest.