Meloidogyne marylandi is Involved in, but not the Primary Cause of Creeping Bentgrass Decline of Putting Greens in Southern California

Turf samples were collected from four declining creeping bentgrass greens with sandy soil several hundred yards apart at a golf course in Indian Wells, Riverside County, California. From each green, five cores were taken with a soil sampling probe (2 cm diameter, Oakfield Apparatus Inc., Fond du Lac, WI) to approximately 10 cm depth, pooled, and stored in a cooler at approximately 15 °C until processing. Soil was washed from the root-thatch mat layer and roots were examined for the presence of galls. They were incubated in a mist chamber for nematode extraction (modified Seinhorst, 1950). Second-stage juveniles (J2) were counted after 5 days of incubation at 26 °C. Some roots were stained in acid fuchsin (Hooper, 1970) to visualize developing juveniles in the tissues. About 50 J2 were sent to the CDFA for molecular identification.

One RKN egg mass from a creeping bentgrass root was used to inoculate creeping bentgrass cv. Penn A-4 that had been seeded in a 164 ml plastic tube (Ray Leach “Cone-tainers” Nursery, Canby, OR, USA) filled with pasteurized sand. This creeping bentgrass-RKN culture was maintained in a greenhouse at 25 ± 2 °C under ambient light. Eight weeks later, the creeping bentgrass roots were rinsed free of sand with a gentle stream of water and placed in a misting chamber for 2 weeks to obtain J2. This population was further reared in larger pots (800 cm³) filled with pasteurized sand and seeded with creeping bentgrass cv. Penn A-4, bermudagrass (Cynodon dactylon ‘Tifway’), rye (Secale cereale ‘Merced’), and barley (Hordeum vulgare ‘Morex’). Eight weeks later, J2 were extracted from the root systems as described above. Since many J2 were obtained from barley cv. Morex, this plant was used to further multiply and maintain the RKN population as the source of nematodes for morphological examination, molecular identification, and inoculum for experiments.

RKN females, J2, and males were obtained from infested barley cv. Morex roots. Thirteen females were excised from the roots, and perineal patterns were cut and transferred to glycerin on microscope slides for examination. Males (11) and J2 (25) obtained from roots after mist-chamber extraction were heat-killed, processed to glycerin, and mounted on microscope slides for measurement and examination (Seinhorst, 1962).

DNA was extracted from several J2 using proteinase K. DNA extraction and PCR protocols were conducted according to Subbotin (2021). Two primer sets were used for amplification of nematode genes: i) D2A (5′ - ACA AGT ACC GTG AGG GAA AGT TG - 3′) and D3B (5′ - TCG GAA GGA ACC AGC TAC TA - 3′) amplifying the D2-D3 expansion segments of 28S rRNA gene and ii) JB3 (5′ - TTT TTT GGG CAT CCT GAG GTT TAT - 3′) and JB5 (5′ - AGC ACC TAA ACT TAA AAC ATA ATG AAA ATG - 3′) primers for amplifying the partial COI gene. The successfully amplified fragments were purified and then sequenced by Azenta (MA, USA) with the primer pairs used in PCR. A Blastn search was used to compare the sequences obtained with those deposited in the GenBank. The new sequences were submitted to the GenBank database under accession numbers PP949975 (COI gene) and PP951498 (28S rRNA gene).

The host status of 26 plant species (Table 1), comprising several graminaceous and dicotyledonous crop species, was evaluated in 2 separate greenhouse trials for this Meloidogyne species. Plants were seeded in 800 cm³ plastic pots containing pasteurized sandy soil. For each plant species, 3 pots were seeded with an amount of seed depending on the size of the mature plant. Two weeks after the emergence of the seedlings, the pots were inoculated with 3,500 RKN eggs from the previously described barley culture plants. After inoculation, the pots were randomized on a greenhouse bench, watered and fertilized through an automated drip irrigation system, and grown for another 6 weeks at 25 ± 2 °C. The roots were carefully rinsed off, blotted dry, and weighed. Eggs were extracted from the roots by shaking for 3 min in a 0.5% NaOCl solution (Hussey and Barker, 1973) and counted with a Hawksley counting chamber under a dissecting microscope at 40x magnification. The complete experiment was repeated once. The reproduction factor RF was calculated as Pf/Pi (number of eggs per root system at harvest/3,500).

Soil temperature is the main abiotic factor determining the rate of development of RKN (Trudgill, 1995; Giné et al., 2021). Consequently, its effect was the focus of this part of the study. The methodology was similar to the one used by Lahtinen et al. (1988). One-liter plastic pots were filled with 800 cm³ steam-pasteurized river bottom sand (90% sand, 1% silt, 8% clay, 0.2% OM, pH 7.5), seeded with barley cv. Morex (2 seeds/pot) and placed on a greenhouse bench for three weeks to germinate. The pots were then placed in water baths, with the soil level inside the pots lower than the water level outside. The tops of the pots were insulated with polystyrene disks with holes for the barley plants. The pots were then transferred to water baths set at 17, 20, 25, 30, and 35 °C (five pots for each temperature). A temperature reader (HOBO, Spectrum Technologies, Inc., Plainfield, IL, USA) set to record the temperature at one-hour intervals was inserted into one pot in each water bath.

Two days later, 2 ml of a J2 suspension, containing approximately 640 J2 (experiment 1) or 2,000 J2 (experiments 2–4), obtained from infested barley cv. Morex roots, were pipetted into 4 shallow holes close to the stems of the barley plants. Plants were grown and fertilized once with 5 g of controlled-release fertilizer (Osmocote 17-6-10; Scotts-Sierra Horticultural Products Co, Marysville, OH) until 20 days after nematode inoculation. Plants were carefully removed from the pots, and the roots were washed free of soil with a gentle stream of water. They were transferred to new, similar pots with five 3-mm-diam. holes drilled in the bottom of each pot. The pots were carefully three-quarters filled with washed small (ca. 5-mm-diam.) stones, placed inside similar pots without holes in the bottom, and returned to the water baths. To collect newly-hatched J2, 300 ml water was added to the inside pot every day and allowed to percolate through the pot. The J2 in the percolate were collected on two 500-mesh (25-micron-pore-size) sieves and counted under a dissecting microscope at 40× magnification.

This entire experiment was done four times at four-week intervals. After the first experiment, the transfer of the barley plants to the pots with stones was not done on the same day for each temperature, but was based on results obtained from the first experiment – i.e., four days prior to the expected first appearance of J2. The second experiment was an exact repeat of the first experiment, but due to a technical problem with the 17 °C water bath, this treatment was lost in the second experiment. The third experiment included 17 °C and temperatures between 25 °C and 33 °C with 2 °C intervals (17, 25, 27, 29, 31, and 33 °C). The fourth experiment was basically a repeat of the third experiment, but with only the 27, 29, and 31 °C treatments.

To qualify as the “first day of life cycle completion”, J2 had to be recovered from at least two out of five replicates, and the number of recovered J2 on each of the two days following the initial recovery had to be higher than on the previous day. In each experiment, J2 were collected for at least one week following the initial recovery.

To estimate the base temperature and degree-days required for life cycle completion, a regression analysis was done according to: R=1/S×Ts−Tb/S {\rm{R}} = 1/{\rm{S}} \times {\rm{Ts}} - {\rm{Tb}}/{\rm{S}} With R = reproductive rate [1/ (days to life cycle completion)], S = required heat sum (DD = degree-days), Ts = average soil temperature (°C), and Tb = base temperature (°C).

Creeping bentgrass cv. Penn A-4 was grown from seed in 164 cm³ plastic cones (Ray Leach “Cone-tainer” Nursery, Canby, OR) filled with pasteurized plaster sand. The cones were inoculated at seeding with 0, 100, 1,000, 5,000, or 10,000 M. marylandi J2 and placed in a greenhouse at 25 ± 2 °C under ambient light. The cones were fertilized once every four weeks by drenching with a solution of 5g/L Miracle-Grow (The Scotts Company LLC, Marysville, OH, USA). The trial design was a randomized complete block with 6 replications per treatment, and the trial was repeated once. The grass in each cone-tainer was cut back to approximately 1 cm every 2 weeks, and the fresh weights of the clippings were determined. Nematode damage was expected to reduce the fresh weight of the clippings in the inoculated treatments compared to non-inoculated controls. Both trials were terminated 180 days after seeding. At this time, the roots were carefully rinsed and placed in a mist chamber for 14 days at 26 °C to extract J2.

A Shapiro test (at P = 0.05) was applied to the collected data to test if the data conformed to a normal distribution. If data were normally distributed, ANOVA tests were done to determine the significance of the treatment effects, the repeated experiments, and their interactive effects. Post-hoc separation of means was done using Fishers’s Least Significant Difference (LSD) test (at P = 0.05). If the data were not normally distributed, a non-parametric Kruskall-Wallis test was used to determine the significance of treatment effects, and post-hoc separation of means was done using a Bonferroni test (at P = 0.05). R-language (R Core Team, 2013) was used for statistical analysis.

Above-ground symptoms on creeping bentgrass greens were typical for nutritional problems, with chlorosis, stunted growth, and dieback. Mist chamber extraction of soil and root samples from affected creeping bentgrass areas yielded several hundred RKN J2 per 100 cm³ soil. The roots showed galls typical of root-knot nematode infestation and exhibited light-to-dark-brown discoloration. Staining the roots with acid fuchsin revealed numerous advanced juvenile stages and females of root-knot nematodes.

Males were fairly common in the population that was multiplied on barley cv. Morex. The J2 (n = 25) mean body length was 396.7 ± 3.7 (352.8 – 438.1) μm, and the mean body width was 16.1 ± 0.12 (15.1 – 16.9) μm. Male (n = 11) mean body length was 1127.6 ± 34.9 (864 – 1273) μm. Perineal patterns (n = 13) were rounded-to-ovoid with coarse striae (Figure 1) and conformed to the description of Meloidogyne marylandi Jepson and Golden, 1987.

Figure 1:

Example of two perineal patterns of a Meloidogyne marylandi population isolated from creeping bentgrass in Indian Wells, California.

The sequence of the D2-D3 expansion segments of the 28S rRNA gene showed 100% similarity with that of M. marylandi from bermudagrass, USA (KP901066). The COI gene sequences showed the highest similarity (98.8%) with M. marylandi from Japan (ON453615 and ON171452).

Statistical analysis was applied to Log₁₀[(eggs per root system) +1]-transformed data. ANOVA analysis showed that the experiment (Exp1, Exp2), the crop cultivar, and their interaction were all significant at the 95% confidence level. However, examination of the interaction plot (Figure 2) showed that the results were very similar between the two replicated experiments, with a few exceptions: reproduction on creeping bentgrass, tall fescue, and onion was considerably lower in experiment 1 than in experiment 2. Despite these minor differences, data between the two replicated experiments were combined and analyzed as the average of six replicates. Because the non-transformed and the Log₁₀[eggs per root system +1]-transformed data did not fulfill the requirements of a normal distribution (Shapiro test; P ≤ 0.05), the non-parametric Kruskall-Wallis test was used to test the crop-cultivar effect on the nematode reproduction. The Bonferroni test (at P = 0.05) was used to separate the means. The reproductive factor (RF = Pf / Pi) for host plants was calculated by dividing the number of eggs extracted from the root systems per pot by the number of eggs used as inoculum.

Figure 2:

Average (n = 3) egg production [Log₁₀(eggs per root system) + 1] of Meloidogyne marylandi on 28 crop cultivars in two replicated experiments, 6 weeks after inoculation with 3,500 eggs.

The host status of the different crop cultivars was evaluated based on the reproduction factor (RF) of M. marylandi on those crop cultivars (Table 1). An RF ≥ 10 was considered an excellent host, 10 < RF < 1 a good host, RF ≈ 1 a maintenance host, and RF < 1 a poor to non-host (Ferris et al., 1993). The grain crops barley, oats, corn, and wheat were excellent hosts for M. marylandi and were the only crops with RF-values > 10. Most grasses (except Kentucky bluegrass, tall fescue, and colonial bentgrass) were good hosts, allowing the population to increase. All dicotyledonous plants tested were poor or non-hosts (Table 1). Surprisingly, rye, a crop closely related to wheat and barley (Clifton and Keogh, 2016), was not a host for M. marylandi in these tests.

The life cycle was completed at each of the nine tested soil temperatures, and the results were generally very similar between the experiments. One exception was the result obtained at 31 °C; in experiment 3, it took 24 days for life cycle completion, whereas in experiment 4, it took 32 days (see Table 2). At the lowest temperature tested (17 °C), the number of days required for life cycle completion was the longest (55 and 59 days in exp. 1 and exp. 3; Table 2). Life cycle completion was fastest at 29 °C (23 and 24 days in exp. 3 and exp. 4; Table 2). At higher soil temperatures, the number of days required to complete the life cycle increased again, and it took about 6 days more at 35 °C than at 29 °C (Table 2).

In general, the effects of soil temperature on nematode reproduction during the first five days after life-cycle completion corresponded with the temperature effects on the life-cycle duration. At the highest (35 °C) and lowest (17 °C) temperatures tested, nematode reproduction was low, with only about 3% and 10% of the number of nematodes recovered relative to the number inoculated. In the temperature range from 27 to 30 °C (around the optimum temperature for life cycle completion), the reproduction factor RF (nematode recovered/nematodes inoculated) was greater than 1 in each experiment (Table 3).

Plotting the reproductive rate (1/day) against soil temperature showed that the reproductive rate at 31 °C from experiment 4 did not correspond with the rest of the data (Figure 3). This data point was omitted in the regression analysis. Figure 3 also shows that the reproductive rate starts to decrease at 30 °C and higher soil temperatures. A regression line was thus fitted through the data points over the 17 to 29 °C range. The resulting regression function R=0.0020T−0.0168 {\rm{R}} = 0.0020{\rm{T}} - 0.0168 explained over 99% of the observed variation (correlation coefficient R² = 0.998). According to this function, M. marylandi has a required heat sum of 493 degree-days and a base temperature of 8.3 °C, below which reproduction will not occur.

Figure 3:

Effect of soil temperature on the reproductive rate R (days-1) of Meloidogyne marylandi in four experiments. Line represents regression analysis : R = 0.0020 × Ts-0.0168 for Ts = 17 – 29 °C, according to R = 1/S × Ts - Tb/S, with S = required heat sum (degree-days), Ts = soil temperature (°C), and Tb = base temperature (°C) for reproduction. Correlation coefficient R² = 0.998.

The effects of the two trials – the different inoculation levels and their interactive effect on the total cumulative weight of the grass clippings, the root weight at harvest, and J2 levels at harvest – were statistically analyzed. Data on the total fresh weight of the clippings, root weight, and Log₁₀[x+1]-transformed J2 data satisfied normal distribution requirements (Shapiro test; P > 0.05).

ANOVA analysis showed that inoculum density did not significantly affect the total weight of the grass clippings (P > 0.05). The total weight of the grass clippings was significantly higher in the first than in the second trial (8.54 and 7.71 gram, respectively). The interaction between the experiment and inoculum density was not significant (P > 0.05). Therefore, the weights of the grass clippings under the different inoculum levels are shown as the average of the two replicated trials (n = 12; Figure 4). The average root weight at harvest was not significantly affected by the experiment, the inoculum density, or their interaction (P > 0.05). No J2 were found in the roots from the non-inoculated treatments. In the inoculated treatments, the average number of J2 per root system at harvest (overall: 2,264) was not significantly different in the two replicated experiments or among the different inoculum levels (P > 0.05). The interactive effect between the experiment and inoculum level was also not significant (P > 0.05).

Figure 4:

Average (n = 12) total fresh weight of creeping bentgrass (Agrostis stolonifera) cuttings over a 180-day period under a range of Meloidogyne marylandi second-stage juvenile inoculum densities. Bentgrass was grown in 164 cm³ plastic cones in a greenhouse and cut every two weeks.