Influence of crop production practices on Pasteuria penetrans and suppression of Meloidogyne incognita

Accepted Manuscript Influence of crop production practices on Pasteuria penetrans and suppression of Meloidogyne incognita Patricia Timper, Chang Liu, Richard F. Davis, Tiehang Wu PII: DOI: Reference:

S1049-9644(16)30063-9 http://dx.doi.org/10.1016/j.biocontrol.2016.04.013 YBCON 3426

To appear in:

Biological Control

Received Date: Revised Date: Accepted Date:

11 September 2015 11 April 2016 29 April 2016

Please cite this article as: Timper, P., Liu, C., Davis, R.F., Wu, T., Influence of crop production practices on Pasteuria penetrans and suppression of Meloidogyne incognita, Biological Control (2016), doi: http://dx.doi.org/10.1016/ j.biocontrol.2016.04.013

This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.

1 Timper et al.

Influence of crop production practices on Pasteuria penetrans and suppression of Meloidogyne incognita

Patricia Timpera,*, Chang Liub, Richard F. Davisa, and Tiehang Wuc a

USDA ARS, Crop Protection of Management Research Unit, Tifton, GA

b

Department of Plant Pathology, University of Georgia, Tifton, GA

c

Department of Biology, Georgia Southern University, Statesboro, GA

* Corresponding author. E-mail address: [email protected] FAX: 229-387-2321 Phone: 229-387-2377 Mailing address: P.O. Box 748, Tifton, GA 31793, USA

2 Timper et al.

ABSTRACT Pasteuria penetrans is a parasite of root-knot nematodes (Meloidogyne spp.). Infected nematodes are not killed by the bacterium, but instead of producing eggs, females produce millions of infectious endospores. In addition to sterilizing females, P. penetrans can reduce nematode infection of roots when spore densities in soil are high because juveniles become so heavily encumbered, movement is restricted. A 4-year field study was conducted to determine 1) if fumigation with 1,3-dichloropropene (1,3-D, 28 L/ha) would have a negative effect on P. penetrans and general suppression of nematodes, 2) if the occasional use of 1,3-D would be as detrimental to P. penetrans as a yearly application, 3) if tillage influenced the abundance of P. penetrans spores, and 4) if the P. penetrans at the field site was contributing to suppression of M. incognita. Fumigation with 1,3-D reduced the abundance of P. penetrans spores in the soil compared to the no-fumigation control and there was no difference between a yearly application of the fumigant and occasional applications. Tillage (conventional and strip) did not affect spore abundance. The reduction of P. penetrans spores by fumigation was small compared to the year-to-year fluctuations in spore densities. Spores per assay nematode varied from 6.3 in 2012 to 0.8 in 2014. In 2012, P. penetrans appeared to suppress populations of M. incognita to very low levels (≤ 10% of the root system with galls). The extreme changes in spore densities over the course of this study were likely due to a combination of factors including the density-dependent dynamics between the nematode and bacterium, the self-limiting effect of the bacterium at high spore densities, and leaching of spores out of the root zone during rain and irrigation events. Within each fumigation treatment, there was an inverse relationship (P ≤ 0.003, r = -0.40 to -0.58) between spore abundance and root galling indicating that the bioassay to estimate spore numbers in the spring was a good predictor of the level of nematode suppression.

3 Timper et al.

Keywords: Gossypium hirsutum, nematode suppression, Meloidogyne incognita, Pasteuria penetrans, Root-knot nematode

4 Timper et al.

1. Introduction Numerous organisms are capable of parasitizing or preying on plant-parasitic nematodes in soil. Many of these organisms, such as predatory nematodes and mites as well as trapping fungi tend to be generalists, consuming many different types of nematodes. Generalist parasites and predators of nematodes are common to most agricultural soils and have been shown to result in lower populations of plant-parasitic nematodes than would occur in the absence of these generalists. In contrast to the generalists, a smaller subset of antagonistic organisms are specialized for parasitizing a particular taxon of plan-parasitic nematodes. Bacteria in the genus Pasteuria are among the most host-specific of these specialists (Stirling, 2014). Pasteuria penetrans is a parasite of root-knot nematodes (Meloidogyne spp.). Under natural conditions, the bacterium can only complete its lifecycle within a host nematode. Second-stage juveniles (J2) of Meloidogyne spp. acquire the adhesive endospores of P. penetrans as they migrate through soil in search of a plant host. After a spore-laden J2 establishes a permanent feeding site within the vascular system of a root, the bacterium produces a germination tube penetrating the nematode cuticle and then grows vegetatively within the nematode body. The infected nematode is not killed and continues to develop into an adult. Endospores are produced within the body of the female and are released into the soil when the nematode cuticle is ruptured. In addition to sterilizing females, P. penetrans can reduce nematode infection of roots when J2 become so heavily encumbered with spores, movement is restricted (Davies et al., 1991; Stirling, 1984). Suppression of Meloidogyne populations by P. penetrans has been documented in both annual and perennial crops; the level

5 Timper et al.

of nematode suppression can be as high as 80-90% (Bird and Brisbane, 1988; Chen et al., 1996; Kariuki and Dickson, 2007; Oostendorp et al., 1991; Stirling, 1984; Timper et al., 2001; Weibelzahl-Fulton et al., 1996). Crop production practices such as crop rotation, pesticides, and tillage can have indirect and direct effects on P. penetrans and generalist predators and parasites of nematodes. Because P. penetrans only reproduces on Meloidogyne spp., any pest management practice which reduces populations of this nematode will reduce recruitment of new hosts for the bacterium and production of spores. Several studies have shown that both crop rotation with nonhosts of Meloidogyne spp. (Madulu et al., 1994; Timper et al., 2001; Timper, 2009) and the application of nematicides (Kariuki and Dickson, 2007; Timper et al., 2012) resulted in lower spore densities of P. penetrans than when continuous hosts for the nematode were grown and no nematicides were used. Similarly, pesticides and tillage can have a direct negative effect on the organisms involved in general suppression of plant-parasitic nematodes. Following application of nematicides, carnivorous nematodes (predators and omnivores) were reduced by 75% 2 weeks after application compared to the no-nematicide control (Timper et al., 2012). Suppression of these carnivores corresponded to diminished general suppression of the reniform nematode in a bioassay. Predatory mites and carnivorous nematodes are particularly sensitive to the physical disturbance that occurs when soil is tilled and their populations are often lower in conventional tillage than in no tillage (Koehler, 1999; Lenz & Eisenbeis, 2000; Okada and Harada, 2007; Sanchez-Moreno et al., 2009; Wardle et al., 1995). In a previous field study, we focused on the effect of tillage and 1,3-D application on carnivorous nematodes; the presence of P. penetrans at the field site was noted only after the

6 Timper et al.

study was initiated (Timper et al., 2012). Nevertheless, we were able collect data showing that spore densities were lower following repeated application of 1,3-D. The rate of fumigant used (56 L/ha), however, was greater than the recommended rate for cotton in Georgia. Our objectives in this study were to determine 1) if the recommended rate of 1,3-D (28 L/ha) would also have a negative effect on P. penetrans and general suppression of nematodes, 2) if the yearly application of 1,3-D is more detrimental to P. penetrans than less frequent applications, 3) if tillage influenced the abundance of P. penetrans spores, and 4) if the P. penetrans at the field site was contributing to suppression of M. incognita. 2. Materials and methods 2.1.

Study site and experimental design

The field site was the same as described in an earlier study (Timper et al., 2012). The soil was a Tifton loamy sand (85% sand, 11% silt, 4% clay, and 1% organic matter). A split-plot design with six replications was used to determine the effect of tillage (main plots, two treatments) and 1,3-D application (subplots, five treatments) on abundance of P. penetrans spores, general suppression of plant-parasitic nematodes, and suppression of M. incognita. The tillage treatments were conventional tillage (rip-bedded) and strip tillage. Strip tillage is commonly used in cotton production in the southeastern United States and is considered a conservation tillage practice. The subplot treatments were the following frequency of 1,3-D application (= F) or no fumigant control (= C) over a 4-year period: F-F-F-F, C-F-F-C, F-C-F-C, F-FC-F, and C-C-C-C. The tillage and fumigant treatments were not re-randomized every year.

7 Timper et al.

The main plots were 7.3 m wide by 15.2 m long, and each subplot consisted of two rows spaced 91 cm apart. There were two border beds on the field margins. In the spring, the conventional plots were harrowed using an S-tine cultivator and then a ripper-bedder was used to form the planting bed. The ripper-bedder has a single sub-soil chisel per row which tears and loosens the soil to a depth of 41 cm. Strip tillage also has a single sub-soil chisel per row with shallow (10 cm) discs and rollers which make a smooth seed-bed 25-cm wide; the remaining space between rows is undisturbed. In both conventional and strip tillage, the fumigant 1,3dichloropropene (1,3-D; Telone II®, Dow AgroSciences) was applied during tillage by injecting 28 L/ha (33.1 kg a.i./ha) 41-cm deep under the rows 9 – 16 days before planting cotton. Cotton (Gossypium hirsutum) was planted on 26 May 2011, 10 May 2012, 30 May 2013, and 3 June 2014. The cotton cultivars planted varied among years, but were all susceptible to M. incognita. The cultivars were Delta and Pine Land DP0935 in 2011 and 2012, DP1048 in 2013, and Phytogen 499 in 2014. Applications of fertilizer, insecticides and herbicides followed University of Georgia Extension Service recommendations and were the same for all plots. Irrigation was applied as needed through overhead sprinklers to the cotton crop. Cotton was harvested on 14 November 2011, 2 November 2012, 21 November 2013, and 9 December 2014. Rye (Secale cereale) ‘Wrens Abruzzi’ was planted in the winter at a broadcast seeding rate of 101 kg/ha. Nitrogen (34 kg/ha) was applied to the rye approximately 1 month after planting to increase the biomass of the rye for building organic residue in the soil and improving weed suppression (Webster et al., 2013). 2.2.

Meloidogyne incognita data

8 Timper et al.

Soil samples to determine population densities of M. incognita J2 were collected before tillage and fumigation with 1,3-D (19 April 2011, 27 April, 2012, and 14 May 2013 and 2014), post plant (31 May 2011, 21 May 2012, 13 June 2013, and 2 July 2014), and after cotton harvest (1 December 2011, 19 November 2012, 16 December 2013, and 17 December 2014). Soil samples consisted of a composite of 10 to 12 cores per subplot (2.5-cm diam. and 20-cm deep) collected from the root zone. Nematodes were extracted from 150 cm3 soil by centrifugal flotation (Jenkins, 1964); the number of J2 were converted to 100 cm3 of soil for presentation. In the fall sample (after cotton harvest), 20 random J2 were examined for the number of attached P. penetrans spores and the average spores per J2 was determined for each subplot. Hereafter, these are referred to as spores per native J2. Root galling on cotton caused by M. incognita was determined after harvest by rating eight root systems from each subplot on a 010 scale based on the percentage of the root system with galls, where 0 = no galling, 1 = 1 to 10% of the root system galled, 2 = 11 to 20%, etc., with 10 = 91 to 100%. The average of these eight ratings were compared with statistical analysis. 2.3.

Bioassays

2.3.1. Pasteuria bioassay The abundance of P. penetrans in a 100 cm3 subsample of soil from each subplot was determined using a bioassay described by Timper et al. (2001). The Pasteuria bioassay was conducted post plant in the spring using soil collected for determining population densities of M. incognita J2. Prior to the bioassay, the soil was heated at 50 ˚C for 3 hours to kill native J2, but not P. penetrans spores (Chen et al., 1995). Nematodes for the bioassay were obtained from a greenhouse culture of M. incognita by placing roots with egg masses in a mist chamber

9 Timper et al.

and collecting the hatched J2 3 to 4 days later. A soil subsample from each plot was added to a 250 cm3 flask along with water. The mixture was shaken, allowed to settle for 5 sec, and the soil-water suspension decanted into another flask. The flasks containing the soil-water suspension were inoculated with 1,500 J2 and shaken for 24 hours. The nematodes were extracted by centrifugal flotation and the number of spores adhering to 20-30 individual J2, selected at random, was determined using an inverted microscope at 400x magnification. 2.3.2. Nematode survival bioassay Starting in 2012, soil samples were collected for a survival bioassay from conventional tillage plots treated every year with 1,3-D (F-F-F-F) and control plots (C-C-C-C) for a total of 12 samples collected at three times: pre-fumigation (same day as samples for M. incognita), at plant (22 May 2012, 6 June 2013, and 25 June 2014), and at midseason (28 August 2012, 27 August, 2013, 17 September 2014). Soil samples consisted of a composite of 5 cores per plot (6.5-cm diam and 16.3-cm deep) collected from the root zone. The bioassay, which was intended to measure general suppression of plant-parasitic nematodes, was previously described by Timper et al. (2009). Briefly, a subsample of soil was split evenly between two jars. The soil in one jar was defaunated by heating for 1 hour in an oven at 65 °C; the other jar was left unheated (natural). The jars were incubated for 5 days before transferring 2000 vermiform stages of the reniform nematode (Rotylenchulus reniformis) in < 1 ml of water into each of the jars. The soil at the time of nematode transfer was moist; therefore, no additional water was added. The nematodes were extracted by centrifugal flotation 5 days later and the number of live reniform nematodes in the natural and heated soil was counted.

10 Timper et al.

2.3.3. Population increase of M. incognita This bioassay was intended to measure specific suppression of M. incognita by P. penetrans. Soil samples to determine the reproductive potential of the nematode were collected within 5 days after planting except in 2014, when samples were collected 1 month after planting. A shovel was used to remove the top 2.5 cm of soil near the planting furrow and collect approximately 0.5 L of soil to a depth of 15 cm from three locations per plot. The samples were placed in metal trays and heated to 50 ˚C for 3 hours to kill native M. incognita, but not P. penetrans. A single pot per field replicate was prepared by adding 850 cm3 of the heated soil to a 10 cm square pot. The pots were left in the greenhouse for 7 days to dissipate toxins released during the heating process and then two cotton (same cultivar as planted in the field) seed were planted and thinned to one plant after emergence. Two weeks after planting, the pots were inoculated by pipetting 3500 J2 of M. incognita into two holes (2 cm deep) at the base of the plant. The bioassay was terminated 60 days after nematode inoculation (approximately two nematode generations). The root systems were rinsed with water to remove soil, patted dry, and weighed. Eggs were extracted from the roots by cutting the roots into approximately 5-cm pieces, placing them in a 1-liter flask, and agitating for 4 minutes in a 1.2% NaOCl solution (Hussey and Barker, 1973). 2.4.

Statistical analyses

All statistical analyses were performed using JMP Pro (v. 11, SAS Institute). To determine the effect of tillage and fumigation on P. penetrans spores per assay J2 in the spring and per native J2 in the fall and reproduction of M. incognita, mixed model analysis was used. In the

11 Timper et al.

model tillage, fumigation, and tillage*fumigation were classified as fixed effects and year, replication, year*tillage, year*fumigation, year*tillage*fumigation, replication (year), and replication*tillage (year) were classified as random effects in the model. Year was classified as a random effect because fumigation with 1,3-D does not have a direct effect on P. penetrans spores (Mankau and Prasad, 1972) and we were interested in generalizing the effects of tillage and fumigation across all years. Eggs per gram of root from the reproduction bioassay was transformed by square root prior to statistical analysis. Mixed model analysis was also used to determine the effect of fumigation and soil treatment (defaunated vs natural) on survival of the reniform nematode using a similar classification of fixed and random effects as described previously. For root gall indices, densities of M. incognita J2 in the soil, and yield, year was classified as a fixed effect because 1,3-D was not applied in all years of some of the treatments (e.g., F-C-F-C). Correlation analysis was used to determine the relationships between abundance of P. penetrans and root galling, the density of M. incognita J2 in soil, and cotton yield across years but within a fumigation treatments. Four different measures of P. penetrans abundance were used in the analysis: spores per assay J2 in the spring, spores per native J2 in the fall, and the percentage of J2 with spores in the spring (assay) and fall (native). Because the percentage of J2 with spores always provided a better fit for the data than the spores per J2, only the correlation analysis on the former measure will be presented. Correlation analysis was also used to determine the relationship between root galling and yield both within a year and among years. 3. Results

12 Timper et al.

Abundance of P. penetrans spores in the spring varied considerably from year to year (Fig. 1), ranging from 6.3 to 0.8 spores/assay J2 in 2012 and 2014, respectively, when averaged across tillage and fumigation treatments. The percentage of J2 acquiring at least one spore was 47, 86, 65, and 21 in 2011, 2012, 2013, and 2014, respectively. Similarly, root gall indices fluctuated from 0.7 in 2012 to 3.4 in 2014 when averaged across tillage and fumigation treatments. 3.1.

Effect of fumigation and tillage on P. penetrans

In the mixed model analysis, abundance of P. penetrans spores was affected by fumigation (P = 0.044) but not by tillage, and the effects of fumigation were consistent across tillage treatments (i.e., no tillage*fumigation interaction). The number of spores per assay J2 was lower in most of the 1,3-D treatments than in the control; however, the F-C-F-C treatment was intermediate between the control and other 1,3-D treatments (Fig. 2). The number of spores per native J2 from the fall sample was considerably lower than the spores detected in the spring bioassay (approximately 1/5); nevertheless, the trends in spores per native J2 were the same as the spores per assay J2 (Fig. 2). Tillage did not affect the spores per native J2. There was a positive, but weak correlation (P = <0.0001, r = 0.26) between spores per assay J2 in the spring and spores per native J2 in the fall. 3.2.

Root galling by M. incognita Galling of cotton roots caused by M. incognita was not affected by tillage, but was affected

(P < 0.0001) by fumigation with 1,3-D and the effect of fumigation was not consistent among years (year*fumigation, P = 0.0002). In years where no fumigation was applied to a treatment

13 Timper et al.

(e.g., F-F-C-F in 2013), galling from M. incognita was not different from the C-C-C-C treatment (Table 1). In years where fumigation was applied, galling was typically reduced compared to CC-C-C. The exception was in 2014 when only the F-F-F-F treatment differed from the control. Within a fumigation treatment, there was an inverse correlation between the percentage of assay J2 with spores in the spring and root galling in the fall (Fig. 3; Table 2); the greater the percentage of J2 with spores in the spring, the lower the root galling in the fall. There was also an inverse correlation between the percentage of native J2 with spores and galling, but only in the C-C-C-C, F-F-C-F, and F-F-F-F plots (Table 2). 3.3.

Population densities of M. incognita J2

For all sampling times, the effects of fumigation on population densities of M. incognita J2 was not consistent among years (year*fumigation, P ≤ 0.003); therefore, the data were analyzed by year. In the pre-fumigation samples, tillage did not influence densities of J2 in soil; however, fumigation had residual effects in 2012 and 2013, with lower (P < 0.05) densities of J2 in plots that had been fumigated the previous year than in the C-C-C-C plots (data not shown). In the post-plant samples, fumigation reduced (P ≤ 0.0001) densities of J2 compared to the C-CC-C in all years except 2014. Across years, numbers of J2/150 cm3 of soil at plant were 73, 32, 11, 9, and 5 in the C-C-C-C, C-F-F-C, F-C-F-C, and F-F-F-F treatments, respectively. Tillage did not influence densities of J2 at plant except in 2011, when densities were greater in strip than in conventional tillage in the C-F-F-C plots (tillage*fumigation, P = 0.009). In the post-harvest soil samples, densities of J2 among fumigation treatments differed only in 2011 and 2013. In 2011, plots receiving 1,3-D in the spring had lower densities of J2 compared to C-C-C-C plots (Table 3). In 2013, only the F-C-F-C plots had lower densities than the C-C-C-C plots; the F-F-C-F plots had

14 Timper et al.

the greatest densities of J2 because they had not been fumigated that year. Tillage did not affect J2 densities except in 2012, when densities were greater in conventional than in strip tillage (290 vs 168 J2/150 cm3 soil). When analyzed within a fumigation treatment, there was no correlation between the percentage of assay J2 with P. penetrans spores in the spring and densities of J2 in samples collected at pre-fumigation, post plant, or after cotton harvest. However, there were positive correlations between the percentage of native J2 with spores and the densities of J2 at all sampling times. With increasing densities of J2, there was an increase in the percentage of native J2 with spores in the pre-fumigation samples for the C-C-C-C (P = 0.007, r = 0.37), C-F-F-C (P = 0.02, r = 0.32), and F-F-C-F (P = 0.0007, r = 0.47) plots; and in the CC-C-C-C plots in the post-plant (P = 0.03, r = 0.28)and after harvest samples (P = 0.02, r = 0.31). 3.4.

Cotton yield

Cotton yield differed (P < 0.0001) among years with yields averaging 1276, 1204, 1664, and 1347 kg lint/ha in 2011, 2012, 2013, and 2014, respectively. The effect of fumigation treatment was not consistent among years (year* fumigation, P = 0.016). In 2011, yield was greatest in plots that had been fumigated with 1,3-D (Table 4). In 2012 and 2013, yields were greater in all the fumigation treatments and lowest in the non-fumigated control. The trends were similar in 2014, though they were not significant (P = 0.06). The effect of tillage on yield was not consistent among years (year*tillage, P < 0.0001). In 2012 and 2013, cotton yield was greater (P ≤ 0.01) in conventional than in strip tillage (data not shown). When analyzed across years, galling was not correlated with yield; however, within a year, galling was negatively correlated with yield in all years; the greater the galling the lower the yield of lint per ha. The strongest correlation between galling and yield was in 2011 (P = 0.0003, r = -0.43) and the

15 Timper et al.

weakest correlation was in 2013 (P = 0.03, r = -0.25). The percentage of assay J2 with P. penetrans spores in the spring was negatively correlated with yield (P = 0.04, r = -0.27) only in the C-C-C-C plots. However, the percentage of native J2 with spores in the fall were negatively correlated with yield in all fumigation treatments; with increasing percentage of J2 with spores, there was a decrease in yield (P ≤ 0.02, r = -0.31 to -0.48). Because the abundance of P. penetrans in the fall is more likely to affect the yield in the subsequent year, the relationship between the percentage of native J2 with spores and yield in the following year was determined. In the new analysis, there was a positive correlation between the percentage of J2 with spores and yield for the C-F-F-C (P = 0.03, r = 0.33) and the F-C-F-C (P = 0.006, r = 0.43) plots. 3.5.

Reproductive potential of M. incognita

When year was a random effect in the mixed model, neither tillage, fumigation, nor their interaction affected reproduction of M. incognita in the soil. However, abundance of P. penetrans varied considerably among the years which may have influenced the outcome of the reproduction bioassay. Therefore, year was assigned as a fixed effect to determine whether year influenced the effect of tillage and fumigation on reproduction of M. incognita. There was a year*tillage (P = 0.0006) and a year*tillage*fumigation (P = 0.007) interaction on nematode reproduction. When analyzed within a year, there were no effects of tillage, fumigation, or their interaction on nematode reproduction in 2011 or 2012. In 2013 fumigation affected (P = 0.0006) reproduction of M. incognita with the greatest reproduction in the F-C-F-C treatment compared to all other fumigation treatments (data not shown). In 2014, soil from the strip

16 Timper et al.

tillage treatment had greater (P = 0.048) reproduction of M. incognita than soil from conventional tillage (data not shown). 3.6.

Nematode survival bioassay

Survival of the reniform nematode was lower (P ≤ 0.008) in the natural soil than in the heated soil for the pre-fumigation and the at-plant soil samples, but not the midseason samples. Compared to the heated soil, nematode survival at all sampling times was reduced by 33-34%. However, yearly fumigation with 1,3-D did not influence survival of the reniform nematode on any of the sample dates. 4. Discussion Fumigation with 1,3-D significantly reduced densities of M. incognita J2 at planting time leading to reduced root galling and generally greater yield of cotton. These results are in agreement with other studies indicating that 1,3-D is an effective nematicide for suppressing populations of M. incognita in sandy soils and improving cotton yield (Ortiz et al., 2012; Overstreet et al., 2014; Thomas and Smith, 1993). The only year in which 1,3-D did not increase cotton yield was in 2014; the fumigant was also less effective in reducing M. incognita populations at plant and root galling compared to the other years. The soil conditions in 2014 may not have been ideal for 1,3-D diffusion through the soil pores because of substantial rainfall (11.4 cm) 5-6 days prior to fumigant application (Thomas et al., 2004). Contrary to our previous study, we did not observe greater galling in conventional compared to strip tillage (Timper et al., 2011). Perhaps the presence of P. penetrans in the soil confounded our ability to detect an effect of tillage on nematode populations. Root galling was negatively correlated

17 Timper et al.

with cotton yield within a year, but not among years, suggesting that the variation in yield among years was due to environmental factors other than M. incognita. Application of 1,3-D in two or more consecutive years reduced the abundance of P. penetrans spores in soil compared to non-fumigated soil. Previous studies showing a reduction in P. penetrans following fumigation with 1,3-D used a high rate of the chemical (Kariuki and Dickson, 2007; Timper et al., 2012). Our study demonstrates that even the low rate of 1,3-D (28 L/ha) recommended for cotton reduces the abundance P. penetrans in soil. However, the yearly application of 1,3-D did not reduce populations of the bacterium more than the alternate year fumigation. The fumigant does not have a direct effect on P. penetrans spores (Mankau and Prasad, 1972), but likely had an indirect effect by reducing the number of hosts available for reproduction of the bacterium. We had anticipated lower densities of P. penetrans spores in conventional tillage compared to strip tillage. Both the conventional and strip tillage rips the soil with a chisel to prepare the planting furrow. This furrow was in the same location in all years of the study; therefore, P. penetrans spores should reach their highest densities in this location. In conventional tillage, the bedding process moves soil from a 45.5-cm area on either side of the furrow and mounds it on top of the chiseled furrow. This surrounding soil should contain lower spore densities than the furrow soil, thus resulting in a dilution of spore densities. However, we were unable to detect an effect of tillage on spore densities. It is possible that tillage had conflicting effects on the nematode and the bacterium. For instance, in previous studies, infection of cotton by M. incognita was lower under strip tillage than under conventional tillage suggesting that conditions were less conducive for nematode infection of roots under strip tillage (Bauer et al.,

18 Timper et al.

2010; Timper et al., 2011). If strip tillage supports greater spore densities than conventional tillage but limits infection of roots by J2 and, therefore, the production of additional spores, then there may be no net effect of tillage on spore densities. In a microplot study, abundance of P. penetrans spores was lower at planting time in soil that had been rototilled than in soil without tillage (Talavera et al., 2002). However, after crop harvest, spore densities did not differ between the tillage treatments. When the effects of tillage were examined at different soil depths after harvest, tillage and tillage plus watering had greater spore abundance at lower depths (20-40 cm) compared to no tillage. Our soil samples were collected only to a depth of 20 cm; consequently, we may have missed differences in abundance of P. penetrans at lower depths. In soybean, suppression of H. glycines by Pasteuria nishizawae was similar in conventional and no tillage plots, though differences in spore densities between tillage treatments were not determined (Noel et al., 2010). We had expected that reproduction of M. incognita in the greenhouse would be lower in the C-C-C-C soil than in soil that had been frequently fumigated because the C-C-C-C soil contained more P. penetrans spores. However, no clear differences in reproduction were observed between treatments containing different spore densities. It is possible that the differences in spore densities in the C-C-C-C and fumigated plots were not great enough to detect differences in nematode reproduction in the greenhouse. Another possibility is that the spores did not persist in the soil. In sandy soils, spores of P. penetrans are readily leached with irrigation water (Dabire and Mateille, 2004; Oostendorp et al., 1990). The pots in the reproduction bioassay were watered at least once per day which may have progressively moved the spores down through the soil and out of the pots.

19 Timper et al.

In a previous study at the same field site, fumigation with 1,3-D reduced general suppression of plant-parasitic nematodes at the time cotton was planted and also at midseason (Timper et al., 2012). In the present study, using the same survival bioassay used in the previous study, we were unable to document an effect of fumigation on general suppression of the reniform nematode. The lower rate of 1,3-D used in this study compared to the previous study may have been less destructive of the soil community. However, the soil was less suppressive than previously reported, which may have made it more difficult to detect the effects of fumigation. Based on survival in the defaunated soil, survival of the reniform nematode in the natural soil was 67% in the present study and 51% in the previous study. The abundance of P. penetrans spores fluctuated considerably among years. In the C-C-C-C plots, spores per assay J2 varied from 11.0 in 2012 to 0.4 in 2014. The significant negative correlation between spores per assay J2 in the spring and root galling in the fall indicates that P. penetrans was suppressing populations of M. incognita at our field site. Several field studies have documented a density-dependent relationship between P. penetrans and root-knot nematodes with rates of parasitism or spore production by the bacterium rising and falling with increases and decreases in the nematode population (Ciancio and Bourijate, 1995; Spaull, 1984; Verdejo-Lucas, 1992). We also observed an increase in spore attachment to native J2 with an increase in populations of J2 in the soil. Thus, the extreme fluctuation in spore densities at our field site can partly be explained by density dependence and by the self-limiting effect of the bacterium at high spore densities. At low densities of P. penetrans, some Meloidogyne J2 acquire one to a few spores, infect roots, and produce 105 - 106 spores/infected female, but produce few to no eggs (Davies et al., 1988; Chen et al., 1994). As the density of spores

20 Timper et al.

increases in soil, more spores are acquired by a greater percentage of J2. Many of the heavily encumbered J2 will fail to penetrate roots resulting in lower root galling. As few as 7 spores/J2 can reduce nematode mobility and penetration of roots (Davies et al., 1991). However, the failure of heavily encumbered J2 to infect roots not only depletes the number of spores in the population, but also limits the production of new spores (Ciancio and Bourijate, 1995). Although endospores of P. penetrans have the potential to survive for several years (Giannakou et al., 1997), they may not persist in sandy soil due to leaching of spores during rain and irrigation events. In a sandy soil, 68% of P. penetrans spores leached 10 cm, whereas in a sandy clay soil, only 39% of spores leached the same distance (Mateille et al., 1996). Increasing the clay content of a sandy soil up to 30% progressively reduced leaching of spores (Dabire and Mateille, 2004). Cetintas and Dickson (2005) also documented substantial leaching of P. penetrans spores in a sandy soil with low clay content; after four irrigation events over a 55-day period, 39-54% of the spores had been leached below 12.5 cm. The soil at our field site was a loamy sand with only 4% clay content. Under these conditions, spores of P. penetrans may be readily flushed from the root zone with irrigation water and rain. Rainfall during the period after cotton harvest and before planting the next cotton crop (15 November to 15 May) was 35.6. 33.0, 76.2, and 63.5 cm in 2011, 2012, 2013, and 2014, respectively. The greater accumulated rainfall in 2013 and 2014 may have contributed to the decline in abundance of P. penetrans spores, particularly if there was a concomitant decrease in spore production. Several studies have found high population levels of Meloidogyne spp. despite the presence of P. penetrans in the populations (Ciancio and Bourijate, 1995; Spaull, 1984; Verdejo-Lucas, 1992). Therefore, it was unclear whether the bacterium was substantially reducing populations

21 Timper et al.

of M. incognita at our field site. Based on 4 years of data, we observed a significant inverse relationship between abundance of spores in the spring and root-gall ratings in the fall indicating that P. penetrans was suppressing populations of M. incognita. Abundance of P. penetrans peaked in 2012 which corresponded to very low gall ratings even in the C-C-C-C plots (1.1 on a 0-10 scale). Conversely, abundance of the bacterium was lowest in 2014 when gall ratings in the C-C-C-C plots reach their highest level (4.1). However, despite the low galling in the C-C-C-C plots in 2012, fumigation with 1,3-D still resulted in increased cotton yield. The fumigant reduces other plant-parasitic nematodes, some fungal plant pathogens, and soil invertebrates (Telone® II label). Perhaps suppression of these other plant pathogens and herbivores contributed to the yield improvements. It is also possible that cotton is sensitive to even low levels of galling (≤ 10%) by M. incognita. It was difficult to relate abundance of P. penetrans to cotton yield because variation in yield among years is due to many factors including differences in climate and plant damage from disease and insects among years in addition to damage from M. incognita. Nevertheless, there was a negative relationship between yield and the percentage of native J2 with spores such that when yields were large, the percentage of J2 with spores in the fall was small. This counter-intuitive trend may be the result of the density-dependent dynamics between the nematode and P. penetrans. When populations levels of M. incognita were lower, spore abundance declined, and yields were greater; however, when populations of the nematode were higher, spore abundance increased, and yields were lower. There was a tendency for greater cotton yields with increasing percentages of J2 with spores in the previous fall samples.

22 Timper et al.

The bioassay to estimate densities of P. penetrans spores in the spring provided a good prediction of root galling in the fall. Similarly, Kariuki and Dickson (2007) observed a positive correlation between spores per assay nematode in the spring and female M. arenaria parasitized by P. penetrans in the fall. Damage thresholds based on the number of Meloidogyne J2 per volume of soil are frequently used by crop consultants and growers to make decisions about whether to apply a nematicide, plant a resistant cultivar, or rotate with a non-host crop for the nematode. Soil samples to estimate damage thresholds are collected soon after crop harvest in the fall because nematode populations may be below detection levels in the spring (Robinson, 2008). In the southeastern United States, damage thresholds for M. incognita in cotton are 50-100 J2/100 cm3 of soil. In our study, densities of J2 after harvest were typically over the damage threshold. Based on this threshold, the 224 J2/100 cm3 of soil in the C-C-C-C plots in 2011 should have led to substantial root galling in 2012. Yet, the rootgall index was very low in 2012 (1 on a 0-10 scale). Bioassays to estimate the abundance of P. penetrans in the soil could be used in conjunction with densities of J2 to more accurately predict crop damage from root-knot nematodes. In our bioassay, we counted the number of spores on 30 random J2; however, counting each spore is time consuming and tedious. The percentage of J2 with spores is easier to determine and was a better predictor of root galling than was spores per J2. Our results support those of Stirling (1984) and Chen et al. (1996) who found that good nematode suppression was achieved when 70-90% or more of bioassay J2 had spores. Further research is underway to determine whether a bioassay for P. penetrans in the fall can predict damage from root-knot nematodes in the subsequent year.

23 Timper et al.

Figure captions

Figure 1. Abundance of Pasteuria penetrans (spores/assay juvenile) in the spring and root-gall indices at harvest in cotton caused by Meloidogyne incognita over 4 years. Data points are the average of all subplots (N = 60). The gall index is a 0-10 scale based on the percentage of the root system with galls, where 0 = no galling, 1 = 1 to 10% of the root system galled, 2 = 11 to 20%, etc., with 10 = 91 to 100%.

Figure 2. Effect of fumigation with 1,3-dichloropropene on the number of Pasteuria penetrans spores per assay juvenile (J2) in the spring and native J2 in the fall. The x-axis indicates the frequency of 1,3-dichloropropene application (= F) or no fumigant control (= C) over a 4-year period. Bars with the same letter are not significantly different (P ≥ 0.05).

Figure 3. Relationship between the percentage of assay juveniles (J2) with Pasteuria penetrans spores in the spring and root galling in the fall caused by Meloidogyne incognita.

24 Timper et al.

Table 1. Effect of fumigation with 1,3-dichloropropene on root gallinga resulting from infection of cotton by Meloidogyne incognita. Year Mean

Fumigationb

2011

C-C-C-C

2.42 ac

1.10 a

2.93 a

4.11 a

2.64 a

F-C-F-C

0.33 b

0.97 ab

1.29 c

3.36 ab

1.49 cd

C-F-F-C

2.43 a

0.56 bc

1.12 c

4.38 a

2.16 b

F-F-C-F

0.62 b

0.35 c

2.26 ab

3.13 ab

1.59 c

F-F-F-F

0.32 b

0.16 c

1.55 bc

2.42 b

1.11 d

P-value

<0.0001

0.0003

0.0004

0.048

< 0.0001

2012

2013

2014

a

The index is a 0-10 scale based on the percentage of the root system with galls, where 0 = no galling, 1 = 1 to 10% of the root system galled, 2 = 11 to 20%, etc., with 10 = 91 to 100%. b

Frequency of 1,3-dichloropropene application (= F) or no fumigant control (= C) over a 4-year period. There was a year*fumigation interaction, P = 0.0002. c

Means within a column with the same letter are not significantly different (P ≥ 0.05).

Table 2. Relationship between the percentage of Meloidogyne incognita juveniles (J2) with Pasteuria penetrans spores in the spring (assay J2) or fall (native J2) and root galling of cotton caused by M. incognita. Fumigationa

Assay J2 with spores

Native J2 with spores

r

P-value

r

C-C-C-C

-0.57

< 0.0001

-0.50

0.0002

F-C-F-C

-0.53

< 0.0001

-0.21

0.08

C-F-F-C

-0.64

< 0.0001

-0.23

0.07

F-F-C-F

-0.43

0.001

-0.51

0.0002

F-F-F-F

-0.54

< 0.0001

-0.44

0.001

a

P-value

Frequency of 1,3-dichloropropene application (= F) or no fumigant control (= C) over a 4-year period.

25 Timper et al.

Table 3. Effect of fumigation with 1,3-dichloropropene on the population density of Meloidogyne incognita juveniles (J2/100 cm3 of soil) after cotton harvest. Year Fumigationa

2011

2012

2013

2014

Mean

C-C-C-C

224 ab

194

192 b

100

177 a

F-C-F-C

105 b

151

87 c

95

110 b

C-F-F-C

267 a

157

137 bc

115

169 a

F-F-C-F

110 b

133

315 a

79

159 a

F-F-F-F

87 b

129

137 bc

60

103 b

P-value

<0.0001

0.0005

0.07

0.001

0.36

a

Frequency of 1,3-dichloropropene application (= F) or no fumigant control (= C) over a 4-year period. There was a year*fumigation interaction, P < 0.0001. b

Means within a column with the same letter are not significantly different (P ≥ 0.05).

Table 4. Effect of fumigation with 1,3-dichloropropene on yield of cotton (kg lint/ha). Year Fumigationa

2011

2012

2013

2014

Mean

C-C-C-C

1115 cb

964 b

1404 b

1285

1192 c

F-C-F-C

1329 ab

1213 a

1818 a

1292

1413 ab

C-F-F-C

1238 bc

1235 a

1678 a

1379

1382 b

F-F-C-F

1300 ab

1302 a

1691 a

1377

1417 ab

F-F-F-F

1397 a

1305 a

1729 a

1402

1458 a

P-value

0.002

< 0.0001

0.0002

0.06

< 0.0001

a

Frequency of 1,3-dichloropropene application (= F) or no fumigant control (= C) over a 4-year period. There was a year*fumigation interaction, P = 0.015. b

Means within a column with the same letter are not significantly different (P ≥ 0.05).

26 Timper et al.

Acknowledgments The authors thank David Clements and Susan Drawdy for technical assistance and Dr. Zhongxiao Chen for reviewing an earlier draft of this manuscript.

References Bauer, P.J., Fortnum, B.A., Frederick, J.R., 2010. Cotton responses to tillage and totation during the turn of the century drought. Agron. J. 102, 1145-1148.

Bird, A.F., Brisbane, P.G., 1988. The influence of Pasteuria penetrans in field soils on the reproduction of root-knot nematodes. Rev. Nematol. 11, 75-81.

Cetintas, R., Dickson, D.W., 2005. Distribution and downward movement of Pasteuria penetrans in field soil. J. Nematol. 37, 155-160.

Chen, S.Y., Dickson, D.W., Mitchell, D.J., 1995. Effects of soil treatments on the survival of soil microorganisms. J. Nematol. 27, 661-663.

Chen, S.Y., Dickson, D.W., Whitty, E.B., 1994. Response of Meloidogyne spp. to Pasteuria penetrans, fungi, and cultural practices in tobacco. J. Nematol. 26, 620-625.

Chen, Z.X., Dickson, D.W., McSorley, R., Mitchell, D.J., Hewlett, T.E., 1996. Suppression of Meloidogyne arenaria race 1 by soil application of endospores of Pasteuria penetrans. J. Nematol. 28, 159-168.

27 Timper et al.

Ciancio, A., Bourijate, M., 1995. Relationship between Pasteuria penetrans infection levels and density of Meloidogyne javanica. Nematol. medit. 23, 43-49.

Dabire, K.R., Mateille, T., 2004. Soil texture and irrigation influence the transport and the development of Pasteuria penetrans, a bacterial parasite of root-knot nematodes. Soil Biol. Biochem. 36, 539-543.

Davies, K.G., Kerry, B.R., Flynn, C.A., 1988. Observations on the pathogenicity of Pasteuria penetrans: a parasite of root-knot nematodes. Ann. Appl. Biol. 112, 491-502.

Davies, K.G., Laird, V., Kerry, B.R., 1991. The motility, development and infection of Meloidogyne incognita encumbered with spores of the obligate hyperparasite Pasteuria penetrans. Rev. Nematol. 14, 611-618.

Giannakou, I.O., Pembroke, B., Gowen, S.R., Davies, K.G., 1997. Effects of long term storage and above normal temperatures on spore adhesion of Pasteuria penetrans and infection of the root-knot nematode Meloidogyne javanica. Nematologica 43, 185-192.

Hussey, R.S., Barker, K.R., 1973. A comparison of methods of collecting inocula for Meloidogyne spp., including a new technique. Plant Dis Rep 57, 1025-1028.

Jenkins, W.R., 1964. A rapid centrifugal-flotation technique for separating nematodes from soil. Plant Dis. Report. 48, 692.

Kariuki, G.M., Dickson, D.W., 2007. Transfer and development of Pasteuria penetrans. J. Nematol. 39, 55-61.

28 Timper et al.

Koehler, H.H., 1999. Predatory mites (Gamasina, Mesostigmata). Agric. Ecosys. Environ. 74, 395-410.

Lenz, R., Eisenbeis, G., 2000. Short-term effects of different tillage in a sustainable farming system on nematode community structure. Biol. Fert. Soils 31, 237-244.

Madulu, J.D., Trudgill, D.L., Phillips, M.S., 1994. Rotational management of Meloidogyne javanica and effects on Pasteuria penetrans and tomato and tobacco Yields. Nematologica 40, 438-455.

Mankau, R., Prasad, N., 1972. Possibilities and problems in the use of a sporozoan endoparasite for biological control of plant parasitic nematodes. Nematropica 2, 7-8.

Mateille, T., Duponnois, R., Dabire, K., Ndiaye, S., Diop, M.T., 1996. Influence of the soil on the transport of the spores of Pasteuria penetrans, parasite of nematodes of the genus Meloidogyne. Europ. J. Soil Biol. 32, 81-88.

Noel, G.R., Atibalentja, N., Bauer, S.J., 2010. Suppression of Heterodera glycines in a soybean field artificially infested with Pasteuria nishizawae. Nematropica 40, 41-52.

Okada, H., Harada, H., 2007. Effects of tillage and fertilizer on nematode communities in a Japanese soybean field. Appl. Soil Ecol. 35, 582-598.

Oostendorp, M., Dickson, D.W., Mitchell, D.J., 1990. Host range and ecology of isolates of Pasteuria spp. from the southeastern United States. J. Nematol. 22, 525-531.

29 Timper et al.

Oostendorp, M., Dickson, D.W., Mitchell, D.J., 1991. Population development of Pasteuria penetrans on Meloidogyne arenaria. J. Nematol. 23, 58-64.

Ortiz, B.V., Perry, C., Sullivan, D., Lu, P., Kemerait, R., Davis, R.F., Smith, A., Vellidis, G., Nichols, R., 2012. Variable rate application of nematicides on cotton fields: A promising site-specific management strategy. J. Nematol. 44, 31-39.

Overstreet, C., McGawley, E.C., Khalilian, A., Kirkpatrick, T.L., Monfort, W.S., Henderson, W., Mueller, J.D., 2014. Site specific nematode management-development and success in cotton production in the United States. J. Nematol. 46, 309-320.

Robinson, A.F., 2008. Nematode management in cotton, in: Ciancio, A., Mukerji, K.G. (Eds.), Integrated Management and Biocontrol of Vegetable and Grain Crops Nematodes. Springer, pp. 149-182.

Sanchez-Moreno, S., Nicola, N.L., Ferris, H., Zalom, F.G., 2009. Effects of agricultural management on nematode-mite assemblages: Soil food web indices as predictors of mite community composition. Appl. Soil Ecol. 41, 107-117.

Spaull, V.W., 1984. Observations of Bacillus penetrans infecting Meloidogyne in sugarcane fields in South Africia. Rev. Nematol. 7, 277-282.

Stirling, G.R., 1984. Biological control of Meloidogyne javanica with Bacillus penetrans. Phytopathology 74, 55-60.

30 Timper et al.

Stirling, G.R., 2014. Biological Control of Plant-parasitic Nematodes: Soil Ecosystem Management in Sustainable Agriculture, 2nd Ed., CABI, Wallingford

Talavera, M., Mizukubo, T., Ito, K., Aiba, S., 2002. Effect of spore inoculum and agricultural practices on the vertical distribution of the biocontrol plant-growth-promoting bacterium Pasteuria penetrans and growth of Meloidogyne incognita-infected tomato. Biol. Fert. Soils 35, 435-440.

Thomas, J.E., Allen, L.H., McCormack, L.A., Vu, J.C., Dickson, D.W., Ou, L.T., 2004. Diffusion and emissions of 1,3-dichloropropene in Florida sandy soil in microplots affected by soil moisture, organic matter, and plastic film. Pest Manag. Sci. 60, 390-398.

Thomas, S.H., Smith, D.W., 1993. Effects of 1,3-dichloropropene for Meloidogyne incognita management on cotton produced under furrow irrigation J. Nematol. 25, 752-757.

Timper, P., 2009. Population dynamics of Meloidogyne arenaria and Pasteuria penetrans in a long-term crop rotation study. J. Nematol. 41, 291-299.

Timper, P., Davis, R., Jagdale, G., Herbert, J., 2012. Resiliency of a nematode community and suppressive service to tillage and nematicide application. Appl. Soil Ecol. 59, 48-59.

Timper, P., Davis, R.F., Webster, T.M., Brenneman, T.B., Meyer, S.L.F., Zasada, I.A., Cai, G., Rice, C.P., 2011. Response of root-knot nematodes and Palmer amaranth to tillage and rye green manure. Agron. J. 103, 813-821.

31 Timper et al.

Timper, P., Minton, N.A., Johnson, A.W., Brenneman, T.B., Culbreath, A.K., Burton, G.W., Baker, S.H., Gascho, G.J., 2001. Influence of cropping systems on stem rot (Sclerotium rolfsii), Meloidogyne arenaria, and the nematode antagonist Pasteuria penetrans in peanut. Plant Dis. 85, 767-772.

Verdejo-Lucas, S., 1992. Seasonal population fluctuations of Meloidogyne spp. and the Pasteuria penetrans group in kiwi orchards. Plant Dis. 76, 1275-1279.

Wardle, D.A., Yeates, G.W., Watson, R.N., Nicholson, K.S., 1995. The detritus food-web and the diversity of soil fauna as indicators of disturbance regimes in agroecosystems. Plant Soil 170, 35-43.

Webster, T.M., Scully, B.T., Grey, T.L., Culpepper, A.S., 2013. Winter cover crops influence Amaranthus palmeri establishment. Crop Prot. 52, 130-135.

Weibelzahl-Fulton, E., Dickson, D.W., Whitty, E.B., 1996. Suppression of Meloidogyne incognita and M. javanica by Pasteuria penetrans in field soil. J. Nematol. 28, 43-49.

Figure 1

Timper et al.

8

Spores/J2

Gall index

4

6

3 4

2 2 0 2011

1 0 2012

2013 Year

2014

Gall index

Spores/J2

5

Timper et al.

Spores per assay J2

Figure 2

6

a ab

4

b

b b

2 0

Spores per native J2

C-C-C-C F-C-F-C F-F-C-F C-F-F-C F-F-F-F 1.5 a 1

ab

b

b

b

0.5 0 C-C-C-C F-C-F-C F-F-C-F C-F-F-C F-F-F-F Frequency of fumigation

Figure 3

Timper et al.

Y = 4.45 – 0.03* (%J2 w spores) R = -0.57

8

Gall index

6 4 2 0 0

20

40 60 80 J2 with spores (%)

100

32 Timper et al.

Highlights •

Fumigation with 1,3-dichloropropene reduced abundance of Pasteuria penetrans.

•

The reduction was small compared to the large year-to-year variation in abundance of P. penetrans spores.

•

The bacterium appeared to suppress nematode populations in some but not all years.

•

Estimates of spore abundance in the spring were a good predictor of root galling in the fall.