Depth distribution of plant-parasitic nematodes on bentgrass golf greens in Missouri and Indiana

Sample collection: Twenty golf putting greens established with creeping bentgrass or a mix of creeping bentgrass and Poa annua were sampled in April, June, August and October of 2021 and 2022. Based on their broad geographic distribution and previously observed high lance nematode populations, seven putting green sites in Missouri and three in the Kansas City area of Kansas [KC: Mission Hills (2) and Olathe (1)] were sampled in 2021. In 2022, ten additional sites in Indiana without historic nematode population data were sampled based on their broad geographic distribution across the state (Table 1). The length of each of the putting greens was measured and four equal transects were created across the green in a “W” shaped pattern. Each transect was randomly divided by four to create three points of even distribution across the green, for a total of 12 sampling points per green. Soil samples were extracted using a 1.9 cm diameter soil probe to a depth of 25 cm. Samples were separated into five groups by depth: 0–5, 5.1–10, 10.1–15 15.1–20 and 20.1–25 cm. Each group of twelve subsamples was aggregated into a single sample (100 cm³) and stored in a small plastic bag at a cool temperature (<20°C) prior to analysis. Samples were shipped immediately or driven to the University of Missouri SCN Diagnostics Clinic for processing.

Sample processing: All samples were homogenized before extraction. Samples from the 0–5 cm group were blended using a food processor to break up verdure, thatch, and organic material, and samples from the remaining depth groups were mixed thoroughly, breaking any soil clumps with the aid of a rolling pin. For each sample, 100 cm³ of soil was separated by water displacement in a 1000 ml plastic beaker filled with 400 ml of tap water and mixed before being processed using a semiautomatic elutriator (3), followed by isolation using the centrifuge flotation method. Individual PPNs were identified to the genus in a 1 ml aliquot and counted with a hemocytometer under an inverted microscope at 40x to 400x magnification. Individual PPNs were identified to genus based on general morphological characteristics such as cephalic framework, stylet, esophagus type, sexual structure and position, tail shape, and cuticle, among others, with the aid of a pictorial key to genera (20).

Statistical analysis: Populations at the five sampling depths were compared to assess the vertical population distribution and change in population density among months. Differences in sampling depth-group populations during different sampling months were evaluated using PROC GLIMMIX in SAS 9.4 (SAS Institute). MO/KC nematode populations were analyzed together and grouped separately from Indiana. Least squared means of PPN population per 100 cm³ of soil were subjected to analysis of variance in PROC GLIMMIX (SAS 9.4; SAS Institute), and means were separated using Fisher's protected LSD (P < 0.05). Variables included were sampling depth, sampling month and sampling depth by month, with site set as a random variable.

Phylogenetic analysis: A subset of lance and root-knot nematodes were selected for molecular analysis from one site in STL and ten sites in Indiana, respectively (Table 2). Thirty-two individual lance and 13 root-knot nematodes (J2) were hand-picked and placed into centrifuge tubes with 200 μL of nuclease-free water. DNA was extracted from individual lance and root-knot nematodes using the EXTRACT-N-AMP kit (Sigma, St. Louis, MO), with the manufacturer's protocol modified as seen in Ma et al. (19).

The rDNA ITS region from morphologically identified lance nematodes was amplified with genus-level primer sequences Hoc-1f (5′- AACCTGCTGCTGGATCATTA-3′) and LSUD-03r (5′- TATGCTTAAGTTCAGCGGGT-3′) and were subsequently sequenced (1). The D2/D3 region of the 28S gene from morphologically identified root-knot nematodes was amplified with primer sequences ((RK28SF (5′- CGGATAGAGTCGGCGTATC-3′) and MR (5′- AACCGCTTCGGACTTCCACCAG-3′)) designed by Ye et al. (36). Amplification was conducted in a 25 μl mixture containing 10 μl of Taq Ready Mix (Sigma), 13 μl of nuclease-free water, 1 μl of DNA and 0.5 μl of each primer. PCR amplification of the ITS rDNA region of lance nematode samples was conducted with the following cycle design: initial denaturation at 95°C for 3 min, followed by 35 cycles of 95°C for 45 s, 59°C for 1 min 30 s, 72°C for 2 min and a final extension at 72°C for 10 min. PCR amplification of the D2/D3 region of root-knot nematode samples was conducted with the following cycle design: initial denaturation at 95°C for 5 min, followed by 40 cycles of denaturation at 94° for 30 s, 55°C for 45 s, 72°C for 1 min and a final extension at 72° for 10 min. Amplification was confirmed via electrophoresis on a 1% agarose gel. Amplicons were purified with ExoSAP-IT (ThermoFisher, Waltham, MA) following the manufacturer's recommendations and sent to Eurofins Genomics (Eurofins Genomics, Louisville, KY) for sequencing. Lance and root-knot sample DNA was also amplified using species-specific primers (Table 3) developed for H. stephanus, H. columbus and H. galeatus (19,1) and M. naasi, M. marylandi, M. graminis and M. incognita (36), respectively. Species-specific amplicons were visualized after electrophoresis on a 1% agarose gel for further evidence of speciation.

Phylogenetic trees were developed with consensus sequences using ClustalW in MEGA (Pennsylvania State University, State College, PA). The ITS sequences of Hoplolaimus spp. described in Holguin (16) were downloaded from GenBank and included in the phylogenetic analysis of lance nematodes. The 16S (21) and 28S (21, Tian, 2017, Ye, 2016) sequences of Meloidogyne spp. were downloaded from GenBank and included in phylogenetic analysis of root-knot nematodes. Both phylogenetic trees were constructed with the neighbor-joining algorithm using the Kimura two-parameter model with Litylenchus spp. (LC383724) as the outgroup. Bootstrap values on branch nodes are based on 1,000 random samples of the data set. Morphological characterization: One female lance nematode from site 5 (Table 2) was used for morphological analysis at the Purdue Electron Microscopy Facility (West Lafayette, IN). Micrographs were acquired using a FEI Nova NanoSEM (Thermo Fischer, Waltham, MA) scanning electron microscope (SEM). The lance nematode was processed with a 2.5% glutaraldehyde fixation for 15 minutes, 1% osmium tetroxide for 15 minutes, a five-minute water rinse, then a series of five-minute ethanol dehydrations (50%, 75%, 95%, 100%) followed by dehydration via hexamethyldisilazane (HMDS) and a final platinum coating.

Overall PPN populations: PPNs were more abundant in Indiana in 2022 than in MO/KC in 2021 (59,229 and 47,242, respectively) (Fig. 1). The lack of nematicide application in Indiana compared to MO/KC may be the cause of this disparity in populations (Table 1). Higher lance nematode populations were found in MO/KS than in Indiana (average of 1,225 and 462, respectively), potentially due to targeting sites with historically high lance nematode populations. Root-knot nematodes were more abundant in Indiana than in MO/KC (average of 1,997 and 857, respectively). Ring nematodes were more abundant in Indiana than in MO/KC (average of 2,617 and 1,432, respectively).

Figure 1:

Lance nematode population dynamics: A significant depth-by-month interaction was observed in MO/KS lance populations in 2021 (P = 0.0487, Tables 4,5). Populations sampled from the upper 0–5 cm of the soil in October of 2021 in Missouri were significantly higher than any other depth and month (Tables 4,5). Lance populations sampled across Indiana in 2022 showed no significant depth-by-month interaction. Lance nematode populations were higher in August 2022 in Indiana than the other three months sampled (P = 0.003, Tables 4,6). As opposed to MO/KS, Indiana lance populations were statistically higher at the 5.1–10 cm sampling depth (P < 0.0001, Tables 4,6)

Hoplolaimus speciation and phylogeny: In total, 31 ITS sequences were obtained using the DNA isolation and PCR amplification methods described above. Of the 31 sequences, 26 grouped in a clade with Genbank accessions of H. stephanus, and five grouped with H. magnistylus (Fig. 2). Amplicons from the HS-1f/HS-1r primer set designed by Ma et al. (19) amplified H. stephanus and no other species (Fig. 3). In reactions containing species-specific primer sets designed for H. columbus and H. galeatus, no amplicons were visualized. H. magnistylus was identified following successful amplification utilizing the genus-level Hoc-1f/LSUD-03r pairing and sequencing, but species-specific primers targeting H. magnistylus were not used for confirmation. DNA extracts of ITS sequences in the H. magnistylus clade did not have visualized amplicons after PCR with species-specific primers designed to amplify H. galeatus, H. stephanus or H. columbus.

Figure 2:

Figure 3:

Morphological traits characteristic of H. stephanus were observed with SEM. Specifically, the presence of four lip annules (Fig. 4A), the presence of an epiptygma (Fig. 4B), 25 longitudinal striae on the basal lip annule that were within the range of that of H. stephanus and not H. galeatus (Fig. 4C) and four lateral incisures (Fig. 4D) (16,32).

Figure 4:

Root-knot nematode population dynamics: Populations of root-knot nematodes in MO/KC were not significantly different between sampling months (Tables 4,5). Populations were higher at the 0–5 cm sampling depth in 2021 than all other depths, and the 5.1–10 cm depth had higher populations than deeper samples (P = 0.0004, Table 5). In contrast, root-knot nematode population levels in Indiana were significantly higher in April 2022 than any other month in 2022 (P < 0.0001, Tables 4,6). Root-knot nematode populations in Indiana were higher at the 0–5 cm and 5.1–10 cm sampling depths than at the lowest three depths (P < 0.0001, Tables 4,6).

Meloidogyne speciation and phylogeny: In total, 13 28S D2/D3 sequences were obtained using the primer pair RK28SF/MR as described by Ye et al. (36). Of the 13 D2/D3 sequences obtained, six grouped in a clade with Genbank accessions of M. naasi, four with M. graminicola and three with M. marylandi (Fig. 5). Additional species confirmation with species-specific primers was provided for M. naasi and M. marylandi. (Fig. 6). M. graminicola was not tested with species-specific primer sets for gel confirmation in this study.

Figure 5:

Figure 6:

Other nematode population trends: Free-living, or non-plant parasitic nematodes, were the most abundant nematode in these samples at 10 times or more the number of PPNs found. The most abundant PPN in MO/KC and Indiana was the ring nematode (Criconemoides spp.) (Fig. 1). In both MO/KC and Indiana, free-living and ring nematodes had a statistically significant depth-by-month interaction (P = 0.0254, P = 0.0001; P < 0.0001, P < 0.0001, respectively, Tables 4–6). In MO/KC, ring and free-living nematodes were significantly more abundant in the 0–5 cm range during October than any other depth–month combination (Tables 4,5). In Indiana, higher ring nematode populations were observed during August at a depth of 0–5 cm than any other depth–month combination (Tables 4,6). In Indiana, free-living nematodes were most abundant during the months of June and August at the 0–5 cm sampling depth, with no significant difference between the two (Tables 4,6).

Populations of lance, root-knot, ring and free-living nematodes fluctuated throughout the season across all sampling sites and sampling depths. Other researchers have also demonstrated considerable variation in plant-parasitic nematode (PPN) populations seasonal dynamics (2,10,18,22,25,26). Some trends agree with one another despite differences in their cropping systems, and some are vastly different. In turfgrass, lance nematode population dynamics have been tied to soil temperature and natural reproductive cycles, while some studies demonstrated no association (18,26,33). The relationship between the decline of root-biomass and lance populations indicates a self-regulating feedback loop of PPN population density. As the competition for food increases through changes in nutrient density and root-decline, a constraint on the reproductive factor of the PPNs is more prevalent (10). Evidence also indicates a potential shift to endoparasitism when soil temperatures reach a certain threshold (26), which was not accounted for in this study. Lance nematode populations steadily rose throughout the year, and a reduction in lance nematodes in midsummer following root decline was not observed in this study. Lance nematode populations in Indiana in 2022 peaked significantly in August, when soil temperatures most likely are not conducive for creeping bentgrass root development and subsequent increases in lance nematode populations.

On average, root-knot nematode populations were below the threshold in which turfgrass managers are recommended to apply a nematicide (37). Significantly more root-knot populations were found in MO/KC at the 0–5 cm depth, and it is widely reported that these species tend to aggregate at this depth. The presence of significantly more root-knot nematodes in April agrees with an early-year population density described by Barker (2) but contrasts with populations recorded by Sasser (25). In his study, Barker indicates moisture levels were high enough in January to induce egg hatch in root-knot populations in February, so populations recorded then would be higher, followed by a reduction as competition for food increases. Sasser observed population peaks in November, but indicated this population fluctuation may be specific to the host crop. Additionally, Sasser sampled only during the months of February, May and November of one year, and March of the following year. The time gaps between these selected sampling months could be a time in which Meloidogyne spp. populations peaked.

Populations of ring nematodes shared a depth-by-month interaction, but the month in which the most ring nematodes were found in the 0–5 cm range differed between MO/KC and Indiana. While ring populations peaked in August and October in Indiana and MO/KC, respectively, Davis (6) reported ring nematode populations on a mixed bentgrass/bluegrass green in Chicago peaked in June. Davis was able to analyze populations of ring nematodes at the same sampling site through two years and found their populations did not peak in the same month two years in a row. However, Davis noted that trends did not agree year over year, and populations analyzed during the same year on two different greens within the same golf course changed together. Wick (33) observed populations on the same green did not change synchronously, and thus discredits evidence that population dynamics may be extrapolated across greens on the same course.

The disagreement of this study with other similar research indicates attempting to predict PPN populations based solely upon time of the year may be ineffective. In addition, Settle (26) describes the spatial aggregation of PPN communities across a putting green may vastly change the populations observed. Because of this, efforts to predict overall PPN populations within a green may be ineffective with current sampling methods, as one area of the putting green may contain significantly more PPNs than an area just a few meters away. The results of this study indicate a spring nematicide application would lower the PPN's ability to increase their populations throughout the summer, dependent on the nematicides ability to remain undegraded and maintain residual efficacy. However, differences in the half-life of common nematicides, e.g., the relatively short half-life of abamectin compared to longer half-life of fluopyram, is an important consideration when deciding which chemistry to employ (23,35). The presence of significantly more lance and ring nematodes in MO/KC found at 0–5 cm during October in this study indicates an additional early fall nematicide application date may be necessary to suppress these population peaks and potentially reduce these populations the following year. If turfgrass damage in Indiana can be accurately correlated to high lance nematode populations, a late season nematicide application may be more effective in controlling lance populations the following year. However, to target and control other PPNs throughout the summer season, an early spring application may be more effective.

Hoplolaimus spp. and Meloidogyne spp. exhibited differences between species across and within sampling sites. Meloidogyne spp. analyzed in this study were more diverse than Hoplolaimus spp. M. naasi, M. marylandi and M. graminicola are known parasites of creeping bentgrass that were found in this study (21,36). The geographic distribution and host species of several Meloidogyne species seemed to overlap, indicating root-knot species have a mixed distribution and may commonly form a species complex on golf greens. Ye et al. (36) described the most prominent species on a mix of warm and cool season greens in North Carolina were M. marylandi and M. graminis, whereas McClure et al. (21) described the most prominent species obtained from golf course greens in the Western United States as M. naasi. M. naasi is a well-described parasite of turfgrass in the United States, and McClure et al. (21) described that two different Meloidogyne species (M. graminis and M. marylandi) cohabitated a single green in California, suggesting species complexes of Meloidogyne on golf greens are common.

With ITS sequence data as sole evidence of speciation, this is the first report of H. magnistylus associated with bentgrass putting greens to the authors' knowledge. The species was putatively found in a mixed population with H. stephanus at a site in both Missouri and Indiana and alone at another site in Indiana. The species has previously been reported in Arkansas, Illinois and Tennessee parasitizing corn, cotton, soybean and willow (5,16). Morphologically, it differs from H. concaudajuvencus by the possession of rounded tails in second-stage juveniles vs. conically pointed tails with acute termini, having fewer subdivisions in female basal lip annules, and the greater distance from female anterior end to posterior end of esophageal lobes (24). Further micromorphological observation is necessary to confirm this result.

H. stephanus was the most numerous lance nematode characterized on bentgrass putting greens in the Midwest. Ma et al. (19) described an under-representation of H. stephanus in the literature, indicating reports of H. galeatus as the most often reported species responsible for damage on golf putting greens may be inaccurate. The over-representation of H. galeatus may be due to an overlap of host plants and geographic distribution and the difficulty of using morphological traits for speciation. Holguin et al. (16) noted that while their host plant specificity may overlap, geographic distribution may be the driving force for species presence. Specifically, while H. stephanus and H. galeatus were both found to be parasites of Agrostis spp., H. stephanus was the only lance nematode found on bentgrass greens in Ohio and Kansas, and H. galeatus was the only lance nematode found in South Carolina and Florida in their study. In South Carolina and Florida, H. galeatus was found parasitizing warm-season turfgrass species and cool-season Agrostis spp. Recently, H. stephanus was discovered parasitizing creeping bentgrass in Georgia using similar molecular and morphological identification diagnostic techniques to what was used in this study (17). Further characterization of lance nematodes collected from golf courses across the United States is needed to assess whether lance nematode species distributions are determined by geography or by host range.

This study reiterates literature on Hoplolaimus stephanus, regarding it as an under-represented, yet seemingly widely distributed, lance nematode species within the United States (16,19,27,32). The presence of multiple lance and root-knot species, some obtained from the same putting green, indicates a potential overlap in both geographic distribution and host preference. The broad geographic distribution could be due to several factors described by McClure et al. (21) such as topdressing with nematode contaminated sand from other regions, or golf shoes and/or clubs introducing juveniles or eggs when playing golf. Additionally, putting green proximity to agricultural systems and/or native plant species where particular plant-parasitic nematode species may be abundant and exhibit a wide host range could further expose the putting green to the introduction of detrimental, native nematodes over time.