1http://dx.doi.org/10.4322/nematoda.00417 Nematoda, e042017

Nematoda. ISSN 2358-436X. This work is licensed under a Creative Commons Attribution International License (https://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

Potential of Crotalaria juncea as cover crop for cyst nematode suppression under Central Europe conditionsSomayyeh Sedaghatjooa, Hans-Peter Söchtinga and Andreas Westphalb*

a Julius Kühn-Institut, Institute for Plant Protection in Field Crops and Grassland, Braunschweig, Germanyb Department of Nematology, University of California, Riverside, Kearney Agricultural Research and Extension Center, Parlier (CA) United States

*andreas.westphal@ucr.edu

HIGHLIGHTS• Crotalariajuncea was resistant to Heteroderaavenae,H.filipjevi,H.schachtii,Globoderapallida and G.rostochiensis.• C.juncea grew vigorously when planted in May and June in northern Germany.• Cropping of C.juncea kept infectivity rates of soil H.schachtii egg populations low.

ABSTRACT: Interest in the use of cover crops for soil health improvement and soil amelioration with organic matter is increasing. In addition to more general benefits, cover crops have also long been implemented for biological nematode suppression. For example for more than three decades, Heteroderaschachtii has been managed by cultivating nematode-resistant brassica cover crops in Central Europe and more recently in North America. Production systems may require additional cover crop species when brassica cover crops are undesirable because of the potential increase of other soil-borne diseases, e.g., club root, that could negatively impact brassica cash crops. Alternative plant species fit for nematode suppression are needed. The tropical legume Crotalariajuncea has been successfully implemented for the suppression of Meloidogyne spp. in climates most typical to its growth requirements. This plant combines the benefits of nitrogen fixation with nematode antagonistic properties. Crotalariajuncea poses no consumption risk for ruminant animals in contrast to other species of this genus. The current study determined the C.juncea host status to five cyst nematodes, and examined the vegetation period requirements of the crop of exemplary two commercial cultivars. Both cultivars were resistant to the nematode species tested, and grew most vigorously when planted in May or June when soil temperatures were conducive for germination. This widely known nematode-suppressing plant can be grown in Central Europe. Further field studies will need to address the implementation of this plant species.

Keywords: cultural practice, Globodera spp., Heterodera spp., Heteroderaschachtii.

Cite as Sedaghatjoo S, Söchting H-P, Westphal A. Potential of Crotalariajuncea as cover crop for cyst nematode suppression under Central Europe conditions. Nematoda. 2017;4:e042017. http://dx.doi.org/10.4322/nematoda.00417

Received: Mar. 24, 2017 Accepted: May 31, 2017

ORIGINAL ARTICLE

INTRODUCTIONPlant-parasitic nematodes resemble a long-lasting production challenge for almost all crops. In many

areas of the world, nematode infections are suppressed by chemical control options[1]. In Europe for decades, such strategies are under critical reconsideration in light of environmental, consumer and applicator health issues. For example in Germany, only Fosthiazate as nematicidal compound for use in potato (Solanum tuberosum L.) is currently registered[2]. Implementation of host plant resistance and tolerance of cash crops has been successful to reduce the yield-reducing effects of nematode pests[3]. Such strategies require a long breeding process to develop high-yielding cultivars and are often threatened by the presence of new nematode pathotypes that can overcome the resistance[4].

2http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

Use of suppressive soil concepts along with directed implementation of biological control carries great promise but needs to be developed into management recommendations before comprehensive usefulness can be secured[5]. Sustainable production systems require integrated management options that predictably reduce the negative effects of nematode infestations[6].

Application of organic amendments and cropping of catch crops as cover or companion crops has a long tradition in integrated nematode management[7, 8]. One success story of implementing resistant cover crops as integral component of a production system is the use of nematode-resistant brassica hosts as catch crops to reduce population densities of Heterodera schachtii Schm. below biological and economic threshold levels in sugar beet (Beta vulgaris L.) production in Germany and the U.S[9,

10]. In this system, nematode-resistant brassica cover crops are grown prior to sugar beet that then can be produced every three years. Heterodera schachtii negatively impacts yields of sugar beet and brassica vegetable crops[11, 12]. Protected by its cyst, this parasite survives for several years in the field, and various field parameters impact how natural decline and spontaneous hatch reduce the inoculum potential relative to threshold levels[13]. In addition to nematode suppression, cover cropping provides benefits in other ways for yield improvements in sugar beet[14, 15].

In central Europe, cropping of sugar beet and oilseed rape (Brassica napus L.), another host of H. schachtii, in the same rotation has long been discouraged because the feared build-up of nematode populations but recently, it has been more frequently practiced[16, 17]. No detrimental nematode reproduction under the production rapeseed was detected, and heavy reproduction under the inevitable volunteer rape seed after harvest of the cash crop can be averted by volunteer destruction after 250 degree days[18, 19]. A crop rotation scheme using sugar beet, brassica cover crops, and rapeseed with high frequency, increases the potential risk of build-up of Plasmodiophora brassicae Woronin on the brassica, which is an increasing risk factor for rape seed production[20]. Clearly alternative cover crops with H. schachtii-reducing potential are urgently needed.

Worldwide, various plant species are used as cover crops. Crotalaria juncea L. is of particular interest because of its poor-host status to many plant-parasitic nematodes[21]. Other benefits of this plant are its nematode antagonistic properties via secondary metabolites and its nitrogen fixation capacities being a tropical legume[21]. The plant is well adapted to sub-tropical and tropical climates where it is reported effective against the most important nematodes, namely Meloidogyne spp[21]. This plant has been successfully used as far North as Maryland, US (39°2’ N) and the Po Valley of Italy (45°0’ N)[22, 23] but very little is known if it could be grown in Central Europe and whether it would provide its nematode suppressing capacities against cyst nematodes.

We hypothesized that C. juncea was resistant and suppressive against key nematode pests of Central Europe, and that it could be grown successfully in northern Germany. The objectives of the current study were to (a) determine the host status of C. juncea to Globodera pallida (Stone) Behrens, G. rostochiensis (Woll.) Behrens, Heterodera avenae Wollenweber, H. filipjevi (Madzhidov) Stelter, and H. schachtii; (b) determine the adaptability in germination, nitrogen fertilizer requirements, and growth pattern to Central Europe, and (c) measure its nematode suppressing capacities under field conditions in microplots.

MATERIAL AND METHODSTwo cultivars of C. juncea were used in the study, ‘large cap Tillage Sunn’ (Cover Crop Solutions,

Lititz, PA, US) and ‘Tropic Sun’ (Molokai Seeds, Kaunakakai, Hawaii, US). Seeds of each cultivar were inoculated with Rhizobium leguminosarum Frank biovar viceae (N-Dure, INTX Microbials LLC, Kentland, IN, US) at 0.6 g material per 100 g of seed before introducing into the various experimental contexts unless otherwise noted.

Resistance determination to Globodera pallida (pathotype Pa 3) and G. rostochiensis (Ro1, Ro 3)

This test followed the standard protocol as outlined in the EU Directive[24]. Eighteen seeds of each of the Crotalaria ‘Tropic Sun’ and ‘Tillage Sunn’ were planted per 1.4-L pot into loess soil amended with slow release fertilizer (1.5 g slow-release fertilizer Osmocote/kg of loess soil) while being amended with cysts containing eggs from pure inocula raised on potato ‘Desiree’ in the year prior (Ro1: 31 cysts containing 14,942 eggs; Ro3: 36 cysts containing 14,904 eggs; Pa3: 34 cysts containing 15,215 eggs). After emergence, plants were reduced to fifteen per pot. Controls were pots planted with a

3http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

single seed piece of the susceptible potato ‘Desiree’ and a fallow treatment. Four replicate pots were arranged on greenhouse benches initially at 18 °C and later as ambient temperature allowed below 25 °C, and incubated from March 11 until June 23. At harvest, plant tops were severed and discarded. The entire pot content was used for nematode extraction. Cysts were washed off the root system with a jet stream of water while this was placed in a large-mesh sieve nested on top of a 5-L bucket with a 250-μm aperture sieve bottom. The separated cysts were transferred onto a filter paper to then be separated by the ethanol method[25], counted, and broken open in a custom-made cyst crusher[26] for egg counts. Egg numbers were compared to those under a single plant/pot of the susceptible potato ‘Desiree’ standard, and a tentative resistance class assigned as prescribed by the EU Directive.

Resistance determination to Heterodera avenae and H. filipjeviThis test followed the standard protocol implemented for resistance determination of small grains

towards cereal cyst nematodes (Westphal, unpublished). Briefly, twelve PVC folding boxes holding 100 mL soil (20 × 40 × 120 mm) filled with subsoil loess amended with nutrient solution were planted to the seeds. Deviating from that protocol, seeds of crotalaria were germinated in a greenhouse at 20 °C because laboratory tests had documented the need of 20 °C for high germination rates. Only after successful germination and plant establishment (two weeks after seeding), ten to twelve plants per treatment (depending on the experiment) were transferred into an environmental growth chamber for incubation at the preset published temperature regiment. Plants were inoculated with 1000 second-stage juveniles (J2) of either H. avenae or H. filipjevi. A subset of plants was harvested ten days after inoculation for root staining for nematode penetration following the protocol by Byrd et al.[27].

Resistance determination to Heterodera schachtiiIn a first experiment (experiment 1), the test plants were planted into the folding boxes as described

for the cereal cyst nematode tests to allow for recovery of the young plants, and root staining according to Byrd et al.[27]. All following tests for resistance against H. schachtii (experiment 2-4) followed the standard protocol[28]. Briefly, eighteen seeds of each of the crotalaria ‘Tropic Sun’ and ‘Tropic Sun’ were planted into 1.4-L pots filled with loess material. This material had been sieved, and mixed with slow-release fertilizer (Osmocote, containing macro-nutrients at 11:11:13:2) and cysts with contained eggs of H. schachtii from greenhouse culture before potting. After emergence fifteen plants per pot were maintained. Four replicate pots were prepared for each of the three independent runs of this experiment with the following initial population densities: 1,174, 1,174, or 1,695 eggs/100 g of soil, respectively. After a temperature sum of approximately 350 DD, tops were severed, and the test soil incubated for two weeks to allow for completion of the one nematode generation expected in such growing period. Cysts were extracted form composites of two pots, and the eggs counted and classified in healthy and diseased after crushing the cysts[26]. Healthy eggs were counted and reported on per 100 g of soil basis.

Suppressing capacity of planting Crotalaria juncea at different times on Heterodera schachtii

In a microplot experiment in Braunschweig, Germany (52°16’ N, 10°34’ E, 71 m a.s.l.), bottomless pots of 32 cm diameter were inserted 26 cm deep in soil spaced at 70 by 75 centre-to-centre distance. Pots were bottomless to allow for free water infiltration. Soil at each plot location was removed to 60 cm depth to remove native nematode populations. The depth of 30-60 cm below the soil surface was filled with steamed sandy loam soil (70.4% sand, 23.8% silt, 5.8% clay, 1% O.M., pH 6.2). The layer of 0-30 cm depth was made up of similar soil but infested with 2,695 eggs in 112 cysts/100 g of soil by amendment with infested sandy loam soil before placing into the plots. For the deep and the shallow layer, soil was mixed per each experimental block in a commercial soil-mixing bin. Treatments were assigned in a randomized complete block design with six replications. Plantings of 28 seeds per plot were executed on April 8, May 14, June 6, July 8, and attempted several times unsuccessfully in August, all for a mutual harvest date in October when growth slowed because of quickly declining ambient temperatures. Controls were a fallow treatment and one with H. schachtii-resistant mustard Sinapis alba L. ‘Serval’ planted on August 28 at the same plant density to simulate the current commercial

4http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

practice of cover cropping. Intermittendly during the growing season, plant emergence was observed, and plant establishment was counted when plant height was measured.

On October 2, all plants of the experiment were harvested. Plant tops were severed at ground level and weighed. Soil samples were taken (six cores per plot, 30 cm deep) and composited. Plant tops were shredded with a garden chopper (natura 2800L, Gloria GmbH, Witten, Germany) to approximately 3-5 cm long pieces, spread on the plot surface, and incorporated to a 20 cm depth by digging with a fork. After incorporation, each plot received 3 L of water to seal the soil surface, and to encourage soil-plant material contact. Soil samples of October 2 were submitted to a bioassay for determination for infective J2 of H. schachtii following procedures of Meinecke et al.[29]. Three subsamples per plot were processed by this radish bioassay following the published protocol[29]. On October 27 after 25 days of incubation, soil in each of the microplots was dug with the digging fork to 20 cm depth to promote tissue break down. On November 7, twelve soil cores were taken from each plot to provide material for another soil bioassay for infective H. schachtii by the bioassay[29] and for cyst extraction followed by egg counting[26].

Additional planting date examinationsSmall plots were arranged in split plot design in Braunschweig, Germany, in an area of sandy loam

soil (84% sand, 12.1% silt, 3.9% clay, 1.6% O.M., pH 6.5) that had been pre-season pasteurized. The main factor was the planting date (April 8, May 8, June 10, July 8, and August 7). Subplot factor was the plant cultivar. Each subplot was 2 m2, and there were four replicate plots for each planting date with the two subplots each. Before the first planting in spring, fertilizer (35 kg/ha P) was evenly spread on the experimental area. In each plot, 300 seeds per m2 were planted to six parallel 15-cm distance rows at 2.5 cm depth. At harvest on September 23, plants were excised at ground level from a 0.25-m2 area exhibiting typical growth. Plants were counted and dry weight was determined.

Nitrogen fertility requirements in microplotsA total of sixteen microplots of 1 m2 filled with sandy loam soil (72.9% sand, 20.5% silt, 6.6% clay,

1% O.M., pH 7.3) were used to examine the potential benefit of administering nitrogen fertilizer at planting of the cover crop. Right after small grain harvest, plots were lightly tilled, and mineral PK fertilizer (26.2 kg P/ha, 100 kg K/ha, and 26.5 kg Mg/ha) was administered uniformly. Nitrogen fertilizer treatments and cultivars were arranged in a split plot design: mineral nitrogen was supplied as calcium ammonium nitrate at 60 kg N/ha or no nitrogen fertilizing as the main factor and crotalaria cultivars ‘Tillage Sunn’ and ‘Tropic Sun’ as the subplot factor. Seeds of the crotalaria cultivars at 300 seeds/m2 were planted in four parallel rows of 10-cm spacing at a depth of 2.5 cm. After emergence, plant stands were repeatedly counted. Plantings were done on August 7, and the entire experiment repeated on August 14. On October 25, plants of the first planting date were excavated, plant tops taken for dry weights, and after counting, roots were washed and examined for rhizobium nodulation. On November 1, plants of the second planting date were processed similarly as those of the first planting date. Subsamples of each of the dried plant tops were processed for carbon and nitrogen content.

Statistical analysisExperimental data of the resistance determination were summarized as customary in the routine

testing program (Westphal, unpublished). Non-parametric analysis was conducted in Minitab 17 (State College, PA). For Globodera spp., median cyst and egg numbers were compared to the numbers under ‘Desiree’, and the relative susceptibility calculated according to Niere[30]. In Heterodera avenae and H. filipjevi, counts of penetrated J2, females, cysts and eggs were reported. For H. schachtii evaluations, medians were compared and reproductive values calculated and compared to the resistant standards used in the official resistance determination assay. Plant growth data of the field plots were submitted to analysis of variance (ANOVA) using Proc Mixed in SAS (SAS Institute, Cary, NC). Differences were tested at P = 0.05. Nematode counts were analysed after proper log-transformation [x = log10(xoriginal + 1)] if necessary to improve the homogeneity of error variances. The percentage of infectivity of J2/eggs found at harvest was calculated, and analysed by ANOVA.

5http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

RESULTS AND DISCUSSIONThe two cultivars of C. juncea ‘Tillage Sunn’ and ‘Tropic Sun’ were non-hosts or poor hosts of G. pallida,

G. rostochiensis, H. avenae, H. filipjevi and H. schachtii. Both cultivars responded similarly to nematode infestations. Previously, few studies had been conducted on the interaction of crotalaria and cyst nematodes, probably because legumes are generally viewed as poor hosts for cyst nematodes[31]. Only in H. glycines, juvenile penetration was measured but reproduction was severely reduced compared to fallow[32]. The findings reported here are valuable additions to the currently known susceptibility range of C. juncea[21]. For implementing a potential cover crop, confirmation of host status to target nematodes is crucial as nematodes may infect plants that are outside their presumed host range, and such could nullify any cover crop benefit.

For all pathotypes of Globodera spp. tested, lower cyst and egg numbers were found under the C. juncea cultivars than under potato ‘Desiree’, and numbers declined to the level of the fallow treatment (Table 1). In the official testing protocol for Globodera spp., the classes range from 9 (highest level of resistance) to 1 (fully susceptible)[30]. If this test was conducted twice and the class ranges for Globodera spp. were used to chararterize C. juncea, ‘Tillage Sunn’ and ‘Tropic Sun’ would be assigned to class 9. This resistance is especially important because management options for these quarantine pests are limited. In potato, the use of resistant cultivars is central for any management[24]. Crop sequences with non-hosts reduce numbers but the long survival capacity of viable cyst content makes rotation times disproportionately long[33]. Use of Solanum sisymbriifolium Lam has been proposed[34] but is limited by undesirable agronomic and horticultural characteristics of this spiny and potentially weed seed-producing plant. Use of brassica cover crops showed some promise but is still in experimental phases[35]. Brassica crops are limited by overabundance of brassica species within intensive crop sequences. The high value of potato probably allows for cropping C. juncea at the favorable planting dates during May, June or July.

In tests with H. avenae and H. filipjevi, cereals were penetrated by each of the nematode species that developed into healthy females, cysts and eggs only under the susceptible standard ‘Hanka’. No nematode reproduction was detected under the resistant cereal and both of the C. juncea cultivars (Table 2). Such lack of reproduction may allow for the successful incorporation of C. juncea into production systems as has been demonstrated in wheat producing systems for fertility benefits of southern US systems[36]. For northern Europe conditions, inclusion of C. juncea as a novel cover crop would be a true enrichment of production systems by increasing crop diversity for managing pests and diseases[37]. This chararacteristic allows for short growing periods with copious amounts of biomass, but also for weed suppression[38]. The fast juvenile development distinguishes C. juncea from other legumes that typically develop slowly. This may allow for cultivating after cereal harvest in late July and fall planting of a new crop in late September/early October. When cropped this way, its vigorous growth can exert some weed suppression[38, 39, 40, 41].

Table 1. Median numbers of cyst and egg population densities/100 g of soil of Globodera pallida (Pa3; Pi = 1,087 eggs/100 g of soil) and Globodera rostochiensis (Ro1, Ro3; Pi = 1,067; 1,065 eggs/100 g of soil, respectively) on potato ‘Desiree’, Crotalaria juncea ‘Tillage Sunn’ and ‘Tropic Sun’, and fallow in a greenhouse experiment conducted according to standard protocols*. Cysts and eggs were extracted from the entire pot volume, and numbers converted to counts per 100 g of soil.

Globodera pallida Pa3 Globodera rostochiensis Ro1 Globodera rostochiensis Ro3

Cysts Eggs Susc. [%]† Cysts Eggs Susc. [%]† Cysts Eggs Susc. [%]†

‘Desiree’ 416 a 118,125 a 100 376 a 118,821 a 100.0 290 a 68,464 a 100.0

‘Tillage sunn’

2 b 903 b 0.8 2 b 700 b 0.6 3 b 654 b 1.0

‘Tropic sun’

2 b 760 b 0.7 2 b 639 b 0.6 3 b 602 b 0.9

Fallow 2 b 846 b 0.8 2 b 446 b 0.4 3 b 671 b 0.9

P <0.01 <0.01 <0.01 <0.01 <0.01 <0.01

* Within a column, medians followed by the same letter were not significantly different at P = 0.05; †Relative susceptibility was calculated as egg numbers per those under the susceptible standard ‘Desiree’.

6http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

Parameters collected for plant response to H. schachtii included early root penetration and reproductive values during one nematode generation. Both crotalaria cultivars had much lower penetration by J2 then the two oilseed radishes (Table 3). Reproductive values were around the value 1. When resistance grades were assigned following the standard protocol[28], tentatively the grade 2 was assigned to the crotalaria (Table 3). Only ‘Resoluti’ received the highest resistance grade 1 in these assays (Table 3). In H. schachtii the success of using brassica cover crops is based on the trap crop concept in that initial attraction and penetration by the host plant is followed by prevention of reproduction because of the resistant phenotype of selected plant cultivars[9]. Thus minor penetration of the C. juncea roots by H. schachtii could be favorable by attracting the J2 Low nematode reproduction on C. juncea similar to that on the known resistant cover crop cultivars of Raphanus sativus L. were found but if this initial penetration is coupled with reproduction suppression needs further investigation.

Under microplot conditions, suppression of H. schachtii by C. juncea was similar to the resistant mustard and the fallow control, and intermittently reduced infectivity of the soil-inhabiting egg populations (Table 4). At harvest, final cyst and egg population densities indicated some trends but only few were significantly different at P = 0.05 (Table 4). Two bioassays were conducted: one at harvest (October) and a second test one month after incorporation of crotalaria tissues into the soil (November). In the bioassay with soil of the October sampling, a significantly higher percentage of J2/soil populations of the eggs were detected in the radish bioassay in the oilseed radish treatment than in the

Table 2. Root penetration and reproduction of Heterodera avenae, and Heterodera filipjevi on Crotalaria juncea ‘Tillage Sunn’ and ‘Tropic Sun’ in greenhouse experiments conducted following standard protocols.

Heterodera avenae Heterodera filipjevi

Penetration* Females† Cysts† Eggs† Penetration* Females† Cysts† Eggs†

‘Hanka’ 2.9 14.8 50.0 14,134 14.8 5.8 25.8 7,001

‘Isotta’ 4.5 0.0 0.0 -- 10.7 0.0 0.0 --

‘Tillage sunn’

0.1 0.0 0.0 -- 0.0 0.0 0.0 --

‘Tropic sun’

0.0 0.0 0.0 -- 0.0 0.0 0.0 --

*Second-stage juveniles that had penetrated the roots were counted after staining with acid fuchsin solution; †Nematode numbers counted per folding box (=100 mL of loess substrate): females on the root ball surface, cysts and eggs after the corresponding extraction.

Table 3. Root penetration and reproductive rate (R) of Heterodera schachtii on Crotalaria juncea ‘Tillage Sunn’ and ‘Tropic Sun’ compared to the official testing standards in greenhouse experiments based on standard protocols.

Experiment 1 Experiment 2 Experiment 3 Experiment 4 Summary

Penetration* Eggs† R‡ Eggs† R mean Eggs† R‡ R mean Grade§

‘Siletina’ 25.8 a 51,500 30.4 44,438 44.3 75,750 64.5 44.3 9

‘Adagio’ -- 903 0.5 588 0.8 1,612 1.4 0.8 2

‘Diabolo’ -- 1,441 0.9 1,393 1.6 3,140 2.7 1.6 2

‘Radical’ 35.3 a 1,103 0.7 1,126 1.0 1,663 1.4 1.0 2

‘Resoluti’ -- 639 0.4 431 0.5 784 0.7 0.5 1

‘Tillage sunn’

4.3 b 1,369 0.8 1,513 1.2 1,856 1.6 1.2 2

‘Tropic sun’

5.2 b 1,020 0.6 1,144 0.9 1,268 1.1 0.9 2

Fallow -- 1,351 0.8 1,110 1.0 1,363 1.2 1.0 -

*Medians with the same letter were not significantly different at P = 0.05; †Number of H. schachtii eggs/100 g of soil after one nematode generation; ‡The original R-value was calculated as R = Pf/Pi, where Pf = the number of eggs counted after one nematode generation, and Pi = the number of eggs at planting. The standards of the closest related experiment were used for comparisons with the test cultivars; § A R-value relative to the four resistant oilseed radish standards (Adgio, Diabolo, Radical and Resoluti) was calculated using the standard protocol; this relative R-values was used to assign a resistance grade between “1” most resistant and “9” most susceptible following standard procedures (Müller and Rumpenhorst[28]).

7http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

other four main treatments (Table 4). No differences in this parameter were observed in the soil of the November sampling (Table 4). A major proportion of the beneficial nematode suppression by C. juncea is attributed to nematode-resistance. Tissues of C. juncea contain alkaloids[42]. Content of pyrrolizidine alkaloids especially in ‘Tropic Sun’ was low[43], and it remains to be clarified how important these are in the nematode suppression. When we incorporated plant tops in our microplot studies after cover crop harvest we did not see additional nematode suppression (Table 4).

While the key components of nematode suppression are generally affected by these direct plant characteristics of host status and content of nematicidal compounds, microbial shifts could be important as well. For example, soil cropped with crotalaria experienced a stimulation of nematode-trapping fungi[44]. At the same time, Crotalaria spp. do not only have a close association with rhizobia but also mycorrhizal fungi[45] illustrating the far-reaching effects of growing this cover crop. The initial infection combined with lack of reproduction was found for Meloidogyne incognita (Kofoid & White) Chitwood and Rotylenchulus reniformis Linford & Oliveira[45, 46], and it may be important for the active trap crop character of C. juncea.

Crotalaria spp. is frequently used as cover crop throughout the western world including temperate regions[22] but it is also an important cash crop in India[47]. There, it grows well and produces seeds. Even at generally favorable growth conditions of South Texas, planting date had a huge impact on productivity[48]. The productivity is best described by a cumulative degree day model that predicts biomass production of C. juncea[49]. Thus it was not surprising that only selected planting dates provided the largest biomass in our studies. Both cultivars grew most vigorously when planted in early summer, and established only poor plant populations when planted in April or August. Crotalaria juncea prefers temperatures of 20 °C for proper germination (Westphal, unpublished). Under controlled conditions, the C. juncea cultivars germinated at >90% only at 20 °C whereas germination was <80% at 17.5 °C (Westphal, unpublished) indicating the high temperature requirements of this tropical legume.

In the outcome of this project, implementing crotalaria in northern Europe is an alternative to the use of known cover crops. One obvious limitation for the introduction of new plant species into a given area is the risk of weediness. For example in Nigeria, Crotalaria spp. are considered weeds but these plants are yet promoted for their nematode-trapping capacity[50]. The risk for weediness in central Europe is not given because C. juncea does not set viable seeds North of the northern latitude of 28° (NRCS 1999 cited in Stallings[36]). It withstands a number of herbicides[41], and thus could be used as uniform cover. In some studies, C. juncea provided weed suppression and reduced different Meloidogyne spp. while in other studies the leguminous cover crop cowpea [Vigna unguiculata (L.) Walp.] ‘Iron Clay’ was only resistant to one species[51].

In general, northern Europe (52° 16’ N) is out of the range of preferred cultivation of C. juncea[52] but it is grown successfully in the temperate climate of Maryland, US and the Po Valley of Italy[22, 23]. In addition to less than optimal soil temperatures in northern climates, damage from game and slugs not encountered in traditional areas may impact planting success. For example, deer will browse plants, turkey and quail will use C. juncea for shelter and food[53]. On average, both cultivars established the highest plant population densities when planted in June and July (Figure 1a). May plantings had the most biomass (Figure 1b). Although there was some differential response for the May and the June

Table 4. Population densities of Heterodera schachtii/100 g of soil and percentage of juveniles from eggs that penetrated bioassay radish roots under fallow, oilseed radish ‘Resoluti’, C. juncea ‘Tillage Sunn’ and ‘Tropic Sun’ treatments in microplots of initial population densities of 28 ± 1 cysts/100 g of soil containing 2,695 ± 297 eggs*.

Nematodes/100 g of soil Bioassay J2†

Cysts Eggs October November

Fallow 26 ± 1 535 ± 113 0.6 ± 0.1b 0.5 ± 0.4

Oil radish ‘Resoluti’ 24 ± 2 351 ± 54 2.4 ± 0.1a 0.2 ± 0.0

‘Tillage Sunn’ 24 ± 1 424 ± 45 0.3 ± 0.1 0.5 ± 0.1

‘Tropic Sun’ 26 ± 1 455 ± 35 0.7 ± 0.4 0.2 ± 0.1

Pculitvar

0.4054 0.1100 <0.0289 0.1754

*Means and standard errors of the means were shown. For C. juncea ‘Tillage sunn’ and ‘Tropic sun’ averages of plots from three planting dates were used in the analysis; †Number detected in radish bioassay roots as percentage of those that were present in 100 g of soil. In the October sampling a significant difference between fallow and oilseed radish could be ascertained, and were indexed with a different letter after the median, other trends could not be statistical estimated.

8http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

plantings, overall numbers were similar for both cultivars (Figure 1b). Plant population densities were highest in June and July (Figure 2a). Plantings in April and August mostly failed, probably because of inclement weather conditions coupled with a partially unexplained crop failure of dislodging tops (Figure 3). Biomass production was higher in earlier planting date treatments of May and June that had the highest amounts of biomass (Figure 2b).

Figure 1. Planting date study in non-infested, preseason steamed small plots with Crotalaria juncea ‘Tillage Sunn’ (solid black) and ‘Tropic Sun’ (hatch-marked). Plant establishment and growth was evaluated: (a) plant population densities, and (b) top biomass. Significant difference between the cultivars is indicated by “*”. Bars with the same letter within the panel were not significantly different at P = 0.05. Where appropriate numbers were analyzed separately for the two cultivars.

Figure 2. Planting date study in Heterodera schachtii-infested soil in microplots. Plant establishment and growth was evaluated: (a) plant population densities, and (b) top biomass. Cultivars responded similarly so values were combined. Bars with the same letter within the panel were not significantly different at P = 0.05.

Figure 3. Daily air and soil temperature, precipitation (incidence and daily accumulation) and planting dates of Crotalaria juncea in Braunschweig, Germany.

9http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

In arable soil, the amount of rhizobium nodulation was limited. Within the genus Crotalaria, C. juncea was a fast growing and high N-binding species[54]. No detailed evaluations of the test soil concerning N status were conducted. We speculate that high levels of this plant nutrient and fairly late seeding (associated with declining soil temperatures) may have contributed to the lack of nodulation. Counteracting of such environmental stresses to nodulation is well documented[55]. In the fertility tests, cultivars responded similarly to the nitrogen fertilizer amendments. On average of the two experiments and for the two cultivars, the percent nitrogen per dry matter of the tops was increased from 3.61 ± 0.10 to 4.31 ± 0.12 (P = 0.01), and the nitrogen content in the roots from 1.84 ± 0.05 to 2.15 ± 0.06 (P = 0.0239; data not shown). Other yield parameters were similar. No increased N contribution via the plant material was found (data not shown).

The experiment in non-infested ground had fairly suppressed growth at the planting times examined. Penetrometer readings throughout the experimental area revealed the presence of severe soil compaction below 20-25 cm depth. Such soil problems would probably be alleviated in sugar beet fields where a deep rooting horizon is mandatory for the cash crop. Similarly heavy August rains probably encumbered the August planting that failed in one of the contexts.

CONCLUSIONSTwo cultivars of C. juncea were non-hosts or poor hosts for several cyst nematodes. The cultivars

grew vigorously at supportive planting dates. The combined potential benefits of this tropical legume of nitrogen fixation with nematode suppression appear to make it a valuable enrichment of intensive production systems in northern Europe.

ACKNOWLEDGEMENTSPartial support by Cover Crop Solutions made this project possible. The authors thank C. Blümcke,

T. Blöthe-Hartmann, S. Deyhle, K. Hauffe, Salewski, P. Schwabe, S. Therre, and W. Baar’s field crew, J. Ripken’s greenhouse group and the Weed Science group at JKI for technical assistance. The nitrogen content determination by M. Kücke’s group is greatly appreciated. Seed provision by Molokai Seeds is appreciated. The experimental part of this study was conducted during the employment of the corresponding author Andreas Westphal with the Julius Kühn-Institut, Institute for Plant Protection in Field Crops and Grassland.

REFERENCES1. Hague NGM, Gowen SR. 1987. Chemical control of nematodes. In: Brown RH, Kerry BR (eds.). Principles and practice

of nematode control in crops. Academic Press, Sydney, p. 131-178.

2. Bundesamt für Verbraucherschutz und Lebensmittelsicherheit – BVL. 2016. Mittel gegen Nematoden. In: Bundesamt für Verbraucherschutz und Lebensmittelsicherheit – BVL (ed.). Pflanzenschutzmittel – Verzeichnis 2016 [on line]. BVL Braunschweig, Germany, p. 186-186. Accessed on 18 June, 2016. Available from: http://www.bvl.bund.de/SharedDocs/Downloads/04_Pflanzenschutzmittel/psm_verz_1.pdf?__blob=publicationFile

3. Müller J. 1998. Resistance and tolerance to beet cyst nematodes (Heterodera schachtii) in sugar beet cultivars. Zuckerindustrie, 123: 688-693.

4. Müller J, Klinke A. 1996. Selection of virulent Heterodera schachtii populations and their use in characterizing resistacne genes in the genus beta. Mitteilungen aus der Biologischen Bundesanstalt für Land- und Forstwirtschaft, 317: 102-116.

5. Stirling GR.2014. Biological control of plant-parasitic nematodes: soil ecosystem management in sustainable agriculture. 2nd ed. CAB International, Wallingford, 510 pp.

6. Westphal A. 2011. Sustainable approaches to the management of plant-parasitic nematodes and disease complexes. Journal of Nematology, 43: 122-124. PMid:22791923.

7. Oka Y. 2010. Mechanisms of nematode suppression by organic soil amendments: a review. Applied Soil Ecology, 44: 101-115. http://dx.doi.org/10.1016/j.apsoil.2009.11.003.

8. McSorley R. 2011. Overview of organic amendments for management of plant-parasitic nematodes, with case studies from Florida. Journal of Nematology, 43: 69-81. PMid:22791915.

9. Müller J, Steudel W. 1983. Der Einfluss der Kulturdauer verschiendener Zwischenfrüchte auf die Abundanzdynamik von Heterodera schachtii Schmidt. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 35: 103-108.

10. Koch DW, Gray FA. 1997. Nematode-resistant oil radish for control of Heterodera schachtii. I. sugarbeet-barley rotations. Journal of Sugar Beet Research, 234: 31-43. http://dx.doi.org/10.5274/jsbr.34.1.31.

11. Müller J. 1999. The economic importance of Heterodera schachtii in Europe. Helminthologia, 36: 205-213.

10http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

12. Abawi GS, Mai MF. 1980. Effect of initial population densities Heterodera schachtii on yield of cabbage and table beets in New York State. Phytopathology, 70: 481-485. http://dx.doi.org/10.1094/Phyto-70-481.

13. Roberts PA, Thomason IJ, McKinney HE. 1980. Influence of nonhosts, crucifers and fungal parasites on field populations of Heterodera schachtii. Journal of Nematology, 13: 164-171. PMid:19300739.

14. Arndt M. 2002. Einfluss von Fruchtfolge, chemischen und biologischen Bekämpfungsmaßnahmen auf die Befallsentwicklung von Rübennematoden (Heterodera schachtii) und den Zuckerrübenertrag. Gesunde Pflanzen, 54: 74-79. http://dx.doi.org/10.1046/j.1439-0345.2002.02003.x.

15. Held LJ, Jennings JW, Koch DW, Gray FA. 2000. Economics of trap cropping for sugar beet nematode control. Journal of Sugar Beet Research, 37: 45-55. http://dx.doi.org/10.5274/jsbr.37.1.45.

16. Nielsen EL, Baltensperger DD, Kerr ED, Rife CL. 2003. Host suitability of rapeseed for Heterodera schachtii. Journal of Nematology, 35: 35-38. PMid:19265972.

17. Niere B. 2008. Grundlagen einer nachhaltigen Bekämpfung von Rübenzystennematoden. Sugar Industry, 133: 53-60.

18. Landwirtschaftlicher Informationsdienst Zuckerrübe – LIZ. 2017. Ausfallrapsmanger [on line]. Accessed on March 19, 2017. Available from: http://raps.rheinmedia.de/raps/index.php

19. Daub M, Westphal A. 2011. Integriertes nematodenmanagement in fruchtfolgesystemen mit zuckerrüben. Special Issue Sugar Industry, 136: 41-50.

20. Diederichsen E, Frauen M, Ludwig-Müller J. 2014. Clubroot disease management challenges from a German perspective. Canadian Journal of Plant Pathology Supplement, 36: 85-89. http://dx.doi.org/10.1080/07060661.2013.861871.

21. Wang K-H, Sipes BS, Schmitt DP. 2002. Crotalaria juncea as a cover crop for nematode management: a review. Nematropica, 32: 35-57.

22. Hinds J, Wang KH, Marahatta SP, Meyer SL, Hooks CRR. 2013. Sunn hemp cover cropping and organic fertilizer effects on the nematode community under temperate growing conditions. Journal of Nematology, 45: 265-271. PMid:24379485.

23. Curto G, Dallavalle E, Santi R, Casadei N, D’Avino L, Lazzeri L. 2015. The potential of Crotalaria juncea L. as a summer green manure crop in comparison to Brassicaceae catch crop for management of Meloidogyne incognita in the Mediterranean area. European Journal of Plant Pathology, 142: 829-841. http://dx.doi.org/10.1007/s10658-015-0655-2.

24. Leibniz-Informationszentrum Technik und Naturwissenschaften und Universitätsbibliothek – TIB. 2007. Council Directive 2007/33/EC of 11 June 2007 on the control of potato cyst nematodes and repealing Directive 69/465/EEC. Official Journal of the European Union, 156: 12-22.

25. Seinhorst JW. 1970. Separation of Heterodera cysts from organic debris in ethanol 96%. Nematologica, 16: 330-330.

26. Müller J. 1980. Ein verbessertes Extraktionsverfahren für Heterodera schachtii. Nachrichtenblatt des Deutschen Pflanzenschutzdienstes, 32: 21-24.

27. Byrd DW Jr, Kirkpatrick T, Barker KR. 1983. An improved technique for clearing and staining plant tissues for detection of nematodes. Journal of Nematology, 15: 142-143. PMid:19295781.

28. Müller J, Rumpenhorst HJ. 2000. Rübenzystennematoden: Die Prüfung von Pflanzen auf ihre Widerstandsfähigkeit gegen Schadorganismen in der Biologischen Bundesanstalt Teil 1 - Prüfung von Kulturpflanzen auf Resistenz gegen pflanzenparasitäre Nematoden. Mitteilungen aus der Biologischen Bundesanstalt für Land- und Forstwirtschaft, 372: 9-22.

29. Meinecke A, Hermann A, Westphal A. 2013. Using a selective fast turn-around bioassay for population density determination of Heterodera schachtii. Nematology, 15: 809-819.

30. Niere B. 2006. Zur Beurteilung der Resistenz von Kartoffelsorten gegen die Kartoffelzystennematoden Globodera pallida und Globodera rostochiensis. In: J. Hallmann und B. Niere (eds.). Aktuelle Beiträge zur Nematodenforschung. Vol. 404. Mitteilungen aus der Biologischen Bundesanstalt für Land- und Forstwirtschaft, Quedlinburg, p. 31-39.

31. Subbotin S, Mundo-Ocampo M, Baldwin JG. 2010. Systematics of cyst nematodes (Nematoda: Heteroderinae). In: Hunt DJ, Perry RN (eds.). Nematology Monographs and Perspectives. Vol. 1-2. Leiden, Brill. 512 pp.

32. Kushida A, Suwa N, Ueda Y, Momota Y. 2003. Effects of Crotalaria juncea and C. spectabilis on hatching a population density of the soybean cyst nematode, Heterodera glycines (Tylenchida: Heteroderidae). Applied Entomology and Zoology, 38: 393-399. http://dx.doi.org/10.1303/aez.2003.393.

33. Evans K, Haydock PPJ. 2000. Potato cyst nematode management: present and future. Aspects of Applied Biology, 59: 91-97.

34. Scholte K. 2000. Screening of non-tuber bearing Solanaceae for resistance and induction of juvenile hatch of potato cyst nematodes and their potential for trap cropping. Annals of Applied Biology, 136: 239-246. http://dx.doi.org/10.1111/j.1744-7348.2000.tb00030.x.

35. Ngala BM, Haydock PPJ, Woods S, Back MA. 2015. Biofumigation with Brassica juncea, Raphanus sativus and Eruca sativa for the management of field populations of the potato cyst nematode Globodera pallida. Pest Management Science, 71: 759-769. PMid:24965697. http://dx.doi.org/10.1002/ps.3849.

36. Stallings A. 2015. Sunn hemp (Crotalaria juncea L.) as a Cover Crop for Winter Wheat [dissertation]. Auburn University, Auburn, 161 pp.

37. Ratnadass A, Fernandes P, Avelino J, Habib R. 2012. Plant species diversity for sustainable management of crop pests and diseases in agroecosystems: a review. Agronomy for Sustainable Development, 32: 273-303. http://dx.doi.org/10.1007/s13593-011-0022-4.

38. Price A, Kelton J, Mosjidis J. 2012. Utilization of sunn hemp cover crops and weed control in temperate climates. In: Price A (ed.). Weed control. InTech Europe Rijeka, Croatia, p. 101-113.

39. Collins A, Chase CA, Stall WM, Hutchinson CM. 2008. Optimum densities of three leguminous cover crops for suppression of smooth pigweed (Amaranthus hybridus). Weed Science, 56: 753-761. http://dx.doi.org/10.1614/WS-07-101.1.

11http://dx.doi.org/10.4322/nematoda.00417

Crotalaria juncea and nematodesSedaghatjoo et al.

Nematoda, e042017

40. Morris JB, Chase C, Treadwell D, Koenig R, Cho A, Morales-Payan JP, Murphy T, Antonious GF. 2015. Effect of sunn hemp (Crotalaria juncea L.) cutting date and planting density on weed suppression in Georgia, USA. Journal of Environmental Science and Health. Part. B, Pesticides, Food Contaminants, and Agricultural Wastes, 50: 614-621. PMid:26065522. http://dx.doi.org/10.1080/03601234.2015.1028855.

41. Mosjidis JA, Wehtje G. 2011. Weed control in sunn hemp and ist ability to suppress weed growth. Crop Protection, 30: 70-73. http://dx.doi.org/10.1016/j.cropro.2010.08.021.

42. Colegate SM, Gardner DR, Joy RJ, Betz JM, Panter KE. 2012. Dehydropyrrolizidine alkaloids, including monoesters with an unusual esterifying acid, from cultivated Crotalaria juncea (Sunn hemp cv. ‘Tropic Sun’). Journal of Agricultural and Food Chemistry, 60: 3541-3550. PMid:22429238. http://dx.doi.org/10.1021/jf205296s.

43. Ji X, Khan I, Mosjidis JA, Wang H, Livant P. 2005. Variability for the presence of pyrrolizidine alkaloids in Crotalaria juncea L. Pharmazie, 60: 620-622. PMID:16124407.

44. Wang K-H, Sipes BS, Schmitt DP. 2001. Suppression of Rotylenchulus reniformis by Crotalaria juncea, Brassica napus, and Tagetes erecta. Nematropica, 31: 235-249.

45. Germani G, Plenchette C. 2005. Potential of Crotalaria species as green manure crops for the management of pathogenic nematodes and beneficial mycorrhizal fungi. Plant and Soil, 266: 333-342. http://dx.doi.org/10.1007/s11104-005-2281-9.

46. Marla SR, Huettel RN, Mosjidis J. 2008. Evaluation of Crotalaria juncea populations as hosts and antagonistic crops to manage Meloidogyne incognita and Rotylenchulus reniformis. Nematropica, 38: 155-161.

47. Khandelwal V, Sorathiya MB, Chauhan DA. 2014. Evaluation of agronomic and nitrogen fixation traits in sunnhemp, Crotalaria juncea L. Bioscience Biotechnology Research Communication, 7: 108-113.

48. Cook BG, Pengelly BC, Brown SD, Donnelly JL, Eagles DA, Franco MA, Hanson J, Mullen BF, Partridge IJ, Peters M, Schultze-Kraft R. 2005. Tropical Forages: an interactive selection tool: Crotalaria juncea [CD-ROM]. CSIRO, DPI&F(Qld), CIAT and ILRI, Brisbane. Accessed on 19 April 2016. Available from: http://www.tropicalforages.info/key/Forages/Media/Html/Crotalaria_juncea.htm

49. Schomberg HH, Martini NL, Diaz-Perez JC, Phatak SC, Balkcom KS, Bhardwaj HL. 2007. Potential for using sunn hemp as a source of biomass and nitrogen for the Piedmont and Coastal Plain regions of the southeastern USA. Agronomy Journal, 99: 1448-1457. http://dx.doi.org/10.2134/agronj2006.0294.

50. Danahap LS, Wonang DL. 2016. Antinematicidal efficacy of root exudates of some Crotalaria species on Meloidogyne incognita (root-knot nematode) (Kofoid and White) Chitwood isolated from infected Lycopersicum esculentum L. (tomato) plant. International Journal of Science Technology Research, 5: 79-84.

51. Wang K-H, McSorley R, Gallaher RN, Kokalis-Burelle N. 2008. Cover crops and organic mulches for nematode, weed and plant health management. Nematology, 10: 231-242. http://dx.doi.org/10.1163/156854108783476412.

52. Cook CG, Scott AW Jr, Chow P. 1998. Planting date and cultivar effects on growth and stalk yield of sunn hemp. Industrial Crops and Products, 8: 89-95. http://dx.doi.org/10.1016/S0926-6690(97)10013-9.

53. United States Department of Agriculture – USDA. 2012. Plant Guide: SUNN HEMP [on line]. Natural Resources Conservation Service, Washington. Accessed on 11 June 2016. Available from: http://plants.usda.gov/plantguide/pdf/pg_crju.pdf

54. Daimon H. 2006. Traits of the genus Crotalaria used as a green manure legume on sustainable cropping systems. Japan Agricultural Research Quarterly, 40: 299-305. http://dx.doi.org/10.6090/jarq.40.299.

55. Zahran HH. 1999. Rhizobium-legume symbiosis and nitrogen fixation under severe conditions and in an arid climate. Microbiology and Molecular Biology Reviews, 63: 968-989. PMid:10585971.