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Dep. of Hortic. and Crop Sci., Ohio State Univ., 202 Kottman Hall, 2021 Coffey Road, Columbus, OH 43210
* Corresponding author (danneberger.1{at}osu.edu)
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ABSTRACT |
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 TOPNOTES ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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Abbreviations: bp, base pair • DOT, day of treatment • G-2, ‘Penn G-2’ • RAPD, random amplified polymorphic DNA • TE, Tris EDTA • USGA, United States Golf Association
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INTRODUCTION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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Turfgrass renovation through chemically killing or stripping the existing turf is the most effective method for establishing a new turf stand. Total renovation, however, necessitates the closure of the putting greens for several months during establishment. The financial loss due to the lack of play is significant to the golf club.
Intraseeding is an alternative approach to establishment of a new turfgrass stand. The concept of intraseeding is to slowly introduce, with minimal disturbance to the existing turf, a new creeping bentgrass cultivar into an established stand of creeping bentgrass. Procedures for intraseeding are similar to those for winter overseeding of common bermudagrass [Cynodon dactylon (L.) Pers.] with a cool-season turfgrass, such as perennial ryegrass (Lolium perenne L.) The difference between winter overseeding and intraseeding are significant. Exploiting adaptive differences between bermudagrass and perennial ryegrass can favor one species more than the other. Intraseeding creeping bentgrass into an existing stand, however, results in competition between individuals of the same species with few adaptive differences.
Factors that influence intraseeding include priority of emergence and gap size. Priority of emergence becomes an important factor in intraspecific competition. Mortality and overall growth of a population is greatly influenced by quicker developing individuals (Smoliak and Johnston, 1975; Weaver and Cavers, 1979). Intraseeded cultivars are at a distinct disadvantage, since the existing cultivars already are established. No published research was found that focused on the intraseeding of putting greens. Other studies, however, investigating the seedling survival of grass species sown into established swards were of limited success due to the asymmetrical competition between seedlings and adult plants (Cook and Ratcliff, 1984; Howe and Snaydon, 1986; Snaydon and Howe, 1986; Aguilera and Lauenroth, 1993). The growth of the smaller seedlings are hindered by the presence of the multitillered adult plants, while the larger plants are relatively unaffected by competition with the juvenile seedlings (Silvertown and Doust, 1993). The greatest limiting factor of seedling survival and growth in existing swards was the inability to compete for soil moisture and nutrients with established plants (Cook and Ratcliff, 1984; Snaydon and Howe, 1986; Aguilera and Lauenroth, 1993).
Gap size is an important factor in seedling survival. Caruso (1970) found that early seedling survival of two sweet clover (Melilotus albus Medik. and M. officinalis Lam.) species in established Canada bluegrass (P. compressa L.) swards was positively correlated with canopy gap size. Large gaps, however, in putting greens may not be conducive to immediate play on a smooth, quality putting green and may result in genetic patchiness within the population.
Determining the success of an intraseeding procedure is extremely difficult because few reliable morphological traits could be used to distinguish a new cultivar from the established, older cultivar on the putting green. The population of creeping bentgrass plants growing in a putting green can be more appropriately assessed with the use of molecular techniques like RAPD to detect polymorphic markers between cultivars (Welsh and McClelland, 1990; Williams et al., 1990; Caetano-Anollés et al., 1991b).
Random amplified polymorphic DNA markers were used to identify cultivars of bermudagrass [Cynodon spp.] (Caetano-Anollés et al., 1995), buffalograss {Buchloe dactyloides (Nutt.) Engelm. [= B. dactyloides (Nutt.) Columbus]} (Wu and Lin, 1994), and zoysiagrass [Zoysia japonica Steud.] (Caetano-Anollés et al., 1991a). Golembiewski et al. (1997) found eight oligonucleotide primers produced characteristic markers that successfully identified 11 of 13 creeping bentgrass cultivars using bulk seed samples. The objective of this study was to determine the competitive success of the cultivar G-2 when intraseeded into an established Penncross creeping bentgrass putting green.
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MATERIALS AND METHODS |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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A Pearson chi-squared test for goodness-of-fit (Ramsey and Schafer, 1997) was used to analyze data, and because replication is not germane to the statistical procedure, the greens were treated and analyzed as separate experiments (Exp. 1 and 2). The greens were divided into five 2.43- by 6.08-m plots and received the following treatments arranged in a randomized complete block design: glyphosate [N-(phosphonomethyl)glycine, in the form of its isopropylamine salt] applications, trinexapac-ethyl [4-(cyclopropyl-
-hydroxymethylene)-3,5-dioxocyclohexanecarboxylic acid methyl ester] applications, coring, scalping, and a control that only received the seed. The glyphosate treatment served as the standard for comparison and is referred to as the check plot. This allowed for the success of seedling establishment to be monitored. The plant growth regulator trinexapac-ethyl was used to slow growth of the existing turf for the purpose of reducing its competitive ability.
Prior to treatment initiation on 22 Sept. 1998, each plot was sampled to provide a genetic identification of the Penncross. Thirty random, individual plant samples were taken from each plot, placed in individual 1.5-mL microfuge tubes and stored at -20°C for subsequent primer screening. Intraseeding treatments were performed on 1 Oct. 1998, 15 Apr. 1999, and 9 Sept. 1999.
Nitrogen fertilizer was applied no less than 4 wk prior to each day of treatment (DOT) to reduce the growth of the existing stand. Glyphosate was applied at 3.43 kg acid equivalent ha-1 with a hand-held CO2 sprayer (2.76 kPa) 7 d prior to DOT. A second application of glyphosate was applied for the September 1999 treatment date due to incomplete kill following the first application. Trinexapac-ethyl was applied at 15.6 g a.i. ha-1 24 hr prior to DOT to allow for maximum plant absorption and translocation within the plant (Fagerness and Penner, 1998).
On DOT, the greens were double cut in two directions to a height of 3.1 mm with a Greensmaster 1000 walking greensmower (The Toro Company, Bloomington, MN). The scalp treatments were mowed to a height of 1.5 mm 
20 to 30 times, such that little to no verdure remained.
The coring treatments were imposed with a Ryan GA-30 aerifier (Cushman, Lincoln, NE) using 1.3-cm diam hollow tines at a 7.6-cm depth. Tine spacing was 5.0 cm (within rows) by 6.4 cm (between rows). After core removal, the plots were rolled with a hollow, steel roller to smooth the surface.
All plots, excluding the control, were vertically mowed in two directions using a Bluebird Lawn Comber Model 720 (Bluebird International, Inc., Englewood, CO) with 3.2-mm wide blades on 1.9-cm spacings set to cut grooves 6.4 mm deep in the greens' surface. Loose debris was removed by mowing to a height of 4.0 mm. Excluding the control, all plots were topdressed with 1.6 mm of sand using a Mete-R-Matic F15B Top Dresser (Turfco MFG., Inc., Minneapolis, MN). The sand was worked into the canopy by hand brooming.
The G-2 creeping bentgrass seed (Lesco, Rocky River, OH) was applied in two directions at 12.2 kg ha-1 per direction using a Scotts drop spreader model PF-2 (The Scotts Company, Marysville, OH). Light brooming was performed in alternating directions after each seeding to work the seed into the coring holes and vertical mowing grooves to maximize seed-soil contact.
Post-treatment management of the plots focused on seedling establishment. Irrigation was applied at 2.67 mm six times per day for the first 4 to 6 wk after seeding with an automated system. Thereafter, irrigation was applied frequently to encourage seed germination and to prevent wilt.
After each intraseeding treatment date, the plots were fertilized with a starter fertilizer (Scotts Contec 19 N–25 P2O5–5 K2O Starter Controlled Release Fertilizer, The Scotts Company) at a rate of 25.4 kg N ha-1 within the first wk of seeding. Three applications at this rate were made in 2- to 3-wk intervals. Thereafter, fertilizer (18 N–4 P2O5–18 K2O) was applied at a rate of 24.4 kg N ha-1 each month of the growing season from April to November. Fertilizer totals from 1 Oct. 1998 to 31 Dec. 1998 were 73.2 kg N ha-1, 97.6 kg P ha-1, and 19.5 kg K ha-1. Totals for 1999 were 278.1 kg N ha-1, 253.7 kg P ha-1, and 146.4 kg K ha-1; while 122.0 kg N ha-1, 53.6 kg P ha-1, and 103.9 kg K ha-1 were applied from 1 Apr. 2000 to 23 May 2000 (project termination date).
After seeding, the greens were mowed three times per week. Mowing height was raised to 4.0 mm for 4 wk to minimize seedling damage from the mower. The mowing height then was gradually lowered to 3.1 mm by 0.4 mm wk-1. Mowing of the glyphosate-treated plots did not commence until the seedlings were 6.4 mm tall. After initial mowing, these mowing heights also were lowered to 3.1 mm by 0.4-mm increments wk-1.
All plots were topdressed after each treatment date with 0.79 mm of sand every 2 to 3 wk during the growing season. In addition, an application of mefenoxam {(R)-2-[(2,6-dimethylphenyl)-methoxy-acetylamino]-propionic acid methyl ester} was applied at 0.1 kg a.i. ha-1 using a hand-held CO2 sprayer during the first 3 wk after seedling emergence to prevent seedling loss from damping-off (Pythium spp.). Following subsequent sampling, fungicides and insecticides were applied on a curative basis. An application of iprodione [3-(3,5-dichlorophenyl)-N-(1-methylethyl)-2,4-dioxo-1-imidazolidinecarboxamide] and chlorothalonil (tetrachloroisophthalonitrile) were applied in December 1999 at 1.0 and 1.5 kg a.i. ha-1, respectively, for the control of Microdochium patch [Microdochium nivale (Fries) Samuels et Hallett]. In December 1998 and 1999, a permeable polyester turf cover was placed over each green to protect young seedlings from winter desiccation and direct low temperature injury (Roberts, 1986; Dionne et al., 1999). The covers were removed in both years during the first week of March.
Field Sampling
Plant samples were taken prior to each treatment date on 28 May 1999 and 24 Mar. 2000. Thirty-two randomly selected plants plot-1 were collected on 28 May 1999 as previously described for 22 Sept. 1998. As a check to the validity of the sampling procedure, line intersect sampling (Battles et al., 1996; Ringvall and Ståhl, 1999) was done on 24 Mar. 2000. This sampling procedure entailed the random selection of eight plants from each of four 1.6-dm2 boxes placed in a representative fashion along a randomly selected 6.0-m transecting line. Areas where sod had been torn by the vertical mower were avoided when placing the boxes along the transecting lines. Such areas, which occurred mostly on both scalp and cored treatment plots, were avoided when sampling since seeding into bare patches of soil is not considered intraseeding. On 12 May 2000, 32 samples were taken from the scalp (Exp. 1) plot in the same fashion as on 22 Sept. 1998 and 28 May 1999, while avoiding the unrepresentative areas. These samples were used to identify any significant effects caused by sampling technique. All samples were stored at -20°C for later RAPD analysis.
Laboratory Analysis
Field samples were used to screen primers for RAPD markers that would distinguish G-2 from Penncross. Bulk samples of the two cultivars were used to make primer screening easier and less time consuming (Sweeney and Danneberger, 1994). Individual samples tested for marker reproducibility and were used to identify population changes after each treatment date.
Two individual leaf blades from each pretreatment plot (20 leaves total from the 22 Sept. 1998 sampling) were cut to 1.0 to 1.5 cm (combined length of all aboveground plant structures) and pooled. The G-2 seed was sown in individual 5-cm3 pots with Metro Mix 360 Growing Medium (The Scotts Company) and grown in the greenhouse at 23 ± 5°C until leaves were large enough for DNA extraction. A 1.0- to 1.5-cm leaf segment was cut from each of 20 different G-2 plants and pooled. Fifty decamers [10 base pair (bp) oligonucleotide] supplied by either Ransom Hill Bioscience, Inc. (Ramona, CA) or J.E. Carlson (Univ. of British Columbia, Vancouver, Canada) were screened. Primers producing polymorphisms were reamplified to ensure that the RAPD markers were reproducible. Primer 703 (CCAACCACCC) was selected because it consistently produced a 1050 bp RAPD marker in G-2 and was absent in Penncross. The primer also was tested on individual samples of each cultivar to determine the band frequency in each cultivar. The remaining 24 pretreatment samples per plot were cut down to equivalent sizes (1.0–1.5 cm combined length of all aboveground plant structures) and amplified separately. Band frequency was determined for each of the 10 pretreatment plots. In addition, one 1.0- to 1.5-cm leaf blade segment was cut from each of 41 G-2 plants also grown in the greenhouse in separate pots. The G-2 seed used for greenhouse samples came from the same seed lot as that used in the field. The G-2 plant samples were amplified separately to determine the band frequency of the 1050 bp fragment. This was repeated with a second, separate set of 41 G-2 samples for replication. The 32 samples from each plot collected from the 28 May 1999 and 24 Mar. 2000 sampling were analyzed individually using Primer 703.
The DNA was extracted from leaf tissue by means of the procedure used by Sweeney and Danneberger (1994), except that the pellet was dried for 20 min under vacuum desiccation, instead of 15 min, and was then suspended in 100 µL Tris EDTA (TE) [10 mM Tris-HCL (pH 7.5), 1.0 mM EDTA] and stored at -20°C. An extraction control consisting of the extraction buffer without DNA was included.
Estimates of DNA concentration of sample extracts and extraction controls were made with either a TKO 100 Mini Fluorometer (Hoefer Scientific Instruments, San Francisco, CA) or a TD-360 Fluorometer (Turner Designs, Sunnyvale, CA). Lambda DNA (GIBCO BRL, Grand Island, NY) was used as a known standard for instrument calibration. Extraction controls did not contain detectable DNA, while sample extracts were diluted to 2 to 5 µg DNA mL-1 with TE for amplification.
The amplification protocol used by Sweeney and Danneberger (1994) and Golembiewski et al. (1997) was followed. The 25-µL amplification reaction mixture for this study contained 20 mM Tris-HCL (pH 8.4); 50 mM KCL; 3.0 mM MgCl2; 0.2 µM primer 703; 0.2 mM each dATP, dGTP, dCTP, dTTP; 1.5 units Taq DNA polymerase (GIBCO BRL, Grand Island, NY); and 2 to 5 ng DNA template. Before the 1-µL DNA template was added to bring the final volume of each 0.5-mL sterile microfuge tube to 25 µL, the reagents were combined, vortexed for 1 min, and then divided into individual aliquots for each reaction. Control samples containing all reagents of the amplification reaction mixture, but without DNA template, were included with each set of amplified samples to ensure that observed bands were amplified genomic DNA from the desired leaf tissue, and not primer artifacts (Williams et al., 1990) or contamination. Each reaction sample was overlaid with a drop of DNAse-free mineral oil (Sigma Chemical Co., St. Louis, MO) to prevent volatilization. After a 3-min soak at 94°C to denature the DNA, amplification was performed in a Thermal Cycler (Perkin-Elmer Corp., Norwalk, CT) programmed for 40 cycles of 1 min at 94°C, 1 min at 40°C, and 1 min at 72°C. A 15-min extension period at 72°C followed the last cycle.
Amplification products were resolved using gels containing 1.5% agarose, 0.4 mg L-1 ethidium bromide, and TBE (89 mM Tris-borate, 89 mM boric acid, and 2 mM EDTA). The agarose was melted and poured into a 15- by 15- by 0.6-cm gel tray with two 20-well combs. Samples were prepared for electrophoresis by adding 4.0 µL sample buffer III (2.4 g L-1 bromophenol blue, 2.5 g L-1 xylene cyanol, and 3 g L-1 glycerol) (Sambrook et al., 1989) to each amplification mixture. Samples were vortexed to mix the loading dye, while brief centrifuging at 15 600 x g collected the sample under the mineral oil. After the gel had solidified (0.5 h), each well was loaded with 14 µL of the amplification mixtures from under the mineral oil. Gels containing the 28 May 1999 and 24 Mar. 2000 post-treatment field samples also were loaded with samples from the greenhouse G-2 tissue that knowingly produced the 1050-bp band for easier scoring of amplification products (Fig. 1)
. Electrophoresis was conducted at 4 V cm-1 for 2 h in a Horizontal Gel Electrophoresis System Model H4 (Bethesda Research Laboratories, Gaithersburg, MD). Amplification products were viewed under ultraviolet light and photographed using FOTO UV 310 DNA Transilluminator and a FOTO/Analyst CCD Video Camera Module (Fotodyne, Inc., Hartland, WI), respectively.
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Changes in plot populations were identified by comparing the number of hits and misses from each pretreatment plot with those from the 28 May 1999 and 24 Mar. 2000 sampling dates. The 24 Mar. 2000 samples were compared with those of 28 May 1999 to determine any significant changes from one treatment date to the next. In addition, samples from all plots showing significant changes were compared with the greenhouse G-2 samples to determine if those plots had completely shifted to the intraseeded cultivar. Data from the pretreatment sampling date and the greenhouse G-2 were modified to 32 samples so that all comparisons were based on the same sample size. The null hypothesis for each test was that no significant change had occurred.
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RESULTS AND DISCUSSION |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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The comparison of the 28 May 1999 samples with those from the pretreatment sampling showed that changes had occurred in the populations of both glyphosate-treated plots and the scalp (Exp. 2) plot (Table 1). The scalp (Exp. 2) plot was not composed of all G-2 since there was a significant (P = 0.001) difference between it and the greenhouse G-2 (Table 2) . The glyphosate-treated (Exp. 1) plot was not different from the greenhouse G-2, indicating that a complete shift to the intraseeded cultivar had occurred. However, the glyphosate-treated (Exp. 2) plot was significantly (P = 0.001) different from the greenhouse G-2. This was probably the result of surviving Penncross plants from incomplete glyphosate kill.
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The only changes from the 28 May 1999 to the 24 Mar. 2000 sampling date were found in the scalp (Exp. 2) and glyphosate-treated (Exp. 2) plots (Table 1), which further support previous results. It was concluded that as of the 24 Mar. 2000 sampling date, the scalp (Exp. 2) plot had shifted back to Penncross since no evidence of G-2 was found. In addition, the glyphosate-treated (Exp. 2) plot had completely shifted to G-2 by the same sampling date. As a result, differences were found between the two sampling dates in these plots.
Results indicated that the mechanical surface disruptive practices used in this study, in particular, scalping, coring, and vertical mowing, and applications of trinexapac-ethyl were quite limited as tools for successful intraseeding. While it seemed these treatments provided greatly reduced competitive pressure from the Penncross, particularly with the almost complete defoliation of plants from scalping, and adequate seed-soil contact from the aggressive vertical mowing, seedling establishment proved to be poor. One possible explanation for this is that root competition from the existing turf was not eliminated. Cook and Ratcliff (1984) reported that root competition for nutrients with established, native tanglehead [Heteropogon contortus (L.) P. Beauv. ex Roem. and Schult.] grassland species was the main factor limiting the growth of green panic [Panicum maximum var. trichoglume Robyns (= P. maximum Jacq.)] seedlings. In a study reported by Snaydon and Howe (1986), shoot competition with established perennial ryegrass had little effect on invading grass seedlings, regardless of ryegrass density or fertilizer applications. They concluded that belowground competition for nutrients (probably for N) was the primary means by which the established ryegrass competed with invading grass seedlings. This would explain why the glyphosate treatments were so successful for G-2 establishment. With complete kill of the existing plants, shoot and root competition was eliminated, providing conditions conducive for seedling growth and establishment.
Initially, it was not uncommon to visually observe seedlings in the vertical mowing grooves in many of the plots, which may explain why the success of intraseeding is often overestimated. As existing plants recover and the canopy closes, the intraseeded cultivar may seem to be present. However, newly germinated seedlings do not necessarily compete with existing plants for resources. Instead, given minimal soil moisture and space, the energy for germination and early growth of seedlings comes from metabolites stored in the endosperm (Langer, 1972). It may not be until the leaves have unfolded and the plant becomes autotrophic that competition with other plants becomes an important factor limiting its growth. Harper (1977) suggested that just the "small size of seedlings and the risks involved in changing from heterotrophic nutrition based on seed reserves to autotrophic life make this stage especially hazardous." Results of this study have shown that once interactions between the G-2 seedlings and neighbor Penncross plants occur, the juvenile plants compete poorly with the established Penncross plants.
Complete shifts in the population are not expected or likely to occur all at once with intraseeding. Rather, it is more probable that a change to the new cultivar will occur during a period of repeated treatments as more and more seedlings become established each time. However, the superficial amounts of G-2 found in the treatment plots (excluding glyphosate plots) at the termination of the project using our techniques and timings led us to conclude that additional treatments will probably yield little additional success. Therefore, until more effective intraseeding methods are developed, greens dominated by existing cultivars that respond poorly to improved management regimes or have become extremely difficult to maintain may benefit more from complete renovation. While a higher cost may be incurred, complete renovation, unlike intraseeding, offers the added advantage of knowing that the new cultivar will dominate the population.
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ACKNOWLEDGMENTS |
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NOTES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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Received for publication June 22, 2001.
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REFERENCES |
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TOPNOTESABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTS AND DISCUSSIONREFERENCES |
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