Plant-parasitic nematodes associated with the root zone of hop cultivars planted in a Florida field soil

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Plant-parasitic nematodes associated with the root zone of hop cultivars planted in a Florida field soil

Tristan T. Watson / Marco Suarez / Zhanao Deng / Johan A. Desaeger *

Keywords : Hop cultivars, Humulus lupulus , Meloidogyne javanica , Nematode management, Pratylenchus brachyurus

Citation Information : Journal of Nematology. Volume 52, Pages 1-10, DOI: https://doi.org/10.21307/jofnem-2020-040

License : (CC-BY-4.0)

Received Date : 19-September-2019 / Published Online: 23-April-2020

ARTICLE

ABSTRACT

In early 2016, hop plants were introduced into Florida. By late 2016, the hop plants were showing stunted growth and were heavily parasitized by Meloidogyne javanica. In this study, we determined host susceptibility of 14 hop cultivars to M. javanica in a greenhouse experiment and monitored population development of plant-parasitic nematode species in the root zone of 17 hop cultivars planted in three newly established hop yards in Florida. Plant-parasitic nematodes in the rooting zone soil of field grown hop plants included M. javanica, Pratylenchus brachyurus, Paratrichodorus minor, Belonolaimus longicaudatus, Xiphinema setariae/vulgare complex, Mesocriconema xenoplax, and Helicotylenchus dihystera; however, soil population densities of P. minor, B. longicaudatus, X. setariae/vulgare complex, M. xenoplax, and H. dihystera remained low through the study. Root galling, M. javanica egg production, and soil population densities of M. javanica were consistently large on the ‘Canadian Red Vine’, ‘Centennial’, ‘Chinook’, and ‘Comet’ cultivars, and small on the ‘Galena’ and ‘Triple Perle’ cultivars. No differences were observed in soil population densities of P. brachyurus among hop cultivars. Overall, our study provides the first report of plant-parasitic nematode population development in the root zone on hop cultivars planted in Florida.

Graphical ABSTRACT

Hop cones from the hop plant (Humulus lupulus L.) are a fundamental ingredient of the brewing process, imparting bitterness, aroma, and flavor to beer. In the United States, hop cultivation began when the first colonialists arrived in the 1600s (Turner et al., 2011). By the late 1800s, hop production spread to states in the Pacific Northwest (Steiner, 1973). Modern hop production is largely concentrated in Washington, Oregon, and Idaho, where favorable day length, climate, and low disease pressure allow for production of excellent quality cones and high yields; however, many other regions are now exploring hop as an alternative crop. The United States is the world’s leading hop producer, responsible for 41.3% of the crop produced worldwide (Hope Growers of America, 2018). Some of the most commonly planted hop cultivars include Citra®, Cascade, and Centennial (USDA-NASS 2018).

Although several bacterial, fungal, and viral diseases have been documented on hop (Mahaffee et al., 2009), there are relatively few reports on the plant-parasitic nematode species associated with this crop. The hop cyst nematode (Heterodera humuli Filipjev) is the most well studied nematode associated with hop worldwide (De Grisse and Gillard, 1963), and has been reported in United States hop production in Pierce County, Washington (Cobb, 1962). Other plant-parasitic nematodes associated with hop in the United States include Meloidogyne hapla Chitwood and Xiphinema americanum Cobb, which were found widely distributed within hop yards in Sacramento County, California (Maggenti, 1962), as well as Pratylenchus neglectus Filipjev & Schuurmans-Stekhoven and Pratylenchus thornei Sher & Allen, which have been reported less frequently within Idaho and Oregon (Hafez et al., 1992). In New Zealand, Ditylenchus destructor Thorne was isolated from severely stunted plants from commercial hop yards (Foot and Wood, 1982). Severe nematode damage in a hop yard in Belgium was associated with H. humuli, Tylenchorhynchus dubius Bütschli, Pratylenchus penetrans Filipjev & Shuurmans Stekhoven, and Helicotylenchus sp. (Pelerents et al., 1983). In China, Meloidogyne incognita Kofoid & White was reported on the hop cultivar Comet; however, nematode population development was restricted in a locally bred cultivar (Zhang and Zheng-Li, 1989).

As a result of the high demand for locally produced hop from the micro-brewing industry, hop plants were recently introduced into Florida as an alternative crop. In early 2016, the University of Florida planted hop rhizomes at the Gulf Coast Research and Education Center in Hillsborough County, Florida to evaluate the potential of this crop. By late 2016, some of the hop plants were showing stunted growth and yellow leaves. Uprooted plants exhibited severe root galling, and females extracted from galls, as well as J2s extracted from soil, were subsequently identified as Meloidogyne javanica (Brito et al., 2018).

In Florida, root-knot nematodes (Meloidogyne spp.) are widespread and likely pose a significant obstacle to any hop yard planted in the state. Many other plant-parasitic nematode species that are abundant in the region may also pose a threat to hop production. Using a combination of field and greenhouse experiments, the objectives of this study were to: (i) determine host susceptibility of 14 cultivars of hop to M. javanica in a greenhouse experiment, and (ii) monitor population development of plant-parasitic nematode species in the root zone of 17 cultivars of hop planted in three newly established hop yards in Florida.

Materials and methods

Site description

The studies were conducted at the Gulf Coast Research and Education Center research farm in Wimauma, Florida, United States. Soil at the site is an Arredondo fine sand (95% sand, 3% silt, 2% clay). The field had been fallow for over 10 years under natural grass cover prior to conducting the following trials.

Hop yard planting No. 1

In 2016, a new hop yard was established at the research farm to evaluate plant-parasitic nematode population development on 10 hop cultivars from two planting material sources. Planting rows were 82-m long and were spaced 4.88-m apart, with a 0.91-m wide strip of black geotextile mulch covering the center of each row. Hills were spaced 0.91-m apart within the planting row. A 6.1-m tall trellis wire was installed above each row. Plants were irrigated with two drip tapes (30-cm emitter spacing), each located 0.46-m feet away from the row center along both sides of the planting row. Plants were fertilized with all-purpose fertilizer (20:20:20) at a rate of 149 kg N/ha/season. Ground cover between rows consisted of perennial ryegrass (Lolium perenne) that was mowed periodically throughout the growing season.

The experimental design was a randomized complete block with three blocks and 12 cultivar treatments. In total, 36 plots were overlaid on to three planting rows within the hop yard. Plots consisted of four hills (3.66-m long) and were planted at a density of two rhizomes per hill. A 0.91-m buffer zone was located between plots to minimize edge effects. On April 12, 2016, triplicate plots were planted with rhizomes from each of the following cultivars: (i) ‘CTZ’, (ii) ‘Cascade’ (source: Oregon), (iii) ‘Cascade’ (source: Washington), (iv) ‘Centennial’ (source: Oregon), (v) ‘Centennial’ (source: Washington), (vi) ‘Chinook’, (vii) ‘Fuggle’ (source: Oregon), (viii) ‘Magnum’ (source: Washington), (ix) ‘Nugget’ (source: Washington), (x) ‘Sorachi Ace’ (source: Washington), (xi) ‘Triple Perle’ (source: Washington), or (xii) ‘Willamette’ (source: Washington).

Plants were harvested in August 2016 and December 2016 by cutting the bines by hand at ground level. During the growing season, a number of the cultivars displayed symptoms of a viral disease, which was subsequently confirmed as Apple Mosaic Virus (ApMV). Due to the distribution and severity of the symptoms the hop yard was scheduled to be renovated using clean planting material in 2017.

In December 2016, plants were uprooted from the soil after harvest using a shovel and assessed for root galling. Entire root systems were assessed for galling and given a root gall index rating on a scale from 0 to 10 (Bridge and Page, 1980), with 0 indicating no visible galling and 10 indicating 100% galled. The abundance of plant-parasitic nematodes in soil was also determined for each plot. Eight soil cores (20 cm in length, 2.5 cm in diameter) were obtained from each plot directly from the previous rooting zone (two soil cores per hill). Soil samples were placed into plastic bags and were stored at 4°C for a maximum of 14 days prior to subsequent processing. Nematodes were extracted from a 200-mL subsample of soil from each plot using the Baermann pan technique (Forge and Kimpinski, 2007), with a two-day incubation period. After collecting the nematodes over a 25-µm sieve, the nematodes were transferred in water into plastic scintillation vials and store at 4°C for a maximum of 14 days prior counting on an inverted compound microscope.

Hop yard planting No. 2

In 2017, a new hop planting was established in the previously renovated hop yard. On August 15, 2017, metam potassium (K-Pam® HL; AMVAC Chemical Corporation, Los Angeles, CA) was drip applied to the previous planting rows at a rate of 584 L/ha. The experimental design was a randomized complete block with three blocks and 12 cultivar treatments overlaid on the previous plots from the first field experiment in 2016. Each hill within a plot was planted with two transplants. On September 13, 2017, triplicate plots were planted with tissue-cultured transplants from each of the following cultivars: (i) ‘Cascade’ (source: Washington), (ii) ‘Cascade’ (source: Florida), (iii) ‘Cashmere’, (iv) ‘Centennial’ (source: Washington), (v) ‘Centennial’ (source: Florida), (vi) ‘Chinook’ (source: Washington), (vii) ‘Chinook’ (source: Florida), (viii) ‘Comet’ (source: Washington), (ix) ‘Magnum’ (source: Washington), (x) ‘Nugget’ (source: Washington), (xi) ‘Triple Perle’ (source: Washington), or (xii) ‘Zeus’ (source: Washington).

In February 2018, artificial lights (Philips Flowering Lamp 2.0; Philips, Amsterdam, the Netherlands) were installed every 2.74 m along the trellis wire to extend the daylight length by 5 hr from March to May and from September to November. Plants were harvested by hand in December 2017, July 2018, December 2018, and July 2019 by cutting the bines at ground level.

The abundance of plant-parasitic nematodes in soil was determined on four sampling dates (December 2017, July 2018, December 2018, and July 2019). Soil was sampled and nematodes were quantified as described previously.

Hop yard planting No. 3

In 2017, an additional new hop planting was established on a single planting row adjacent to the previous hop yard from 2016. On August 15, 2017, metam potassium was drip applied to the planting row, as described above. Planting rows were covered with a 0.91-m wide strip of metallic plastic mulch. The experimental design was a randomized complete block with four blocks and five cultivars overlaid on the previous plots from the first field experiment in 2016. Each hill within a plot was planted with two transplants. On September 15, 2017, plots were planted with tissue-cultured hop transplants of each of the following cultivars sourced from Washington: (i) ‘Canadian Red Vine’, (ii) ‘Galena’, (iii) ‘Mt. Rainier’, (iv) ‘Tahoma’, or (v) ‘Willamette’.

Artificial lighting was installed every 2.74 m along the trellis wire to extend the daylight length during March to May and September to November. Plants were harvested by hand in December 2017, July 2018, December 2018, and July 2019 by cutting the bines at ground level.

The abundance of plant-parasitic nematodes in soil was determined on three sampling dates (December 2017, July 2018, December 2018, and July 2019). Six soil cores were obtained from each plot directly from the previous planting holes (two cores per hill). Soil samples were processed, and nematodes were quantified as described previously.

Greenhouse experiment

In 2017, a greenhouse experiment was performed to evaluate the susceptibility of 14 hop cultivars to M. javanica. The experimental design was a randomized complete block with six blocks and 14 cultivar treatments. In total, 84 20-cm diameter circular pots were filled with 3.5 L of soil mix (three parts steam sterilized field soil: two parts potting mix). The nematode inoculum was prepared by extracting eggs from a population of M. javanica maintained in a greenhouse on tomato roots using the sodium hypochlorite technique (Hussey and Barker, 1973), followed by washing the eggs in water over a 25-µm sieve. The eggs were inoculated as 1-mL of nematode egg suspension (2,500 eggs/mL) into each of four 1-cm deep holes made in the top of the soil in each pot (10,000 eggs/pot). After nematode inoculation, six replicate pots were planted on September 18, 2017 with a single tissue-cultured hop transplant from one of the following cultivars sourced from Washington: (i) ‘Canadian Red Vine’, (ii) ‘Cascade’, (iii) ‘Cashmere’, (iv) ‘Centennial’, (v) ‘Chinook’, (vi) ‘Comet’, (vii) ‘Galena’, (viii) ‘Magnum’, (ix) ‘Mt. Rainier’, (x) ‘Nugget’, (xi) ‘Tahoma’, (xii) ‘Triple Perle’, (xiii) ‘Willamette’, or (xiv) ‘Zeus’. The pots were irrigated as required with a drip irrigation system. Pots were fertilized biweekly with all-purpose fertilizer (20:20:20), with a cumulative application of 0.45 g of mineral N supplied to each plant. The plants were grown in a greenhouse for 12 weeks prior to subsequent analysis.

At harvest, plants were carefully uprooted from the soil and assessed for root galling, as described above. The abundance of eggs on each root system was determined by extracting eggs from entire root systems using the sodium hypochlorite technique, as described above. Eggs were collected over a 25-µm sieve, transferred in water into plastic scintillation vials, and store at 4°C for a maximum of 14 days prior counting on an inverted compound microscope.

Molecular identification of plant-parasitic nematodes

In July 2019, dagger (Xiphinema sp.), lesion (Pratylenchus sp.), ring (Mesocriconema sp.), spiral (Helicotylenchus sp.), and stubby-root (Paratrichodorus sp.) nematodes extracted from the root zone of hop plants were identified to the species level by sequencing a partial fragment of the 28S ribosomal RNA gene region and the 18S ribosomal RNA gene region. For each genus, one to seven vermiform nematodes were plucked from the July 2019 sample nematode extractions and combined into a single sample for DNA extraction. Total genomic DNA was extracted using the sodium hydroxide nematode digestion protocol (Floyd et al., 2002). The D2-D3 expansion segments located within the 28S ribosomal RNA gene region was amplified using the primers D2A (5′-ACAAGTACCGTGAGGGAAAGTTG-3′) and D3B (5′-TCGGAAGGAACCAGCTACTAGAT-3′). The 18S ribosomal RNA gene region was amplified using the 1813/2646 primer set (Holterman et al., 2006). Amplification reactions contained 2.5 μL of 10X ThermoPol® Buffer (New England Biolabs, Beverly, MA, USA), 0.5 μL of 10 mM dNTPs, 0.5 μL of 25 mM MgCl2, 0.5 μL of 10 μM forward and reverse primers, 1.0 μL of nematode digestion solution, and 0.125 μL of Taq DNA Polymerase (New England Biolabs, Beverly, MA, USA), brought to a volume of 25 μL. The 28S ribosomal RNA gene region was amplified using the following temperature profile: 5 min at 95°C, 35 cycles (30 sec at 94°C, 45 sec at 55°C, and 1 min at 68°C), followed by a final extension of 5 min at 68°C. The 18S ribosomal RNA gene region was amplified using the following temperature profile: 5 min at 95°C, 5 cycles (30 sec at 94°C, 30 sec at 45°C, and 70 sec at 68°C), 40 cycles (30 sec at 94°C, 30 sec at 54°C, 70 sec at 68°C) followed by a final extension of 5 min at 68°C. All amplification reactions were carried out on a T100 Bio-Rad Thermal Cycler (Bio-Rad, CA, USA), sequenced in both directions by GENEWIZ DNA sequencing services (GENEWIZ Inc., South Plainfield, NJ, USA), and assembled using Geneious Prime v. 2019.2.1 (Biomatters, Auckland, New Zealand). Consensus sequences were compared for similarities within the National Center for Biotechnology Information GenBank database using the Basic Local Alignment Search Tool.

Statistical analysis

Nematode data from the field and greenhouse experiments were subjected to a one-way ANOVA in SAS Studio (SAS University Edition; version 3.3; SAS Institute Inc., Cary, NC, USA) using the PROC GLM procedure. Differences among treatment means were examined using Tukey’s HSD test (p < 0.05).

Results

Nematode species recovered from the rooting zone soil of hop cultivars

Plant-parasitic nematode species recovered from the root zone of hop plants grown in a Florida field soil during the three-year trial period included M. javanica, Pratylenchus brachyurus Filipjev & Schuurmans-Stekhoven, Paratrichodorus minor Siddiqi, Belonolaimus longicaudatus Rau, Xiphinema setariae/vulgare complex Luc, Mesocriconema xenoplax Loof & De Grisse, and Helicotylenchus dihystera Sher (Table 1). Throughout the study, soil population densities of P. minor, B. longicaudatus, X. setariae/vulgare complex, M. xenoplax, and H. dihystera remained low.

Table 1.

Molecular identification of plant-parasitic nematode species in the rooting zone of hop cultivars in a Florida field soil.

10.21307_jofnem-2020-040-t001.jpg

Nematode population development in hop yard planting No. 1

Soil population densities of M. javanica did not differ significantly among the hop cultivars (Table 2); however, populations were large (> 200 nematodes per 200 mL of soil) in the ‘Chinook’, ‘Centennial’ (WA), ‘Centennial’ (OR), and ‘Cascade’ (OR) cultivars. Population densities of M. javanica remained low (< 50 nematodes per 200 mL soil) in the ‘CTZ’, ‘Magnum’, and ‘Triple Perle’ cultivars. Soil population densities of P. brachyurus did not differ among the hop cultivars; however, populations were largest in the ‘Magnum’ cultivar (69 nematodes per 200 mL of soil).

Table 2.

Effect of hop cultivar and planting material source on soil populations of plant-parasitic nematodes and root galling in December 2016 in Hop Yard Planting No. 1.

10.21307_jofnem-2020-040-t002.jpg

Root galling varied considerably among the hop cultivars in Hop Yard Planting No. 1. The most severe root galling was observed on ‘Chinook’, ‘Centennial’ (OR), ‘Fuggle’, and ‘Centennial’ (WA). The lowest amount of root galling was observed on ‘Magnum’, ‘Triple Perle’, ‘CTZ’, and ‘Nugget’.

Nematode population development in hop yard planting No. 2

In December 2017, M. javanica population densities in soil did not differ significantly among the hop cultivars; however, populations were largest in the ‘Cascade’ (FL), ‘Centennial’ (WA), ‘Centennial’ (FL), ‘Chinook’ (FL), and ‘Comet’ cultivars (Table 3). In July 2018, soil population densities of M. javanica were significantly larger in the ‘Centennial’ (WA) cultivar relative to that of the ‘Cascade’ (FL), ‘Chinook’ (FL), ‘Comet’, ‘Magnum’, ‘Nugget’, and ‘Triple Perle’ cultivars. In December 2018 and July 2019, M. javanica populations in soil did not differ significantly among the hop cultivars; however, population densities were consistently large in the ‘Centennial’ (WA), ‘Centennial’ (FL), and ‘Comet’ cultivars.

Table 3.

Effect of hop cultivar on soil populations of Meloidogyne javanica in Hop Yard Planting No. 2.

10.21307_jofnem-2020-040-t003.jpg

Soil population densities of P. brachyurus did not differ significantly among the hop cultivars throughout the first two years of plant establishment in Hop Yard Planting No. 2 (Table 4). Populations of P. brachyurus remained numerically low in soil planted with ‘Cascade’ (WA), ‘Cashmere’, ‘Centennial’ (WA), ‘Nugget’, and ‘Triple Perle’.

Table 4.

Effect of hop cultivar on soil populations of Pratylenchus brachyurus in Hop Yard Planting No. 2.

10.21307_jofnem-2020-040-t004.jpg

Nematode population development in hop yard planting No. 3

Soil population densities of M. javanica did not differ significantly among the hop cultivars (Table 5) but in July 2018 and December 2018 soil population densities were consistently large (> 100 M. javanica per 200 mL soil) in the ‘Canadian Red Vine’, ‘Mt. Ranier’, ‘Tahoma’, and ‘Willamette’ cultivars. Similarly, soil population densities of P. brachyurus did not differ among the hop cultivars throughout the first two years of plant establishment (Table 6) however, population densities were consistently large in the ‘Galena’ and ‘Mt. Rainier’ cultivars.

Table 5.

Effect of hop cultivar on soil populations of Meloidogyne javanica in Hop Yard Planting No. 3.

10.21307_jofnem-2020-040-t005.jpg
Table 6.

Effect of hop cultivar on soil populations of Pratylenchus brachyurus in Hop Yard Planting No. 3.

10.21307_jofnem-2020-040-t006.jpg

Hop cultivar susceptibility to M. javanica in the greenhouse experiment

Root galling differed significantly among the hop cultivars evaluated in the greenhouse experiment (Table 7). The most severe root galling was observed on ‘Comet’, ‘Cashmere’, ‘Canadian Red Vine’, and ‘Chinook’. The lowest amount of root galling was observed on ‘Galena’, ‘Willamette’, and ‘Cascade’. The number of M. javanica eggs extracted per plant also differed significantly among the various hop cultivars. The greatest number of eggs was obtained from the ‘Comet’, ‘Cashmere’, ‘Centennial’, ‘Cascade’, and ‘Chinook’ cultivars. The fewest number of eggs was obtained from the ‘Magnum’, ‘Willamette’, ‘Galena’, and ‘Canadian Red Vine’ cultivars.

Table 7.

Effect of hop cultivar on root galling and the abundance of Meloidogyne javanica eggs per plant in the greenhouse experiment.

10.21307_jofnem-2020-040-t007.jpg

Discussion

A diverse range of plant-parasitic nematodes inhabit Florida soils (Porazinska et al., 1999), likely as a result of favorable biotic and abiotic conditions provided by the sandy soils and warm sub-tropical climate characteristic of this growing region. As a result, plant-parasitic nematodes present a significant obstacle to most crops planted in Florida soils. In this study, plant-parasitic nematode species recovered from the root zone of hop planted in a Florida field soil included M. javanica, P. brachyurus, P. minor, B. longicaudatus, X. setariae/vulgare complex, M. xenoplax, and H. dihystera. With the exception of M. javanica (Brito et al., 2018), this is the first report of P. brachyurus, P. minor, B. longicaudatus, X. setariae/vulgare complex, M. xenoplax, and H. dihystera in the rooting zone soil of hop plants in Florida. For all nematodes, percent identity was greater than 97% for the first five accession numbers in each BLAST search; however, only a single amplicon was sequenced per nematode species, thus restricting the power of this study to identify mixed-species populations within the hop yard. Although these plant-parasitic nematode species were present in the root zone of hop plants, soil population densities remained relatively low for P. minor, B. longicaudatus, X. setariae/vulgare complex, M. xenoplax, and H. dihystera, suggesting that these particular species are unlikely parasites of major concern in the growing region. The recovery of these nematodes from soil in the rooting zone of the hop cultivars may have been an artifact of the previous natural grass cover in the field prior to establishing the hop yard, or the ryegrass groundcover located between the planting rows, as opposed to direct parasitism of the hop roots. Our findings are in agreement with other studies which have also reported lesion and root-knot nematode parasitizing hop plants in significant numbers in the Pacific Northwest of United States (Maggenti, 1962; Hafez et al., 1992), albeit by more temperate climate nematode species (P. neglectus, P. thornei, and M. hapla) than the nematode species observed in this study.

The abundance of particular plant-parasitic nematode species in the root zone differed considerably among the different hop cultivars planted. Root-knot nematodes are widespread in Florida (Garcia and Rich, 1985), and likely pose one of the most significant obstacles to successful production of hop in the growing region. All hop cultivars that were evaluated were hosts to M. javanica to some extent; however, root galling index ratings, egg production, and soil population densities of M. javanica were consistently greater on the ‘Canadian Red Vine’, ‘Centennial’, ‘Chinook’, and ‘Comet’ hop cultivars. Root-knot nematode population development was lowest on the ‘Galena’ and ‘Triple Perle’ cultivars. In China, differential host susceptibility of hop cultivars to M. incognita has been observed, where the hop cultivar ‘Comet’ also supported substantial root-knot nematode population development, whereas nematode population development was restricted on a locally bred cultivar (Zhang and Zheng-Li, 1989).

No consistent differences in lesion nematode (P. brachyurus) soil population densities were observed among the hop cultivars, and destructive sampling to quantify root populations was not performed in order to maintain root health in the newly established hop yard. This makes it difficult to draw definitive conclusions about the host status of P. brachyurus on cultivars used in this study. Conducting additional controlled cultivar screening greenhouse experiments, similar to the greenhouse experiment performed with M. javanica in this study, could help further delineate the host status of hop cultivars to this nematode. Selecting cultivars that are resistant or tolerant to nematode feeding will be vital to establishing a successful hop industry in Florida.

Plant-parasitic nematodes present an obstacle to any crop grown in Florida and finding suitable hop cultivars that can thrive in the sub-tropical climate as well as tolerate or resist nematode feeding will be vital to the establishment of a sustainable hop industry in the region. In a recently conducted field trial in Central Florida, the cultivars ‘Columbus’ and ‘Chinook’ showed the greatest bine lengths and yield, followed by ‘Neo1’ and ‘Amalia’ (Pearson et al., 2016). In addition to finding vigorous growing hop cultivars, developing effective pest management strategies will also be key to sustaining the hop industry. As a herbaceous perennial crop, opportunities for soilborne pest management are limited on hop. In this study, pre-plant fumigation only provided temporary nematode relief at the time of planting and did not prevent subsequent nematode population development later on in the growing season, as has been observed in other studies on perennial crops (Watson et al., 2017). Nematicides may be able to provide effective management options for use by growers (e.g. fluopyram), alongside other management tools, such as the development of tolerant/resistant hop cultivars through hop breeding programs and adopting management practices that promote soil health and disease suppressive soils.

Overall, our study provides the first report of the plant-parasitic nematode species recovered from soil in the rooting zone of hop in Florida and provides a basis for selecting hop cultivars that restrict plant-parasitic nematode population development in Florida field soils.

Acknowledgements

This research was funded in part through two subrecipient grants (‘Establishing a hop production system for Florida growers’ and ‘Development of viable production methods for hop production in Florida’) awarded by the United States Department of Agriculture through the Florida Department of Agriculture and Consumer Services (the Florida Specialty Crop Block Grant Program). The contents in this publication do not necessarily reflect the views or policies of the United States Department of Agriculture nor does mention of trade names, commercial productions, services or organization imply endorsement by the US Government. The authors thank David Moreira, Justin Carter, Hailey Regier, Bonnie Xie, Joyce Jones, Gail Bowman, Keri Druffel, Alan Behrens, Vinny Nguyen, Patrick Hooks, Jared Rogers, Angelica Castro, et al., and the Gulf Coast Research and Education Center farm crew for their assistance.

References


  1. Hope Growers of America. 2018. USA hops 2018 statistical report. Hop Grow. Am. , available at: https://www.usahops.org/img/blog_pdf/168.pdf.
  2. Bridge, J. and Page, S. L. J. 1980. Estimation of root-knot nematode infestation levels on roots using a rating chart. International Journal of Pest Management 26:296–298.
  3. Brito, J. A. , Subbotin, S. A. , Desaeger, J. , Achinelly, F. and Qiu, S. 2018. First report of the root-knot nematode (Meloidogyne javanica) infecting hops (Humulus lupulus) in Florida, USA. Journal of Nematology 50:1–2.
  4. Cobb, S. G. 1962. First record of hop cyst nematode (Heterodera humuli Filipjev, 1934) in the United States. Plant Disease Report 46:541.
  5. De Grisse, A. and Gillard, A. 1963. Morphology and biology of hop cyst eelworm (Heterodera humuli Filipjev 1934). Nematologica 9:41–48.
  6. Floyd, R. , Abebe, E. , Papert, A. and Blaxter, M. 2002. Molecular barcodes for soil nematode identification. Molecular Ecology 11:839–850.
  7. Foot, M. A. and Wood, F. H. 1982. Potato rot nematode, Ditylenchus destructor (Nematoda: Tylenchida), infecting hops in New Zealand. New Zealand Journal of Experimental Agriculture 10:443–446.
  8. Forge, T. A. and Kimpinski, J. 2007. “Nematodes”, in Carter, M. R. and Gregorich, E. G. (Eds), Soil sampling and methods of analysis 2nd ed, CRC Press, Boca Raton, FL, pp. 415–425.
  9. Garcia, M. and Rich, J. R. 1985. Root-knot nematodes in North-Central Florida soybean fields. Nematropica 15:43–48.
  10. Hafez, S. L. , Golden, A. M. , Rashid, R. and Handoo, Z. 1992. Plant-parasitic nematodes associated with crops in Idaho and Eastern Oregon. Nematropica 22:193–204.
  11. Holterman, M. , Van Der Wurff, A. , Van Den Elsen, S. , Van Megen, H. , Bongers, T. , Holovachov, O , Bakker, J. and Helder, J. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23:1792–1800.
  12. Hussey, R. S. and Barker, K. R. 1973. A comparison of methods of collecting inocula for Meloidogyne spp., including a new technique. Plant Disease Report 57:1025–1028.
  13. Maggenti, A. R. 1962. Hot water treatment of hop rhizomes for nematode control. California Agriculture October, 11–12.
  14. Mahaffee, W. , Pethybridge, S. and Gent, D. H. 2009. Compendium of hop diseases and pests American Phytopathological Society, St. Paul, MN.
  15. Pearson, B. J. , Smith, R. M. and Chen, J. 2016. Growth, strobile yield, and quality of four Humulus lupulus varieties cultivated in a protected open-sided greenhouse structure. HortScience 51:838–842.
  16. Pelerents, C. , Heungens, A. , Dejonckheere, W. and Barbez, D. 1983. Control of nematodes in hops. Mededelingen van de Faculteit Landbouwwetenschappen, Rijksuniversiteit Gent 48:75–77.
  17. Porazinska, D. L. , Duncan, L. W. , McSorley, R. and Graham, J. H. 1999. Nematode communities as indicators of status and processes of a soil ecosystem influenced by agricultural management practices. Applied Soil Ecology 13:69–86.
  18. Steiner, S. 1973. Steiner’s guide to American hops Hopsteiner, New York, NY.
  19. Turner, S. F. , Benedict, C. A. , Darby, H. , Hoagland, L. A. , Simonson, P. , Robert Sirrine, J. and Murphy, K. M. 2011. Challenges and opportunities for organic hop production in the United States. Agronomy Journal 103:1645–1654.
  20. USDA-NASS 2018. National hop report USDA National Agricultural Statistics Service, Washington, DC.
  21. Watson, T. T. , Nelson, L. M. , Neilsen, D. , Neilsen, G. H. and Forge, T. A. 2017. Soil amendments influence Pratylenchus penetrans populations, beneficial rhizosphere microorganisms, and growth of newly planted sweet cherry. Applied Soil Ecology 117–118:212–220.
  22. Zhang, H. and Zheng-Li, L. 1989. Histological studies on the root-knot nematodes nonresistsant and resistant root apex of Humulus lupulus L. Acta Botanica Sinica 31:576–580.
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REFERENCES

  1. Hope Growers of America. 2018. USA hops 2018 statistical report. Hop Grow. Am. , available at: https://www.usahops.org/img/blog_pdf/168.pdf.
  2. Bridge, J. and Page, S. L. J. 1980. Estimation of root-knot nematode infestation levels on roots using a rating chart. International Journal of Pest Management 26:296–298.
  3. Brito, J. A. , Subbotin, S. A. , Desaeger, J. , Achinelly, F. and Qiu, S. 2018. First report of the root-knot nematode (Meloidogyne javanica) infecting hops (Humulus lupulus) in Florida, USA. Journal of Nematology 50:1–2.
  4. Cobb, S. G. 1962. First record of hop cyst nematode (Heterodera humuli Filipjev, 1934) in the United States. Plant Disease Report 46:541.
  5. De Grisse, A. and Gillard, A. 1963. Morphology and biology of hop cyst eelworm (Heterodera humuli Filipjev 1934). Nematologica 9:41–48.
  6. Floyd, R. , Abebe, E. , Papert, A. and Blaxter, M. 2002. Molecular barcodes for soil nematode identification. Molecular Ecology 11:839–850.
  7. Foot, M. A. and Wood, F. H. 1982. Potato rot nematode, Ditylenchus destructor (Nematoda: Tylenchida), infecting hops in New Zealand. New Zealand Journal of Experimental Agriculture 10:443–446.
  8. Forge, T. A. and Kimpinski, J. 2007. “Nematodes”, in Carter, M. R. and Gregorich, E. G. (Eds), Soil sampling and methods of analysis 2nd ed, CRC Press, Boca Raton, FL, pp. 415–425.
  9. Garcia, M. and Rich, J. R. 1985. Root-knot nematodes in North-Central Florida soybean fields. Nematropica 15:43–48.
  10. Hafez, S. L. , Golden, A. M. , Rashid, R. and Handoo, Z. 1992. Plant-parasitic nematodes associated with crops in Idaho and Eastern Oregon. Nematropica 22:193–204.
  11. Holterman, M. , Van Der Wurff, A. , Van Den Elsen, S. , Van Megen, H. , Bongers, T. , Holovachov, O , Bakker, J. and Helder, J. 2006. Phylum-wide analysis of SSU rDNA reveals deep phylogenetic relationships among nematodes and accelerated evolution toward crown clades. Molecular Biology and Evolution 23:1792–1800.
  12. Hussey, R. S. and Barker, K. R. 1973. A comparison of methods of collecting inocula for Meloidogyne spp., including a new technique. Plant Disease Report 57:1025–1028.
  13. Maggenti, A. R. 1962. Hot water treatment of hop rhizomes for nematode control. California Agriculture October, 11–12.
  14. Mahaffee, W. , Pethybridge, S. and Gent, D. H. 2009. Compendium of hop diseases and pests American Phytopathological Society, St. Paul, MN.
  15. Pearson, B. J. , Smith, R. M. and Chen, J. 2016. Growth, strobile yield, and quality of four Humulus lupulus varieties cultivated in a protected open-sided greenhouse structure. HortScience 51:838–842.
  16. Pelerents, C. , Heungens, A. , Dejonckheere, W. and Barbez, D. 1983. Control of nematodes in hops. Mededelingen van de Faculteit Landbouwwetenschappen, Rijksuniversiteit Gent 48:75–77.
  17. Porazinska, D. L. , Duncan, L. W. , McSorley, R. and Graham, J. H. 1999. Nematode communities as indicators of status and processes of a soil ecosystem influenced by agricultural management practices. Applied Soil Ecology 13:69–86.
  18. Steiner, S. 1973. Steiner’s guide to American hops Hopsteiner, New York, NY.
  19. Turner, S. F. , Benedict, C. A. , Darby, H. , Hoagland, L. A. , Simonson, P. , Robert Sirrine, J. and Murphy, K. M. 2011. Challenges and opportunities for organic hop production in the United States. Agronomy Journal 103:1645–1654.
  20. USDA-NASS 2018. National hop report USDA National Agricultural Statistics Service, Washington, DC.
  21. Watson, T. T. , Nelson, L. M. , Neilsen, D. , Neilsen, G. H. and Forge, T. A. 2017. Soil amendments influence Pratylenchus penetrans populations, beneficial rhizosphere microorganisms, and growth of newly planted sweet cherry. Applied Soil Ecology 117–118:212–220.
  22. Zhang, H. and Zheng-Li, L. 1989. Histological studies on the root-knot nematodes nonresistsant and resistant root apex of Humulus lupulus L. Acta Botanica Sinica 31:576–580.

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